Corrosion Protection for Pipelines Draft Basis of Design ...
Draft Basis of Design Report - dspace2.creighton.edu
Transcript of Draft Basis of Design Report - dspace2.creighton.edu
VOLTAGE GATED SODIUM CHANNEL REGULATION OF
NEURITE OUTGROWTH: ROLE OF NMDA AND
NEUROTROPHIN RECEPTOR SIGNALING PATHWAYS
___________________________________
By
SAIRAM JABBA
___________________________________
A DISSERTATION
Submitted to the faculty of the Graduate School of the Creighton University in Partial
Fulfillment of the Requirements for the degree of Doctor of Philosophy in the Department
of Pharmacology.
_________________________________
Omaha, NE
(7/6/2012)
II
III
ABSTRACT
Activity-dependent N-methyl D-aspartate receptor (NMDAR) signaling, gene
transcription and protein synthesis play major roles in brain functions that regulate
neuronal morphology. Inasmuch as neuronal activity-induced increments in cytoplasmic
sodium may augment NMDAR-mediated currents (Rose and Konnerth, 2001; Yu and
Salter, 1998; George et al., 2009), we reasoned that intracellular Na+ may function as a
signaling molecule and positively regulate neuronal development in immature
cerebrocortical neurons. The central hypothesis of this study is that sodium channel
activators stimulate neuronal development by elevating [Na+]i , augmenting NMDAR
function in presence of activated SFKs, enhancing BDNF release and activating the
downstream TrkB signaling. The specific objective of the proposed work is to elucidate
the signaling mechanisms by which sodium channel activators influence neuronal
morphology in immature cerebrocortical neurons. More specifically, to understand the
relationship between increases in [Na+]i and NMDAR, brain-derived neurotrophic factor
(BDNF)-TrkB mediated neuronal development.
For these studies, sodium channel activators that increase [Na+]i ,antillatoxin
(ATX) and veratridine (VRT) were used as pharmacological tools to determine their
potential to mimic neuronal activity. VGSCs activators robustly stimulated neurite
outgrowth in a hormetic concentration-response relationship and this enhancement was
sensitive to the VGSC antagonist, tetrodotoxin. To unambiguously demonstrate the
enhancement of NMDA receptor function by ATX, we recorded single-channel currents
from cell-attached patches. ATX was found to increase the open probability of NMDA
IV
receptors. Na+ dependent upregulation of NMDAR function has been shown to be
regulated by Src family kinase (SFK) (Yu and Salter, 1998). The Src kinase inhibitor PP2
abrogated ATX-enhanced neurite outgrowth suggesting a SFK involvement in this
response. ATX-enhanced neurite outgrowth was also inhibited by the NMDAR antagonist,
MK-801, and the calmodulin dependent kinase kinase (CaMKK) inhibitor, STO-609,
demonstrating the requirement for NMDAR activation with subsequent downstream
engagement of the Ca2+ dependent CaMKK pathway.
Activity-dependent neuronal development involves N-methyl D-aspartate receptor
(NMDAR) mediated calcium influx and brain-derived neurotrophic factor (BDNF)-TrkB
signaling. We tested the effect of the VGSC activators on BDNF synthesis and release and
TrkB activation in DIV1 cerebrocortical neurons. Inhibition of TrkB receptors and its
downstream effector pathways, PI3K, and PLCγ inhibited VRT-enhanced NOG. VRT
stimulated phosphorylation of TrkB and its downstream effectors Akt, mTOR, PLCγ1,
ERK1/2 and CREB. VRT increased BDNF synthesis and release in a concentration
dependent manner; however, VRT stimulation of TrkB phosphorylation displayed a
biphasic concentration-response curve. VRT stimulation of BDNF synthesis required
VGSCs and NMDARs.
Taken together, these data suggest that VGSC activators seem to be capable of
mimicking activity-dependent neuronal development and hence may represent a novel
pharmacological strategy to regulate neuronal development through NMDA and
neurotrophin receptor-dependent mechanisms.
V
VI
DEDICATION
I dedicate this dissertation to my respected parents, my lovely wife, my family, all my
teachers and all of my friends for their constant love and support, without which I would
not have completed this degree.
VII
ACKNOWLEDGEMENTS
I express my sincere gratitude to Dr. Thomas F. Murray for his invaluable support
and guidance during my Ph.D years. Not only was he a wonderful teacher, but also an
inspirational role model. I would like to thank him for his patience and commitment
towards making me a good researcher and a better person. I would not hesitate to say that
I would like to take him as a role model for training my students in future.
I would like to thank all my committee members, Dr. D. Roselyn, Cerutis, Dr.
Shashank Dravid and Dr. Yaping Tu for their kind assistance and critical evaluation of my
progress, which helped me in learning good science and in becoming a better researcher. I
thank them for their inputs and guidance into my projects. My special thanks to Dr. Dravid
who was not only a wonderful collaborator, but also was a great friend and badminton
partner. He and his lab conducted some critical electrophysiological experiments
discussed in chapter-III.
A special thank you to members of my laboratory who were very helpful and
always made me feel comfortable in the laboratory during all these years. I thoroughly
enjoyed the insightful scientific interactions with Joju George, Zhengyu Cao, Lakshmi
Kelamangalath and Suneet Mehrotra. Special thanks to Bridget Sefranek (nee Leuschen)
for perfectly organizing the research needs and requirements for my projects. I want to
express my gratitude to the faculty, fellow graduate students and staff of Department of
Pharmacology for their help during my years in the program.
I greatly thank my wonderful wife, Sujatha Nagulapally, for her unconditional love and
encouragement. She was my ‘rock of Gibraltar’ during my most stressful period of my
graduate career. I would like to extend my humblest and sincere gratefulness to my
VIII
parents, Lakshmi Rajyam and Panduranga Rao Jabba, for instilling in me the value of
education in life. They were there for me at every step of my life providing unwavering
love and support. I thank my sisters Suma Jabba and Uma Peddireddy for being my role
models and for showering me with unflinching love. My wishes to my lovely niece
Ramyatha Sai, and nephews, Ravi and Hitesh Sai, who always brightened my mood. My
special thanks to my brother-in-laws Ratna Kishore and Gurunath and to my in-laws
Radha and Venkat Nagulapally, for being ever supportive of my ambitions. I would like to
extend my sincere gratitude to Venkateshwaralu Karanam, who was one of my earliest
mentor and true inspiration to take up science. I take this opportunity to thank all of my
friends for providing me with ever constant love, support, encouragement and above all
friendship. I greatly appreciate Chandra Sekhar Baliwada, Bhanu Telugu, Lokaranjan
Somaraju, Pradeep Gendapodi, Kishor Devalaraja, Kalyan Nannuru and Hima Bindu
Ramachandrareddy along with their spouses. My special thanks to Kiran Kotu for being a
very good friend. I truly appreciate the help and company of Pradeep Malreddy, Raja
Rachaktla, Nithya Raveendran, Niranjan Butchi, Rajkumari Sanginaboyina, Vijay and
Rebekah Golla, Satish and Chitra Medicetty, Keil Regehr, Raj Maganti, Casey Devore,
Prasanna Kankanala, Hyma Gajula, Saurabh Jauhari, Vijay Yajjala, Neha Jain, Laxmi
Fogueri, Srinivas Dannaram, Praveen Ramanan, Anantha Gollapudi, Praneet Bathena and
Vamsi Karuturi.
Finally, last but not the least, I offer my humblest gratitude to God for guiding me all
through my life.
IX
Table of Contents
1 CHAPTER 1 - INTRODUCTION .................................................................. 1-14
1.1 Background ................................................................................................... 1-17
1.2 Significance ................................................................................................... 1-22
2 CHAPTER 2 - LITERATURE REVIEW .................................................... 2-23
2.1 Voltage-Gated Sodium Channels .................................................................. 2-23
2.1.1 Introduction: ........................................................................................ 2-23
2.1.2 Primary structure of VGSCs ............................................................... 2-23
2.1.3 VGSCs: Diversity in expression and function .................................... 2-24
2.1.4 Structure of VGSCs at Atomic Resolution ......................................... 2-25
2.1.5 VGSCs function: Molecular perspective ............................................ 2-26
2.2 Voltage-gated sodium channels activators and gating modifiers .................. 2-29
2.2.1 Antillatoxin ......................................................................................... 2-30
2.2.2 Veratridine .......................................................................................... 2-33
2.3 N-methyl-D-aspartate receptors (NMDARs) ................................................ 2-36
2.3.2 Neurotrophins and their receptors, tropomyosin-related kinases
(TRKs): ........................................................................................................ 2-41
2.3.3 Regulation of BDNF-TrkB signaling pathway ................................... 2-47
2.3.4 Role of BDNF-TrkB signaling in neuronal development ................... 2-50
2.4 References ..................................................................................................... 2-56
X
3 CHAPTER 3 - Antillatoxin, a Novel Lipopeptide, Enhances Neurite
Outgrowth in Immature Cerebrocortical Neurons Through Activation of Voltage–
Gated Sodium Channels .............................................................................................. 3-74
3.1 Abstract ......................................................................................................... 3-75
3.2 Experimental Procedures ............................................................................... 3-79
3.3 Results ........................................................................................................... 3-87
3.4 Discussion ..................................................................................................... 3-96
3.5 References ................................................................................................... 3-116
4 CHAPTER 4 – Sodium channel activator-stimulated neuronal development
involves BDNF-TrkB signaling pathway. ................................................................ 4-121
4.1 Abstract ....................................................................................................... 4-122
4.2 Introduction ................................................................................................. 4-123
4.3 Materials and Methods ................................................................................ 4-126
4.4 Results ......................................................................................................... 4-133
4.5 Discussions .................................................................................................. 4-142
List of Figures
Figure 2-1: Subunit structure of VGSCs. ....................................................................... 2-54
Figure 2-2: A) Voltage dependent activation: outward movement of S4 Voltage sensors 2-
54
Figure 2-3: Intracellular NA+ may act as a signalinging molecule and upregulate
NMDARs. ....................................................................................................................... 2-55
XI
Figure 3-1: ATX increases intracellular sodium levels in DIV-1 cerebrocortical neurons. 3-
103
Figure 3-2: Effect of ATX on neurite outgrowth. ........................................................ 3-104
Figure 3-3. Effect of TTX on ATX-induced neurite outgrowth in immature cerebrocortical
neurons. ......................................................................................................................... 3-106
Figure 3-4:Pharmacological evaluation of signaling pathways involved in ATX-enhanced
neurite outgrowth. ......................................................................................................... 3-107
Figure 3-5: ATX-induced neurite extension involves a Src Family kinase. ................. 3-108
Figure 3-6: Quantification of ATX-induced increase of intracellular sodium levels in DIV-
1 cerebrocortical neurons. ............................................................................................. 3-109
Figure 3-7:ATX evoked change in membrane potential in DIV-1 cereobrocortical neurons.
....................................................................................................................................... 3-110
Figure 3-8: ATX-induced Ca2+ influx and pharmacological evaluation in DIV-1
cerebrocortical neurons. ................................................................................................ 3-112
Figure 3-9:. Increase in NMDA receptor channel open probability by ATX. .............. 3-114
Figure 3-10:Schematic diagram of the pathways involved in ATX-induced neurite
outgrowth ...................................................................................................................... 3-115
Figure 4-1: Veratridine stimulated neurite outgrowth and dendritic arborization in
immature cerebrocortical neurons................................................................................. 4-148
Figure 4-2: Veratridine (VRT) increases intracellular Na+ and Ca2+ in DIV1
cerebrocortical neurons and this Ca2+ influx is TTX-sensitive and NMDAR dependent . 4-
149
Figure 4-3:TrkB is essential for veratridine-induced neurite outgrowthVeratridine .... 4-150
XII
Figure 4-4: In situ BDNF ELISA: Veratridine enhances BDNF release in immature
cerebrocortical neurons ................................................................................................. 4-151
Figure 4-5:Veratridine enhances BDNF synthesis in immature cerebrocortical neurons and
this requires VGSCs and partially involves NMDARs................................................. 4-152
Figure 4-6: Veratridine enhances BDNF synthesis in immature cerebrocortical neurons
and this requires VGSCs and partially involves NMDARs .......................................... 4-153
Figure 4-7: - Veratridine-induced neurite outgrowth involves PI3-kinase activity ...... 4-154
Figure 4-8- : Veratridine stimulated Akt phosphorylation involves TrkB receptors .... 4-155
Figure 4-9 - : Veratridine-induced neurite outgrowth involves the PI3K-Akt-mTOR
pathway ......................................................................................................................... 4-156
Figure 4-10: - Veratridine-induced Ca2+ influx involves PLC mediated release of Ca2+
from intracellular .......................................................................................................... 4-157
Figure 4-11: Veratridine-induced neurite outgrowth requires phospholipase C (PLC) 4-158
Figure 4-12:MAPK pathway has a modest role in veratridine-induced neurite outgrowth. 4-
159
Figure 4-13:Pharmacological characterization of Akt activation by veratridine .......... 4-160
Figure 4-14: – Model ................................................................................................... 4-161
List of Tables
Table 1: Location and pharmacological properties of various VGSCs isoforms……..2-55
Table 2: Receptor sites on VGSCs (modified from Catterall et al., Pharmacol Rev. 2005)…………………………………………………………………………………..2-56
XIII
1-14
1 CHAPTER 1 - INTRODUCTION
Activity-dependent N-methyl D-aspartate receptor (NMDAR) signaling, gene
transcription and protein synthesis play major roles in brain functions that regulate
neuronal morphology. Various neurological disorders including several mental
retardation conditions, learning and memory deficit conditions and traumatic brain injury
may be attributed to loss of one or more of these processes. The mechanisms by which
neuronal activity translate into morphological changes are complex and not completely
understood. Understanding these mechanisms will help to determine potential drug
targets to modulate these pathways and develop therapeutic approaches to these
neurological disorders.
Recent studies have demonstrated that neuronal activity-mediated increases in [Na+]i
in neuronal structures augment NMDAR function and may contribute to activity-
dependent synaptic plasticity (Rose and Konnerth, 2001; Yu and Salter, 1998; George et
al., 2009). Inasmuch as neuronal activity-induced increments in cytoplasmic sodium may
augment NMDAR-mediated currents, we reasoned that intracellular Na+ may function as
a signaling molecule and positively regulate neuronal development in immature
cerebrocortical neurons. In this study, sodium channel activators that increase [Na+]i
antillatoxin(ATX) and veratridine (VRT) will be used as pharmacological tools to
determine their potential to mimic neuronal activity. The findings of this study will be
useful in developing novel therapeutic strategies for better management of neurological
diseases involving abnormalities of neuronal morphologies.
1-15
Our long term research goal is to identify novel compounds that can mimic neuronal
activity-dependent neuronal development, and thereby to develop new therapeutic
strategies. The specific objective of the proposed work is to elucidate the signaling
mechanisms by which sodium channel activators influence neuronal morphology in
immature cerebrocortical neurons. More specifically, to understand the relationship
between increases in [Na+]i and NMDAR and brain-derived neurotrophic factor
(BDNF)-TrkB mediated neuronal development. Our proposal is based on published
observations and preliminary data indicating that sodium channel activators promote
neurite outgrowth.
HYPOTHESIS:
The central hypothesis of this proposal is that sodium channel activators stimulate
neuronal development by elevating [Na+]i , augmenting NMDAR function in presence of
activated SFKs, enhancing BDNF release and activating the downstream TrkB-PI3K-
Akt-mTOR pathway.”
SPECIFIC AIM 1:
Evaluate the influence of [Na+]i on NMDAR signaling and investigate the role of src
family kinases (SFKs) in sodium channel activator-enhanced neurite outgrowth. Our
working hypothesis is, sodium channel activators enhance neurite outgrowth by elevating
[Na+]i and augmenting NMDAR function and this Na+ dependent up-regulation of
NMDAR function is regulated by SFKs. To test this hypothesis, up-regulation of
NMDAR function by antillatoxin will be assessed by recording NMDAR single channel
1-16
currents from cell attached patches. The role of NMDARs in ATX and VRT induced
Ca2+ influx will be investigated. Neurite outgrowth assay and Ca2+ influx assay in
presence of PP2, a specific inhibitor of SFKs will be performed to investigate the role of
SFKs. Also the effect of ATX on activation of Src will be assessed.
SPECIFIC AIM 2:
Determine the significance of BDNF-TrkB signaling and its downstream pathways of
TrkB-PI3K-PLCγ-MAPK in regulation of VGSC activator-induced neurite outgrowth,
spinogenesis and synaptogenesis. Our working hypothesis is that sodium channel
activators potentiate NMDAR function and thereby exert trophic effects through
increased gene expression and release of BDNF. The released BDNF regulates neuronal
development by activating its cognate receptor TrkB and the downstream TrkB signaling
pathways, mainly a combination of the PI3K, MAPK and PLCγ pathways. To test this
hypothesis, the involvement of BDNF-TrkB signaling in sodium channel activator-
stimulated neuroal development will be examined by 1) Overexpression of dominant
negative isoform of TrkB, the TrkB.T1 (truncated TrkB) in immature cerebrocortical
cultures, and study its effects on veratridine-mediated neurite outgrowth, spinogensis and
synaptogenesis . 2) Activation of of TrkB receptors and its major downstream pathways,
the PI3K-Akt, PLCγ, MAPK pathways. Enhanced release and increased gene expression
of BDNF under the influence of veratridine will be studied using ELISA and qRT-PCR
respectively.
1-17
1.1 Background
Antillatoxin is a novel VGSC activator
Antillatoxin (ATX) is a structurally novel lipopeptide with an exceptionally high degree
of methylation unlike any known natural product (Lee and Loh, 2006). Isolated from the
cyanobacterium Lyngbya majuscula, this compound is also distinguished by multiple
stereocenters (Orjala et al., 1995a). The essential role of the asymmetric carbon atoms in
ATX is reflected in the stereoselective effects of ATX enantomers (Li et al., 2001).
ATX is considered to be the second most potent ichthyotoxic compound obtained from
marine sources after only brevetoxin (PbTx)-1 (Orjala et al., 1995b). Exposure to L.
majuscula blooms are associated with adverse human health effects, including respiratory
irritation, eye inflammation, and severe contact dermatitis. Previous work has
demonstrated that ATX is a potent activator of voltage-gated sodium channels (VGSCs)
that elevates intracellular Na+ concentration ([Na+]i) in intact neurons (Li et al., 2001;
Cao et al., 2008). Moreover, ATX has been shown to be neurotoxic in cerebellar granule
cells through an indirect activation of N-methyl-d-aspartate receptors (NMDARs) as a
consequence of glutamate release (Li et al., 2001; Li et al., 2004). ATX precise
recognition site on the channel protein remains to be defined. The structure of ATX
includes asymmetric carbon atoms, and the (4R,5R)-isomer is the naturally occurring
compound. The (4R,5R)-isomer appears in profile as an “L” shape with a hydrophobic
interior and a cluster of hydrophilic groups on the exterior of the macrocycle (Li et al.,
2001). Thus, the (4R,5R)-configuration is important for creating a molecular topology
1-18
that is recognized by the acceptor site on the voltage-gated sodium channel α subunit.
Given the unique structure and mechanism of action of ATX, we sought to further
characterize its pharmacologic actions and the functional consequences in cerebrocortical
neurons.
Role of NMDAR in neuronal development and synaptic plasticity.
Neuronal activity has a major role in the development of dendritic complexity and
neuronal circuits. The mechanisms by which neuronal activity translate into
morphological changes are complex. Numerous studies have shown that activity-
dependent neuronal development involves ionotropic glutamate receptors (NMDAR)
mediated calcium influx (Ghosh and Greenberg, 1995; West et al., 2002). These increases
in [Ca2+]i activate signaling cascades that control the transcriptional regulation of
neuronal development. Increase in cytosolic Ca2+ also involves calcium release form
intracellular stores (ER) and is involved in local effects like dendrtic branching and
stabilization of dendritic structures. Intracellular calcium acts as a signaling molecule
largely through binding to calmodulin, a calcium-binding protein that engages
downstream Ca2+/calmodulin-dependent protein kinase (CaMK) and mitogen-activated
protein kinase (MAPK) signaling pathways (Ghosh and Greenberg, 1995; West et al.,
2002). CaMK kinase (CaMKK) has been demonstrated to be an upstream regulator of
both CaMK- and MAPK-signaling pathways. Moreover, previous studies have
demonstrated that activity-dependent neurite outgrowth (Wayman et al., 2006) and
synaptogenesis (Saneyoshi et al., 2008) are regulated by NMDAR-dependent
CaMKK/calmodulin kinase I-signaling cascades. Therefore, NMDARs play a critical role
1-19
in activity-dependent development and plasticity (Ghosh and Greenberg, 1995), dendritic
arborization (Wayman et al., 2006; Wong and Ghosh, 2002; Miller and Kaplan, 2003),
spine morphogenesis (Ultanir et al., 2007), and synapse formation (Saneyoshi et al.,
2008) by stimulating these calcium-dependent signaling pathways. Preliminary results
show the involvement of NMDAR in VGSC activators-stimulated.
Intracellular sodium as a signaling molecule and its involvement in neuronal development.
Regulation of [Na+]i plays a critical role in the nervous system, not only because Na+
influx through VGSCs is responsible for the initiation and propagation of action
potentials but also because various neuronal cell functions, such as intracellular pH, Ca2+
homeostasis, and reuptake of neurotransmitters, are dependent on the Na+ gradient.
Previous studies have further indicated that intracellular Na+ can also act as a signaling
molecule to modulate cell functions, such as cell proliferation, ion channel permeability,
G-protein function, and opioid ligand-receptor interactions (Yu, 2006). Moreover, recent
studies have demonstrated that neuronal activity-mediated increases in [Na+]i in
structures, including soma, dendrites, and spines, may act as a signaling molecule and
contribute to activity-dependent synaptic plasticity (Rose and Konnerth, 2001). In
cerebellar Purkinje neurons, α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid
receptor-mediated Na+ influx was shown to be required for induction of long-term
depression (Linden et al., 1993). In both hippocampal and immature cerebrocortical
neurons, an elevation in intracellular Na+ was found to increase NMDAR-mediated
whole-cell currents and NMDAR single-channel activity by increasing both channel open
probability and mean open time (Yu and Salter, 1998; George et al., 2009). Yu and Salter
1-20
have demonstrated that an increment of [Na+]i of 10 mM was sufficient to produce
significant increases in NMDA receptor single-channel activity. It has also been reported
that increments of [Na+]i greater than 5 mM represent a critical threshold required to
regulate NMDAR-mediated Ca2+ influx. They used veratridine, a VGSC modulator, to
demonstrate that Na+ influx through TTX-sensitive VGSC was sufficient to up-regulate
NMDAR activity. Moreover, this [Na+]i-mediated up-regulation of NMDAR function has
been shown to require Src kinase activation (Yu and Salter, 1998; George et al., 2009).
Src family kinases (SFKs) act as a crucial point of convergence for signaling pathways
that enhance NMDAR activity, and, by up-regulating the function of NMDARs, Src gates
the production of NMDAR-dependent synaptic potentiation and plasticity (Salter and
Kalia, 2004). Hence it is important to explore the regulatory influence of [Na+]i on
NMDAR function in activity-dependent neuronal development.
Regulation of activity dependent dendritic growth and branching by BDNF-TrkB
signaling.
Neurotrophins cooperate with neuronal activity to modulate neuronal morphology.
Studies in the past few years have shown convincing evidence for the involvement of
BDNF-TrkB signaling in activity-dependent dendritic growth, dendric arborization,
synaptogenesis and synaptic transmission (Reichardt, 2006; Huang and Reichardt, 2003;
Yoshii and Constantine-Paton, 2007). BDNF-TrkB mediates increase in dendritic
complexity by increasing the total dendritic/neurite length, dendritic branching
(arborization) and increase in number of primary neurites (Baker et al., 1998; McAllister
et al., 1996). Activation of NMDARs and intracellular Ca2+ rise is critical for the
1-21
expression and release of BDNF (Reichardt, 2006). The released BDNF binds to its
cognate receptors TrkB and acts in an autocrine/paracrine mode to regulate neuronal
development. The binding of BDNF to full length TrkB (FL-TrkB) induces
autophosphorylation of tyrosine residues in the intracellular kinase domains of TrkB,
which in turn, leads to phosphorylation of two tyrosine residues (Tyr 515, Tyr 816)
located outside the kinase activation domain of TrkB (Reichardt, 2006). Phosphorylation
of these two residues recruit and activate three major intracellular signaling pathways that
mediate BDNF-TrkB regulated neuronal morphology. They are: the PI3K-Akt pathway,
the Ras-MAPK pathway and PLCγ-Ca2+ pathway (Reichardt, 2006). Activation of TrkB
at Y816 recruits and activates PLCγ which in turn hydrolyzes PIP2 to DAG and IP3,
where IP3 releases Ca2+ from intracellular stores. The other TrkB phosphorylation site is
Tyr515 that activates downstream MEK-MAPK/Erk signaling, which promotes neuronal
differentiation and growth by influencing transcription events, such as activation of
CREB transcription factor. Tyr 516 indirectly activates PI3K pathway by interacting with
adaptor proteins that regulate PI3K pathway. Another pathway that is indirectly activated
by Phospho-TrkB (Y516) is the PI3K-Akt pathway. Once activated, Akt can
subsequently phosphorylate a number of substrates (GSK3β, MAP2, mTOR) that are
involved in cell survival, NOG/actin dynamics, dendritogenesis and synaptic plasticity
(Read and Gorman, 2009; Yoshii and Constantine-Paton, 2010). Preliminary results for
the present study show the involvement of TrkB in VGSC activators-stimulated NOG in
immature cerebrocortical cultures. This proposed research project will help us understand
1-22
the role and function of BDNF-TrkB signaling in sodium channel-activators mediated
neuronal development.
1.2 Significance
Inasmuch as activity-dependent N-methyl D-aspartate receptor (NMDAR) signaling,
gene transcription and protein synthesis play major roles in brain functions that regulate
neuronal morphology and synaptic plasticity, disruption of one or more of these
processes gives rise to dendritic abnormalities implicated in mental retardation and
autism-spectrum disorders. On the other hand, normal functioning of these mechanisms
are crucial for structural remodeling of dendritic arbor required in synaptogenisis,
synaptic plasticity, and learning and memory. Recently, there is increasing evidence for
mTOR-mediated protein synthesis being important in long term memory formation, and
misregulation of this protein synthesis playing an important role in autism and mental
retardation disorders. Given the increasingly critical role of NMDAR-BDNF-TrkB-PI3K-
mTOR signaling in synapse maturation, plasticity and neurological diseases, expanding
our knowledge of how these signaling pathways function in the brain is key to
understanding learning and memory, as well as developing novel therapeutic approaches
neurological disorders related to NMDAR dysfunction
2-23
2 CHAPTER 2 - LITERATURE REVIEW
2.1 Voltage-Gated Sodium Channels
2.1.1 Introduction:
Voltage gated sodium channels (VGSCs) play an important role in the initiation
(the rising phase) and propagation of action potentials (AP) in neurons and other
electrically excitable cells like myocytes (especially cardiac myocytes) and
neuroendocrine cells. In response to local membrane depolarization they mediate the
rapid influx of the sodium ions into the excitable cell, generating an action potential.
These action potentials will initiate various physiological events like neuronal firing and
muscle contraction. Seminal work in this regard was performed by Hodgkin and Huxley
in 1952 and showed that electrical signals in nerves are initiated by voltage-dependent
activation of sodium current that carries Na+ inward. This sodium current is followed by
inactivation of VGSCs in a few milliseconds and subsequently terminated by activation
of voltage-gated potassium channels.
2.1.2 Primary structure of VGSCs
VGSCs consists of huge complex of four structurally homologous domains of α-
subunits (each 260 kDa), β1 (36 kDa), β2 (33 kDa) subunits (Hartshorne and Catterall,
1981). The ∝-subunit was sufficient enough for functional VGSC currents (Noda et al.,
1986) but the β subunits are necessary for modulation of kinetics and voltage dependence
2-24
of gating (Isom et al., 1995). All four α subunit domains consists of six helical
transmembrane (TM) segments (S1-S6). The voltage sensor for each of the domain is
located in the S4 which consists of positively charged amino acids at every 3 position of
the segment. The loop between S5-S6 re-enters and gets embedded in the TM region and
four such loops of all domains forms the narrow extracellular side of the ion-selective
pore of the channel. The S6 segments of all four domains make the wider intracellular
portion of the pore. Each of the TM segments is connected by short extracellular loops
except for the large one between S5-S6. The domains are connected to each other by
large intracellular loops. The C & N terminal ends are located intracellularly. The β
subunits are unique ion channel subunits with N-terminal extracellular immunoglobulin-
like fold, a single transmembrane segment and a small intracellular segment (Figure 1)
(Isom et al., 1992; Isom et al., 1995). The hydrophobic interactions hold the 2 beta sheets
like a sandwich to form immunoglobin like fold.
2.1.3 VGSCs: Diversity in expression and function
There are at least 10-12 different kinds of VGSCs expressed in various excitable
tissues with subtle amino acid changes among them. Nav1.1, 1.2, 1.3 and 1.6 are
primarily in central nevous system (CNS) and 1.7, 1.8 and 1.9 in peripheral nervous
system (PNS). Nav1.4 is the primary sodium channel in skeletal muscle and Nav1.5 in
heart. Physiological and pharmacological signatures for some of these VGSCs have been
established which clearly support that different VGSCs have different physiological
characteristics in various excitable tissues. Studies on neuronal electrogenic apparatus
plasticity have confirmed that VGSCs gene expression is dynamic, like in diseased and
2-25
normal states (Nav1.8&1.9 are down regulated in injured neurons-(Dib-Hajj et al., 1998),
different functional states (Nav1.2 & 1.6 are over expressed in fast bursting state
compared to quiescent state-(Tanaka et al., 1999), and also different developmental
states. In regard to Navβ subunits, a total of 4 Navβ subunits have been discovered. They
bind to α subunits either noncovalently (β1 and β3) or by disulfide bonds (β2 and β4).
These Navβ subunits resemble the family of cell adhesion molecules (Isom et al., 1995)
and are required for localization and immobilization of VGSCs in specific locations of
excitable cells (Patino et al., 2011).
Various studies have lead to discovery of various sub-type specific toxins and
some sub-type specific blockers which may further help in discovering therapeutic
alternatives for diseases involving excitable cells and tissues. Table 1 is a compilation of
various isoforms of VGSCs discovered till date and their expressed locations in the body
and their pharmacological significance.
2.1.4 Structure of VGSCs at Atomic Resolution
A high resolution crystal structure of the bacterial sodium channel NaVAb was
determined, (Payandeh et al., 2011) revealing abundant novel information about the
structural basis for sodium selectivity and conductance. It also shed new light on the
mechanism of VGSCs block by therapeutic drugs and mechanisms of voltage-
dependence of gating. A top view of NaVAb structure revealed that, the central pore is
formed by four pore-forming modules (S5-S6 segments) and the intervening pore loop.
The outer rim of the pore module was formed by four voltage- sensing modules (S1-S4
2-26
segments). The transmembrane architecture demonstrates that the voltage-sensing
module of one subunit is closely associated with the pore-forming module of its adjacent
subunit, thereby enforcing a concerted gating of the four subunits.
2.1.5 VGSCs function: Molecular perspective
There are three important features that characterize the sodium channels: 1)
selective permeability for sodium conductance, 2) Voltage dependent activation, 3) rapid
inactivation (HODGKIN and HUXLEY, 1952a; HODGKIN and HUXLEY, 1952b;
HODGKIN and HUXLEY, 1952c).
2.1.5.1 selective permeability for sodium conductance
VGSCs pore and Ion-selectivity filter: Studies utilizing Na+ channel blockers tetradotoxin
(TTX) and saxitoxin (SXT) were responsible in identifying outer pore and selectivity
filter (Heinemann et al., 1992; Noda et al., 1989; Terlau et al., 1991). These initial studies
on pore structure were further confirmed by the crystal structure study of NavAb.
Glutamate 387 on S5-S6 inter segmental loop (membrane-reentrant loop) of domain I has
been revealed as the crucial binding amino acid (aa) for TTX and SXT. Two negatively
charged aa residues present at analogous positions on all 4 domains in S5-S6 loop form
the inner and outer rings for the TTX and SXT receptor binding site and also the
selectivity filter of the outer pore of the Na+ channel. Mutational studies of the residues
forming these rings have strongly supported the idea that these aa residues form the
selectivity filters (Schlief et al., 1996).
2-27
Though all VGSCs have similar Na+ permeation, they have different affinity for
TTX. Nav1.1, 1.2, 1.3 and 1.7 have more affinity for TTX. They are called TTX
sensitive VGSCs and are broadly expressed in neurons. Nav1.5, 1.8, 1.9 have 200 times
lesser affinity for TTX and are called TTX resistant VGSCs. These are mostly expressed
in cardiac myocytes and dorsal root ganglion neurons. This difference is due to a single
aa change at couple of residues preceding the glutamate 387 of domain 1 selectivity filter
from tyrosine/ phenylalanine to cysteine. Similarly, for cadmium (which is a high
affinity cardiac VGSCs blocker), the presence of cysteine at the above said site in the
selectivity filter of heart VGSCs attributes for stronger affinity in cardiac VGSCs over
other VGSCs.
2.1.5.2 Voltage-dependent activation
Voltage dependent activation of VGSCs was first demonstrated by Hodgkin and Huxley
and predicted that it requires the outward movement of gating charges in response to
changes in the membrane electric media (HODGKIN and HUXLEY, 1952a; HODGKIN
and HUXLEY, 1952b; HODGKIN and HUXLEY, 1952c). The S4 is a unique segment
with repeat motifs consisting of positively charged aa residues (usually arginine)
followed by 2 hydrophobic residues, thereby creating a spiral of positive charge around
the helix (Figure 2). The negatively charged internal TM electric field will strongly
attract the positive charges into the plasma membrane. Depolarization of the membrane
will lead to movement of this positively charged S4 segment in a spirally outward
direction bringing a conformational change that makes the pore open. This model is
called the sliding helix or helical screw model of voltage sensing (Catterall et al., 1986;
2-28
Catterall, 1986; DeCaen et al., 2008; DeCaen et al., 2009; DeCaen et al., 2011; Guy and
Seetharamulu, 1986; Kontis and Goldin, 1997; Kontis et al., 1997; Wang et al., 2011;
Yarov-Yarovoy et al., 2012; Zhang et al., 2011). This transmembrane position of the S4
segment was confirmed by mapping the receptor sites for scorpion toxins. β scorpion
toxin enhances the VGSCs activation by pushing the voltage dependence to much
negative membrane potentials. Upon activation the β-scorpion toxins trap the activated
S4 voltage sensor in its outward, activated position and thereby slowing down the
deactivation of the channel. Voltage sensor trapping is a common mechanism employed
by various neurotoxins like β and α scorpion toxins.
2.1.5.3 Inactivation:
VGSCs get inactivated within milliseconds of activation/depolarization. The short
conserved intracellular loop connecting III-IV domains of α subunit functions as the
inactivation gate. The aa triad of isoleucine, phenylalanine and methionine (IFM motif)
play an important role in the inactivation of the channel (West et al., 1992). The IFM
motif binds like a tether and blocks the pore by binding to the receptor site present
intracellular of the pore. (Figure 3)
2.1.5.4 Coupling of activation to inactivation:
S4 sensor outward movement (especially of domain IV) leads to activation of VGSCs
and changes in its voltage dependence leading to their inactivation. This inactivation
signal leads to closure of intracellular inactivation gate. Some toxins like α scorpion
toxin and sea anemone toxins would bind to the receptor site in such a way that it would
2-29
allow only partial movement of the S4 voltage sensor and thereby activating it but will
lead to slow inactivation. This clearly shows that activation couples with inactivation in
the normal functionality of the channel.
2.2 Voltage-gated sodium channels activators and
gating modifiers
VGSCs channel function can be altered by neurotoxins binding to various
molecular targets called receptor sites located on the channels. These receptor
neurotoxins bind with high affinity and specificity and hence provide excellent tools to
study the structure, function, conductance and gating properties of VGSCs.
Pharmacological studies have revealed that neurotoxins act on six unique receptor sites
(Table 2) with the possibility of additional two more sites (pyrethroid and antillatoxin
binding sites). Upon binding, these neurotoxins alter the Na+ conductance and voltage-
dependent gating by bringing about conformational changes at the site of binding and
thereby changing the equilibrium between open and the closed/inactive state of the
VGSCs. Also, upon binding they can alter the affinity for other neurotoxins acting at a
different receptor site. VGSCs modifiers bring about their actions primarily by 1) slowing
the coupling of sodium channel activation to inactivation (reduce the rate of inactivation),
2) increasing the mean open time of the channel 3) inhibiting channel inactivation or 4)
shifting the activation potential to more negative values, thereby augmenting Na+ influx
through VGSCs.
2-30
2.2.1 Antillatoxin
Antillatoxin (ATX) is a structurally novel lipopeptide with an exceptionally high
degree of methylation unlike any known natural product 28 Lee,K.C. 2006. Isolated
from the cyanobacterium Lyngbya majuscula, this compound is also distinguished by
multiple stereocenters 261 Orjala,J. 1995. The essential role of the asymmetric
carbon atoms in ATX is reflected in the stereoselective effects of ATX enantiomers 13
Li,W.I. 2004. ATX is considered to be the second most potent ichthyotoxic compound
obtained from marine sources after only brevetoxin (PbTx)-1 261 Orjala,J. 1995.
Exposure to L. majuscula blooms are associated with adverse human health effects,
including respiratory irritation, eye inflammation, and severe contact dermatitis. Previous
work has demonstrated that ATX is a potent activator of voltage-gated sodium channels
(VGSCs) that elevates intracellular Na+ concentration ([Na+]i) in intact neurons 24
Li,W.I. 2001; 7 Cao,Z. 2008; 296 Jabba,S.V. 2010. Moreover, ATX has been shown to
be neurotoxic in cerebellar granule cells through an indirect activation of N-methyl-d-
aspartate receptors (NMDARs) as a consequence of glutamate release 24 Li,W.I.
2001.
ATX is a novel activator of VGSC; however, its precise recognition site on the
channel protein remains to be defined. The structure of ATX includes asymmetric carbon
atoms, and the (4R,5R)-isomer is the naturally occurring compound. The (4R,5R)-isomer
appears in profile as an “L” shape with a hydrophobic interior and a cluster of
hydrophilic groups on the exterior of the macrocycle (Li et al., 2001). Thus, the (4R,5R)-
2-31
configuration is important for creating a molecular topology that is recognized by the
acceptor site on the voltage-gated sodium channel α subunit. Previous studies from our
lab have shown that ATX allosterically enhances the specific binding of
[3H]batrachotoxin to intact cerebellar granule cells (Li et al., 2001). This effect of ATX
on [3H]batrachotoxin binding was synergistically augmented by brevetoxin. The strong
synergistic interaction of the ATX recognition site with neurotoxin site 5 suggests that
these sites may be topologically close and/or conformationally coupled. The results
obtained using [3H]batrachotoxin as a probe for sodium channel conformation allowed us
to exclude the interaction of ATX with neurotoxin sites 1, 2, 3, and 5 on VGSCs. Site 1
was ruled out because tetrodotoxin and saxitoxin bind to the outer vestibule of the pore of
the ion channel and allosterically inhibit the binding of [3H]batrachotoxin; this is an
effect that is antipodal to that of ATX. We were able to rule out sites 2 and 5 inasmuch as
these sites display positive allosteric coupling to the ATX site. Neurotoxin receptor site 3,
the target for α-scorpion toxins and sea-anemone toxins, was excluded because ATX
enhanced [3H]batrachotoxin binding in the presence of a maximally effective
concentration of sea-anemone toxin. Although we cannot exclude an interaction of ATX
with neurotoxin site 4, the target for β-scorpion toxin, it is reasonable to posit that ATX
binds to a novel recognition domain on the α-subunit of the VGSC. The relatively small
lipotripeptide structure of ATX would not be restricted to an extracellular target, as is the
case for the scorpion toxins, which are composed of 60 to 65 amino acids. Given the
unique structure and mechanism of action of ATX, it would be important to further
characterize its binding site on VGSCs.
2-32
Most recent study on ATX has demonstrated that it displays a unique efficacy
with respect to stimulation of sodium influx in cells expressing rNav1.2, rNav1.4 and
rNav1.5 α-subunits (Cao et al., 2011). The efficacy of ATX was distinctive inasmuch as it
was not shared by activators of neurotoxin sites 2 and 5. It was also demonstrated that
the ATX binding site shares with neurotoxin sites 2 and 5 the phenomenon of partial
agonism. Finally, it was observed a reciprocal allosteric interaction between neurotoxin
site 5 and the ATX binding site. Collectively, these data indicate that ATX is a sodium
channel gating modifier with unique efficacy in cells heterologously expressing VGSC α-
subunits. Defining the molecular determinants and mechanisms of action of ATX may
provide further insight into the gating properties of sodium channels.
Our lab previously demonstrated that NMDA receptor function may be increased
through activation of VGSCs with attendant elevation of intracellular sodium in
cerebrocortical neurons (George et al., 2009). VGSC activators function as gating
modifiers that elevate [Na+]i in the absence of substantial depolarization of neurons (Cao
et al., 2008; George et al., 2009). These findings have been confirmed and extended for
this structurally novel lipopeptide, ATX. Jabba et al., found that ATX promoted
neuritogenesis by elevating [Na+]i, which in turn augmented NMDAR function leading to
Ca2+ influx and engagement of a CaMKK pathway (Jabba et al., 2010) which stimulated
neuritogenesis in DIV-1 cerebrocortical neurons. These data provide further support for
the hypothesis that sodium channel activators seem to be capable of mimicking activity-
dependent neuronal development through potentiation of NMDAR signaling pathways
that influence neuronal plasticity.
2-33
2.2.2 Veratridine
Veratridine (VRT) is a lipid-soluble alkaloid extracted from plants of family Liliaceae
(suborder Melanthaceae) belonging to genus Veratrum (species album, viride etc.).
Veratrine, the alkaloid fraction of the seeds is a mixture that mostly consists of the ester
alkaloids, veratridine, and cevadine and of the alkamine veracevine or its isomer cevine.
Veratridine binds to receptor site 2 of VGSCs and preferentially to activated state of
VGSCs, thereby causing them to stay open (persistent activation via allosteric
mechanism) during a sustained membrane depolarization. This leads to abolishment of
VGSCs inactivation and shift of the voltage dependence of activation to more negative
potentials 395 Ulbricht,W. 1969; 384 Albuquerque,E.X. 1988; 394 Catterall,W.A.
1975; 393 Catterall,W.A. 1975; 392 Catterall,W.A. 1980; 391 Catterall,W.A. 1980; 389
Khodorov,B.I. 1985. This primarily leads to Na+ influx, with secondary effects like
increased Na+-K+ pump activity, increased Ca2+ influx and in turn exocytosis .
Veratridine is a partial agonist at site 2 with batrachotoxin being the full agonist. Most of
the experiments involving localization of receptor site-2 utilized batrachotoxin and
determined that site 2 neurotoxins bind to the S6 TM region of domain I 359
Trainer,V.L. 1996. It is thought that the abolishment of VGSCs inactivation by VRT
and other site 2 neurotoxins is due to its interaction with S6 TM of domain IV (IVS6) that
is required for fast inactivation. Also, site 2 neurotoxins binding to IVS6 may cause the
toxin to change the voltage-dependent movements of the adjacent IVS4 voltage-sensor
and thereby affect activation and coupling of activation to inactivation 351
Linford,N.J. 1998.
2-34
2.2.2.1 Intracellular sodium as a signaling molecule and its involvement in neuronal
development.
The electrical signals of neurons are fundamentally dependent on Na+ influx
through VGSCs and are primarily responsible for the rising phase of action potential and
hence supply the current that drives the membrane potential to the peak of action
potential. Na+ is a major physiological ion present in the extracellular milieu. Apart from
action potential dependent Na+ influx, Na+ can enter into the cells via various routes
including influx through voltage and ligand-gated ion channels, uptake via membrane
exchangers and gradient-driven co-transporters 252 Nicholls,D. 1990. Recent studies
have indicated that changes in intracellular sodium concentration ([Na+]i) produced in the
soma and dendrites as a result of neuronal activity may act as a signaling molecule and
play a role in activity-dependent synaptic plasticity. It has been shown that short burst or
tectonic stimulation of afferents that induce synaptic LTP causes [Na+]i increments of 10
mM in dendrites and of up to 35-40 mM in dendritic spines 93 Rose,C.R. 1999; 91
Rose,C.R. 2001 . In cerebellar Purkinje neurons, α-amino-3-hydroxy-5-methyl-4-
isoxazolepropionic acid receptor-mediated Na+ influx was shown to be required for
induction of long-term depression 249 Linden,D.J. 1993. In both hippocampal and
immature cerebrocortical neurons, an elevation in intracellular Na+ was found to increase
NMDAR-mediated whole-cell currents and NMDAR single-channel activity by
increasing either channel open probability or mean open time or both (Figure 3) 96
Yu,X.M. 1998; 296 Jabba,S.V. 2010; 6 George,J. 2009. Yu and Salter have
demonstrated that an increment of [Na+]i of 10 mM was sufficient to produce significant
2-35
increases in NMDA receptor single-channel activity. It has also been reported that
increments of [Na+]i greater than 5 mM represent a critical threshold required to regulate
NMDAR-mediated Ca2+ influx. They used veratridine, a VGSC modulator, to
demonstrate that Na+ influx through TTX-sensitive VGSC was sufficient to up-regulate
NMDAR activity. Moreover, this [Na+]i-mediated up-regulation of NMDAR function has
been shown to require Src kinase activation 6 George,J. 2009; 96 Yu,X.M. 1998. In
DIV 1 dorsal root ganglion (DRG) neurons obtained from E7 pups, Semaphorin3A-
induced facilitation of axonal guidance involved VGSCs activation and required changes
in [Na+]i 309 Yamane,M. 2012. [Na+]i downregulated the amiloride-sensitive
currents in epithelial sodium channels (ENaCs) 433 Cook,D.I. 2002; 434 Dinudom,A.
2001; 436 Komwatana,P. 1996. Further, studies have demonstrated [Na+]i may also
activate potassium channels 432 Bhattacharjee,A. 2005; 431 Dryer,S.E. 2003. Some
studies have found the role of [Na+]i in regulation of ligand-receptor and ligand-
transporters interactions 440 Pert,C.B. 1974; 439 Pert,C.B. 1974; 438 Bloch,R.J. 1986;
437 Werling,L.L. 1986; 435 Puttfarcken,P. 1986. Also, a significant increase in [Na+]i
is a characteristic feature of tissue injury and various studies have demonstrated the
neuro-protectiveness of blocking Na+ influx during tissue injury. The detailed mechanism
of action for the above listed affects of Na+ are not clearly evident and require further
studies are very important to elucidate them. One important possibility is the manifold
influx of Ca2+ into cytoplasm with subsequent activation of Ca2+-dependent activation.
Also, Na+ entry could change intracellular pH 443 Boonstra,J. 1983; 442
Moolenaar,W.H. 1983; 441 Moolenaar,W.H. 1986, thereby regulating enzymatic
2-36
activity, neuronal growth and death. Hence, it would be critical to determine these
mechanisms and elucidate the potential Na+ binding target sites.
2.3 N-methyl-D-aspartate receptors (NMDARs)
NMDARs are located at excitatory glutamate synapses in the central nervous system
and play several important roles including, but not limited to, excitatory synaptic
transmission, neuronal plasticity, learning and memory formation and excitotoxicity.
They possess unique features like voltage-dependent block by extracellular Mg2+, high
Ca2+ permeability and slow activation/deactivation kinetics. Though NMDARs are
sensitive to various endogenous ligands, a critical requirement for their activation is
binding to the co-agonist glycine. Others that modify the receptor activity are
extracellular Zn2+, polyamines, interactions with scaffolding, anchoring and signaling
molecules associated with postsynaptic density (PSD). Numerous subtypes of NMDARs
have been identified in CNS, distinct in their channel properties, including, but not
limited to, ligand sensitivity, divalent ion block, kinetics, and interaction with
intracellular proteins. Understanding the role of various subtypes of NMDARs in normal
CNS functions like neuronal development, learning and memory and synaptic plasticity
will be important in developing therapeutic strategies for various pathophysiological
conditions involving NMDARs like epilepsy, ischemic brain damage, schizophrenia,
depression and, more speculatively, neurodegenerative disorders such as Parkinson’s and
Alzheimer’s diseases, Huntington’s chorea, and amyotrophic lateral sclerosis.
2-37
2.3.1.1 NMDARs subunits and composition:
Electrophysiological, binding and cloning experiments provided the first evidence for
the presence of diversity in NMDARs and also regarding the heterologous composition of
the NMDARs in forming a functional NMDAR. Three families of NMDAR subunits
having significant sequence homology with other ionotrophic glutamate receptors were
discovered: GluN1, GluN2 (4 distinct GluN2’s-A, B, C and D) and GluN3 (2 different
GluN3’s) (previously called NR1, NR2 and NR3 respectively) 378 Moriyoshi,K. 1991;
368 Monyer,H. 1994; 363 Mori,H. 1995 with each having several splice variants.
NMDARs function as hetertetrameric assemblies, usually comprising of 2 obligatory
GluN1 subunits and 2 GluN2 subunits. GluN3 containing NMDARs contain
diheteromeric (GluN1/GluN3) or triheteromeric (GluN1/GluN2 / GluN3) complexes
430 Traynelis,S.F. 2010. The presence of such diverse NMDAR subunits allow for
various combinations of functionally distinct NMDAR subunit assembly.
The obligatory GluN1 subunit is ubiquitously expressed in the CNS, both in embryo
and adult, but with developmental and spatial variations in regard to GluN1 isoform
expressed. In contrast, GluN2 subunits have expression patterns that differ in
developmental time and space (one brain region to other) 369 Watanabe,M. 1992; 367
Akazawa,C. 1994; 368 Monyer,H. 1994; 373 Monyer,H. 1992. In embryonic CNS,
GLuN2B and GluN2D are preferentially expressed, with GluN2B being widely expressed
all over the brain and with GluN2D exclusively expressed in diencephalon and brain
stem. Postnatal, GluN2 subunit expression undergoes a drastic turnover, especially during
the first two weeks of birth, with gradual increase in GluN2A subunit expression in entire
2-38
CNS. GluN2B expression reaches maximum at P7-10 but gets restricted more to the fore
brain regions (cortex, hippocampus, striatum, olfactory bulb) and remains high
throughout at these regions. GluN2A subunit expression increases steeply at around P7-
10. GluN2C subunit expression starts around P7, exclusively in the cerebellum granule
cells (CGC) and olfactory bulb neurons. GluN2D subunit expression is drastically
reduced postnatally, with little expression in the diencephalon and brainstem in adult life.
These developmental switches in GluN2 subunit expression patterns are important
functionally and bring about a change in the kinetics and sensitivity of NMDAR
mediated currents to various endogenous GluN ligands.
2.3.1.2 NMDAR Transmembrane topology:
NMDARs are integral membrane proteins, and as described earlier, form
heteroterameric channels with a central ion channel pore selective for Na+, Ca2+ and K+
cations. Each GluN subunit consists four distinct domains 81 Mayer,M.L. 1984; 325
Paoletti,P. 2007, 1)N-terminal domain (NTD): first 380 amino acids in the extracellular
region which forms a large clamshell-like domain, 2) Agonist binding domain (ABD): a
300 amino acids region that binds glycine in GluN1 and glutamate in GluN2 subunits, 3)
membrane domain: consists of 3 TM segments (M1, M3 and M4) and a short re-entrant
loop (pore or P-loop; M2) that forms the ion channel, and 4) An intracellular C-terminal
domain: highly variable in length (subunit dependent) and involved in scaffolding,
interaction with signaling molecules and receptor trafficking.
2-39
2.3.1.3 NMDARs role in neuronal development:
Neuronal activity has a major role in the development of dendritic complexity and
neuronal circuits. The mechanisms by which neuronal activity translate into
morphological changes are complex. Numerous studies have shown that activity-
dependent neuronal development involves NMDARs mediated calcium influx 254
Ghosh,A. 1995; 32 West,A.E. 2002. Calcium influx through the NMDA receptor and
the subsequent initiation of signaling pathways have a well established role in activity-
dependent neuronal development and plasticity. NMDAR-mediated increase in cytosolic
Ca2+ also involves calcium release form intracellular stores (ER) and is involved in local
effects like dendrtic branching and stabilization of dendritic structures37 Redmond,L.
2002 . Intracellular calcium acts as a signaling molecule largely through binding to
calmodulin, a calcium-binding protein that engages downstream Ca2+/calmodulin-
dependent protein kinase (CaMK) and mitogen-activated protein kinase (MAPK)
signaling pathways 254 Ghosh,A. 1995; 32 West,A.E. 2002. Also, NMDAR-
mediated activity-dependent signaling to the nucleus may be influenced by the
developmental regulation of NMDAR subunit composition. Similarly, NMDAR-
dependent changes in synaptic function and calcium signaling can mediate the
developmental regulation of activity-dependent transcriptional programs 328
Majdan,M. 2006. Moreover, previous studies have demonstrated that activity-
dependent neurite outgrowth 99 Wayman,G.A. 2006 and synaptogenesis 98
Saneyoshi,T. 2008 are regulated by NMDAR-dependent CaMKK/calmodulin kinase I-
signaling cascades. Also, Epherin B (EphB) receptor tyrosine kinases regulates NMDARs
2-40
function, thereby altering Ca2+ influx mediated gene expression, providing further
evidence for the role of NMDARs in neuronal development 344 Dalva,M.B. 2000.
Moreover, EphB interaction with GluN1 subunit of NMDARs during synaptic
development and plasticity plays an important role in NMDAR-dependent induction of
LTP and LTD 342 Grunwald,I.C. 2001; 341 Henderson,J.T. 2001. More
importantly, Src family tyrosine kinases phosphorylation of GluN2B subunit leads to
NMDARs function up-regulation, thereby increasing activity-dependent gene expression
340 Takasu,M.A. 2002. Moreover, NMDAR function is up-regulated due to changes
in [Na+]i and it has been shown that this requires Src kinase activation96 Yu,X.M.
1998; 6 George,J. 2009 . Src family kinases (SFKs) act as a crucial point of
convergence for signaling pathways that enhance NMDAR activity, and, by up-regulating
the function of NMDARs, Src gates the production of NMDAR-dependent synaptic
potentiation and plasticity97 Salter,M.W. 2004 .
Therefore, NMDARs play a critical role in activity-dependent development and
plasticity (Ghosh and Greenberg, 1995), dendritic arborization 99 Wayman,G.A. 2006;
1 Wong,R.O. 2002; 84 Miller,F.D. 2003, spine morphogenesis 36 Ultanir,S.K.
2007 and synapse formation 98 Saneyoshi,T. 2008 by stimulating these NMDAR-
mediated calcium-dependent signaling pathways.
2-41
2.3.2 Neurotrophins and their receptors, tropomyosin-related kinases
(TRKs):
Neurotrophins play an important role in regulating neural survival, development,
function and plasticity. The discovery of neurotrophins and its cognate receptor marked a
breakthrough in developmental neurobiology. These biochemicals that are secreted in
small quantities regulate both local and global effects, for e.g., growth cone motility and
gene transcription respectively.
As of now, six neurotrophins are identified, out of which, four have been characterized.
1) Nerve growth factor (NGF), 2) Brain-derived neurotrophic factor (BDNF), 3)
Neurotrophin-3 (NT-3), and 4) Neurotrophin-4 (NT-4). All four neurotrophins have
similar sequence and structure and originate from a common ancestral gene. Their genes
share various similarities, including the presence of multiple promoters. The protein
product consists of three different sequences connected in a row. They are 1) signal
sequence 2) prodomain sequence, and 3) mature neurotrophin sequence. Hence, each
gene product has to undergo proteolysis to form a mature protein.
The neurotrophins interact with two different classes of receptors. Primarily, they
activate receptor tyrosine kinases called tropomyosin-related kinases (TRKs) 269
Huang,E.J. 2003 which in turn regulate signaling pathways involved in neuronal
survival, proliferation, neurite and axonal outgrowth, dendritic aroborization and
remodeling, cytoskeleton rearrangement, membrane trafficking and synaptogenesis and
synaptic plasticity. Trk receptors consist of three domains 1) extracellular domain, 2)
2-42
transmembrane domain, and 3) cytoplasmic domain. The extracellular domain consists of
a) cysteine-rich cluster followed by b) three leucine-rich repeats, c) another cysteine-rich
cluster and d) two immunoglobulin-like domains. The cytoplasmic domain consists of a
tyrosine kinase domain and is surrounded by several tyrosines. These tyrosines upon
phosphorylation interact with several cytoplasmic adaptors and enzymes. Upon binding
with neurotrophins, Trk receptors dimerize, leading to the transphosphorylation
activation of the kinases present in the cytoplasmic domain. Three distinct TRKs have
been identified, TrkA, B, and C. Neurotrophins binds to their cognate receptor in an
autocrine or/and pracrine manner, subsequently activating them. NGF binds to TrkA and
activates its tyrosine kinase activity. Similarly BDNF and NT-4 activates TrkB, and NT-3
activates TrkC 380 Kaplan,D.R. 1991; 381 Klein,R. 1991; 379 Klein,R. 1991; 269
Huang,E.J. 2003; 345 Bibel,M. 2000. In addition, NT-3 can activate other Trk
receptors but with lower efficiency. Alternate splicing of the receptors will introduce
additional short amino sequences to the extracellular domain, thereby affecting their
ligand interactions 366 Clary,D.O. 1994; 376 Meakin,S.O. 1992; 358 Strohmaier,C.
1996. Similarly, splicing can also introduce changes in the intracellular catalytic
domain and thereby affecting signaling mechanisms initiated by neurotrophin binding.
TrkB and TrkC isoforms generated by alternate slicing include those which contain
comparatively short cytoplasmic motifs with no tyrosine kinase domains. Those isoforms
that lack the kinase domain inhibit productive dimerization with kinase-containing Trk
receptors, thereby inhibiting responses to neurotrophins 429 Eide,F.F. 1996. More
recent works, however, have demonstrated that these isoforms have a role to play in
2-43
regulating various cellular signaling mechanisms. BDNF mediated activation of T1
isoform of TrkB controls release of Ca2+ from intracellular stores in a G-protein and IP3-
dependent pathway 336 Rose,C.R. 2003. Differential splicing also affects the
substrate specificity of one of the TrkC isoform, inhibiting its activation of several
substrates and thereby its ability to promote neuronal differentiation 364 Guiton,M.
1995; 356 Meakin,S.O. 1997.
Neurotrophins also interact with another family of receptor called the p75
Neurotrophin receptor (p75NTR). These are low affinity receptors, but bind to all
neurotrophins with almost same affinity 382 Rodriguez-Tebar,A. 1990; 353 Frade,J.M.
1998. p75NTR belongs to tumour necrosis receptor superfamily and its structure
consists of three domains. 1) an extracellular domain containing four cysteine-rich
motifs, 2)a transmembrane domain, and 3) cytoplasmic domain, that includes the ‘death’
domain 357 Liepinsh,E. 1997. Although, p75NTR do not have a catalytic domain, it
can interacts with several proteins that play an important role in neuronal survival and
development and synaptic plasticity 330 Makkerh,J.P. 2005; 346 Bentley,C.A. 2000;
347 Harrison,S.M. 2000.
2.3.2.1 Brain Derived Neurotrophic Factor (BDNF):
BDNF is one of the best characterized neurotrophin that plays a critical role in neuronal
development and plasticity. More recent studies in the field of neurotrophins have
associated BDNF deficits with various neurodevelomental, neurodegenerative and
psychiatric disorders. BDNF is widely expressed in most brain areas, with high
2-44
expression in the hippocampus, amygdale, and cerebellum with principal glutamatergic
neurons being the main cell type expressing it 324 Aid,T. 2007; 418 Ernfors,P. 1990;
420 Ernfors,P. 1990; 424 Lindholm,D. 1992. An activity-regulated gene, Bdnf gene is
composed of at least nine exons and is transcribed by many unique promoters leading to
synthesis of distinct 5’untranslated regions (UTRs). Exons I to VIII have their own
distinct promoter and are associated by splicing mechanism to exon IX, which is the only
one being translated into a protein. Also, these mRNA synthesized have two distinct
3’UTRs, with differences due to the presence of two distinct polyadenylation sites,
leading to expression of Bdnf gene with 2 different 3’UTR lengths 324 Aid,T. 2007.
Due to a combination of several 5’UTRs, 10 different exons and 2 distinct 3’UTRs, there
is a possibility of at least 18 distinct pre-mRNA being transcribed, but all translated into
one BDNF protein. This transcription is highly regulated, in a developmental age wise,
tissue-specifically and neuronal activity-dependent manner 332 Rattiner,L.M. 2005;
324 Aid,T. 2007. For e.g., neuronal activity-dependent and Ca2+ influx mediated Bdnf
transcription is controlled by promoter I and IV 321 Kidane,A.H. 2009; 332
Rattiner,L.M. 2005. Bdnf gene with long 3’UTRs are transported to the dendrites and
locally translated to the BDNF protein 417 An,J.J. 2008, where as the short 3’UTR
containing Bdnf gene is localized to the soma. Though most genes that are locally
translated are tightly regulated by local synaptic activity, mechanisms of regulation of
Bdnf gene local translation in the dendrites is not well established. Also, other
mechanisms of transcriptional control of BDNF are not well understood, and given the
utmost biological significance of BDNF in neuronal development and plasticity, it will be
2-45
very important to focus future studies on transcriptional regulation of Bdnf gene synthesis
in the nervous system.
The pre sequence of the pre-RNA directs the translation of the BDNF to endoplasmic
reticulum, giving rise to a dimeric pro-BDNF (29 kDa) that is involved in the sorting of
BDNF into appropriate pathway of secretion 333 Chen,Z.Y. 2005; 287 Lee,C.C.
2005. This pro-BDNF is later cleaved to give rise to mature BDNF 360 Seidah,N.G.
1996. Two different pathways are utilized in the secretion of BDNF 1) constitutive and
2) regulated pathways. In the constitutive pathway, BDNF is packaged into smaller sized
granules (Ø 50-100 nm) and released continuously, in a Ca2+-independent fashion. This
pathway occurs mostly at the soma and proximal parts of the neuron and does not involve
of any specific triggering stimulus 349 Mowla,S.J. 1999; 337 Lessmann,V. 2003; 331
Brigadski,T. 2005. In the regulated pathway, BDNF is packaged into more bigger
sized vesicles (Ø ~300 nm), and following an increase in [Ca2+]i these vesicles fuse to the
plasma membrane and release BDNF. This pathway occurs primarily at the distal parts of
the neuronal processes 331 Brigadski,T. 2005. As mentioned earlier, pro-BDNF is
converted into mature BDNF in the golgi and trans-golgi network and then sorted for
these two different pathways. Two different amino acid motifs, each present in the pro
and mature form are involved in this sorting process. The interaction of the motif
specifically present in the mature form with carboxypeptidase E (CPE) targets it to the
regulated secretory pathway and an absence of the CPE interaction leads to the BDNF
sorted to the constitutive pathway 416 Pang,P.T. 2004 . Similarly, interaction of the
motif specifically present in the pro form with sortilin targets it to the regulated secretory
2-46
pathway. A single nucleotide polymorphism in the motif specifically present in the pre
form (val66met) will lead to the loss of interaction with sortilin, thereby impairing the
sorting mechanisms to the regulated secretory pathway and causing deficits in neuronal
development and cognition 339 Egan,M.F. 2003. Even though both pathways release
both pro and mature form, studies have shown that, comparatively more mature BDNF is
released by the regulated pathway than constitutive pathway. The possible reason is, in
regulated pathway, the time between synthesis and release of BDNF is longer, thereby
providing more time for the intracellular convertases to convert the pro form to mature
form. Vice versa, in constitutive pathway, more pro-BDNF is released due to the less
available time for the convertases to convert the pro form to mature form.
Several studies in the last decade have demonstrated that, BDNF release in regulated
pathway requires increase in [Ca2+]i. The route for increase in [Ca2+]i can be manifold.
Calcium influx through NMDARs or VGCCs or release of Ca2+ from intracellular
calcium stores or a combination of any of the above (depending on the kind of
stimulation or activation), can trigger the release of BDNF. Mechanistic studies on BDNF
release have demonstrated that the rise in the calcium activates the calcium calmodulin II
(CamKII) leading to the fusion of BDNF containing secretory granules with the plasma
membrane and slowly releasing BDNF into the extracellular milieu 331 Brigadski,T.
2005; 323 Kolarow,R. 2007. It has been also demonstrated that protein kinase A
(PKA) have a regulatory role in the release of BDNF through regulated pathway 323
Kolarow,R. 2007 in the dendritic neuronal processes.
2-47
2.3.3 Regulation of BDNF-TrkB signaling pathway
Upon binding with neurotrophins, Trk receptors dimerize, leading to the
transphosphorylation activation of the kinases (activation loop tyrosines) present in the
cytoplasmic domain 415 Huang,E.J. 2003. Proteases and convertases that control the
processing of pro-BDNF to mature-BDNF regulate TrkB receptor responsiveness to
BDNF, as only mature form of BDNF and not the pro-forms of BDNF can activate TrkB
receptors 400 Lee,R. 2001. The TrkB receptor contain additional tyrosine residues in
the cytoplasmic domain that act as phosphorylation substrates for TrkB tyrosine kinase.
These phosphorylated tyrosines then interact with various scaffolding proteins and other
intermediary proteins that are part of intracellular signaling cascades. Y670, Y674 and
Y675 (human TrkA sequence) are present in the autoregulatory loop of the tyrosine kinse
domain. Phosphorylation of these tyrosines increases the Trk tyrosine kinase activity.
Phosphorylation of other tyrosine residues leads to the formation of binding sites for
proteins containing phosphor-tyrosine binding (PTB) or Src-homology-2 (SH2) domains.
Three major intracellular signaling pathways are activated by BDNF-TrkB. They are: 1)
PI3K-Akt pathway, 2) the Ras-MAPK pathway and 3) PLCγ-Ca2+ pathway (Reichardt,
2006). BDNF-TrkB signaling mediated functional changes can involve any one of these
or a combination of these signaling cascades.
2.3.3.1 Phospholipase-C signaling cascade
Activation of TrkB at Y816 recruits and activates PLCγ which in turn hydrolyzes
phosphotidylinositol 4,5-biphosphate (PIP2) to diacylglycerol (DAG) and inositol 1,4,5-
2-48
triphosphate (IP3), where DAG activates protein kinase C (PKC) and IP3 releases Ca2+
from intracellular stores 415 Huang,E.J. 2003; 399 Reichardt,L.F. 2006. Together
these signaling molecules activate many intracellular enzymes, including most PKC
isoforms, Ca2+/CamKs and other Ca2+/Cam-regulated targets. Activation of PLCγ1
activates PKCδ leading to neurotrophin-mediated activation of MEK1 and Erk1/2 401
Corbit,K.C. 1999. Also, BDNF-TrkB-PLC γ1 signaling activates transient receptor
potential canonical subfamily of cation channels 3/6 (TRPC3/6) and contributes to
BDNF-induced Ca2+ elevations at growth cones and synapses 403 Li,Y. 2005; 402
Amaral,M.D. 2007. In nociceptive sensory neurons, the capsaicin receptor VR1, the
heat-activated TRP channel is repressed due to depletion of PIP2. Since activation of Trk-
PLC γ1 signaling leads to depletion of PIP2, Trk receptor activation leads to
hypersensitization of this channel to thermal and mechanical stimuli 404 Chuang,H.H.
2001; 405 Prescott,E.D. 2003. Also, some recent studies have implicated the role of
BDNF-TrkB-PLC γ1 signaling induced Ca2+ increase in synaptic plasticity 406
Nakata,H. 2007; 407 Shaywitz,A.J. 1999.
2.3.3.2 Mitogen-activated protein kinase pathway
The other TrkB phosphorylation site is Tyr515, which activates downstream MEK-
MAPK/Erk signaling and promotes neuronal differentiation and growth by influencing
transcription events, such as activation of CREB transcription factor. Phosphorylation of
Y515 on TrkB by BDNF recruits Shc to TrkB and phosphorylates it. This leads to Shc
interaction with Grb2, an adaptor protein that recruits and activates the guanine
nucleotide exchange factor (GEF) SOS. Subsequently SOS activates Ras which thereby
2-49
activates downstream kinases B-raf, MEK and MAPK/Erk 399 Reichardt,L.F. 2006;
415 Huang,E.J. 2003. MEK-MAPK/Erk signaling pathway activates transcription
factors, such as CREB 407 Shaywitz,A.J. 1999. MAPK/Erk plays a critical role in
protein synthesis dependent plasticity by increasing phosphorylation of eukaryotic
initiation factor 4E (eIF4E), the 4E-binding protein 1 (4E-BP1) and ribosomal protein S6
409 Kelleher,R.J.,3rd 2004; 408 Klann,E. 2004.
2.3.3.3 Phosphotidyl-Inositol-3-kinase pathway
Phosphorylation of Tyr 516 indirectly activates PI3K pathway by interacting with
adaptor proteins (Shc-Grb2-Ras) that regulate PI3K pathway 399 Reichardt,L.F.
2006. PI3K activation leads to the changes in inositol phospholipids composition in the
plasma membrane (cytoplasmic side), resultingin translocation of Akt/protein kinase B
(PKB) to the plasma membrane. Activated Akt/PKB is involved in a variety of functions
such as cell survival, proliferation, neuronal differentiation, neurite outgrowth and protein
translation 410 Read,D.E. 2009; 415 Huang,E.J. 2003; 399 Reichardt,L.F. 2006. The
TrkB-PI3K-Akt mediated protein translation is mediated by signaling cascade called
mammalian target of rapamycin (mTOR), a mjor regulator of protein synthesis 411
Sarbassov,D.D. 2005; 412 Sarbassov,D.D. 2005. Akt activation of mTOR pathway
involves, phosphorylation and inhibition of tuberous sclerosis complex proteins: hamartin
(TSC1) and tuberin (TSC2). TSC1/2 complex is a GTPase-activating protein for Ras
homolog enriched in brain (Rheb), immediate upstream activator of mTOR. PI3K-Akt
mediated inhibition of TSC1/2 complex increases activation of Rheb which subsequently
causes mTOR activation 413 Jaworski,J. 2006. Active mTOR phosphorylates p70S6
2-50
and 4E-BP1 leading to mRNA translation. Also, The PI3K-Akt pathway plays an
important role in long-term maintenance of synaptic plasticity by regulating the
trafficking of synaptic proteins 414 Yoshii,A. 2007.
Activation of TrkB at Y816 recruits and activates PLCγ which in turn hydrolyzes
PIP2 to DAG and IP3, where IP3 releases Ca2+ from intracellular stores. The other TrkB
phosphorylation site is Tyr515 that activates downstream MEK-MAPK/Erk signaling,
which promotes neuronal differentiation and growth by influencing transcription events,
such as activation of CREB transcription factor. Tyr 516 indirectly activates PI3K
pathway by interacting with adaptor proteins that regulate PI3K pathway. Another
pathway that is indirectly activated by Phospho-TrkB (Y516) is the PI3K-Akt pathway.
2.3.4 Role of BDNF-TrkB signaling in neuronal development
Neurotrophins cooperate with neuronal activity to modulate neuronal morphology.
Studies in the past few years have shown convincing evidence for the involvement of
BDNF-TrkB signaling in activity-dependent dendritic growth, dendric arborization,
synaptogenesis and synaptic transmission 399 Reichardt,L.F. 2006415 Huang,E.J.
2003; 414 Yoshii,A. 2007. BDNF-TrkB mediates increase in dendritic complexity by
increasing the total dendritic/neurite length, dendritic branching (arborization) and
increase in number of primary neuritis 295 Baker,R.E. 1998; 275 McAllister,A.K.
1996. Activation of NMDARs and intracellular Ca2+ rise is critical for the expression
and release of BDNF399 Reichardt,L.F. 2006. The released BDNF binds to its
cognate receptors TrkB and acts in an autocrine/paracrine mode to regulate neuronal
2-51
development. Through BDNF-TrkB-PI3K-Akt pathway, Akt can subsequently
phosphorylate a number of substrates (GSK3β, MAP2, mTOR) that are involved in cell
survival, NOG/actin dynamics, dendritogenesis and synaptic plasticity 410 Read,D.E.
2009; 414 Yoshii,A. 2007.
2-52
Table 1: Location and pharmacological properties of various VGSCs isoforms.
Channel name Location Pharmacological significance Nav1.1 Primarily in the cell
bodies of the central neurons; cardiac myocytes
Site of action for epileptic drugs; Site for local anesthetics that enter circulation and Cerebro spinal fluid (CSF) and thereby causing side effects.
Nav1.2 Primarily in the myelinated and premyelinated axons of central neurons of the central neurons
Site of action for epileptic drugs; Site for local anesthetics that enter circulation and CSF and thereby causing side effects.
Nav1.3 Cell bodies of central neurons in embryonic and prenatal life: Cardiac myocytes
Site of action for epileptic drugs; Site for local anesthetics that enter circulation and CSF and thereby causing side effects.
Nav1.4 Skeletal muscle Site for local anesthetics treating myotonia
Nav1.5 Cardiac myocytes; immature and denervated skeletal muscle; Some neurons in the brain
Site for antiarrhythmic drugs; Site for local anesthetics that enter circulation and thereby causing side effects.
Nav1.6 Cerebellum, cerebral cortex, brainstem, spinal cord; DRG; nodes of Ranvier in PNS and CNS.
Target for anti epileptic drugs and analgesics.
Nav1.7 DRG neurons, Schwann cells, sympathetic neurons; neuroendocrine cells
Site for local anesthetics in PNS.
Nav1.8 Small and medium sized DRG neurons and axons.
Target for Analgesic drugs
Nav1.9 Mostly in the nociceptive DRG neurons (c-type); trigeminal neurons
Target for Analgesic drugs
2-53
Table 2: Receptor sites on VGSCs (modified from Catterall et al., Pharmacol Rev. 2005)
Receptor Site Toxin or Drug Domains
Neurotoxin receptor site 1 Tetrodotoxin IS2–S6, IIS2–S6
Saxitoxin IIIS2–S6, IVS2–S6
µ-Conotoxin
Neurotoxin receptor site 2 Veratridine IS6, IVS6
Batrachotoxin
Grayanotoxin
Neurotoxin receptor site 3 α -Scorpion toxins IS5–IS6, IVS3–S4
Sea anemone toxins IVS5–S6
Neurotoxin receptor site 4 β-Scorpion toxins IIS1–S2, IIS3–S4
Neurotoxin receptor site 5 Brevetoxins IS6, IVS5
Ciguatoxins
Neurotoxin receptor site 6 δ -Conotoxins IVS3–S4
Local anesthetic receptor site Local anesthetic drugs IS6, IIIS6, IVS6
Antiarrhythmic drugs
Pyrethroid insecticide receptor site
Deltamethrin and other pyrethroids Unknown site
2-54
Figure 2-1: Subunit structure of VGSCs.
( Source: Yu & Catterall, 2003, Genome Biology 2003, 4:207)
Figure 2-2: A) Voltage dependent activation: outward movement of S4 Voltage sensors
(Source: Yu & Catterall, 2003, Genome Biology)
A
2-55
B) Voltage sensor trapping model of β scorpion toxin action. (Source: Catterall WA,
2002, Nov Found symp.)
Figure 2-3: Intracellular NA+ may act as a signalinging molecule and upregulate
NMDARs. (Courtesy Yu XM, 2006)
2-56
2.4 References
Aid T, Kazantseva A, Piirsoo M, Palm K, Timmusk T (2007) Mouse and rat BDNF gene
structure and expression revisited. J Neurosci Res (United States) 85:525-535.
Akazawa C, Shigemoto R, Bessho Y, Nakanishi S, Mizuno N (1994) Differential
expression of five N-methyl-D-aspartate receptor subunit mRNAs in the cerebellum of
developing and adult rats. J Comp Neurol (UNITED STATES) 347:150-160.
Albuquerque EX, Daly JW, Warnick JE (1988) Macromolecular sites for specific
neurotoxins and drugs on chemosensitive synapses and electrical excitation in biological
membranes. Ion Channels (UNITED STATES) 1:95-162.
Amaral MD, Chapleau CA, Pozzo-Miller L (2007) Transient receptor potential channels
as novel effectors of brain-derived neurotrophic factor signaling: Potential implications
for rett syndrome. Pharmacol Ther (England) 113:394-409.
An JJ, Gharami K, Liao GY, Woo NH, Lau AG, Vanevski F, Torre ER, Jones KR, Feng
Y, Lu B, Xu B (2008) Distinct role of long 3' UTR BDNF mRNA in spine morphology
and synaptic plasticity in hippocampal neurons. Cell (United States) 134:175-187.
Baker RE, Dijkhuizen PA, Van Pelt J, Verhaagen J (1998) Growth of pyramidal, but not
non-pyramidal, dendrites in long-term organotypic explants of neonatal rat neocortex
chronically exposed to neurotrophin-3. Eur J Neurosci (FRANCE) 10:1037-1044.
Bentley CA, Lee KF (2000) P75 is important for axon growth and schwann cell
migration during development. J Neurosci (UNITED STATES) 20:7706-7715.
Bhattacharjee A, Kaczmarek LK (2005) For K+ channels, na+ is the new Ca2+. Trends
Neurosci (England) 28:422-428.
2-57
Bibel M, Barde YA (2000) Neurotrophins: Key regulators of cell fate and cell shape in
the vertebrate nervous system. Genes Dev (UNITED STATES) 14:2919-2937.
Bloch RJ (1986) Loss of acetylcholine receptor clusters induced by treatment of cultured
rat myotubes with carbachol. J Neurosci (UNITED STATES) 6:691-700.
Boonstra J, Moolenaar WH, Harrison PH, Moed P, van der Saag PT, de Laat SW (1983)
Ionic responses and growth stimulation induced by nerve growth factor and epidermal
growth factor in rat pheochromocytoma (PC12) cells. J Cell Biol (UNITED STATES)
97:92-98.
Brigadski T, Hartmann M, Lessmann V (2005) Differential vesicular targeting and time
course of synaptic secretion of the mammalian neurotrophins. J Neurosci (United States)
25:7601-7614.
Cao Z, Shafer TJ, Murray TF (2011) Mechanisms of pyrethroid insecticide-induced
stimulation of calcium influx in neocortical neurons. J Pharmacol Exp Ther (United
States) 336:197-205.
Cao Z, George J, Gerwick WH, Baden DG, Rainier JD, Murray TF (2008) Influence of
lipid-soluble gating modifier toxins on sodium influx in neocortical neurons. J Pharmacol
Exp Ther (United States) 326:604-613.
Catterall WA (1986) Molecular properties of voltage-sensitive sodium channels. Annu
Rev Biochem (UNITED STATES) 55:953-985.
Catterall WA (1980) Neurotoxins that act on voltage-sensitive sodium channels in
excitable membranes. Annu Rev Pharmacol Toxicol (UNITED STATES) 20:15-43.
2-58
Catterall WA (1975a) Activation of the action potential na+ ionophore of cultured
neuroblastoma cells by veratridine and batrachotoxin. J Biol Chem (UNITED STATES)
250:4053-4059.
Catterall WA (1975b) Cooperative activation of action potential na+ ionophore by
neurotoxins. Proc Natl Acad Sci U S A (UNITED STATES) 72:1782-1786.
Catterall WA, Beneski DA (1980) Interaction of polypeptide neurotoxins with a receptor
site associated with voltage-sensitive sodium channels. J Supramol Struct (UNITED
STATES) 14:295-303.
Catterall WA, Schmidt JW, Messner DJ, Feller DJ (1986) Structure and biosynthesis of
neuronal sodium channels. Ann N Y Acad Sci (UNITED STATES) 479:186-203.
Chen ZY, Ieraci A, Teng H, Dall H, Meng CX, Herrera DG, Nykjaer A, Hempstead BL,
Lee FS (2005) Sortilin controls intracellular sorting of brain-derived neurotrophic factor
to the regulated secretory pathway. J Neurosci (United States) 25:6156-6166.
Chuang HH, Prescott ED, Kong H, Shields S, Jordt SE, Basbaum AI, Chao MV, Julius D
(2001) Bradykinin and nerve growth factor release the capsaicin receptor from
PtdIns(4,5)P2-mediated inhibition. Nature (England) 411:957-962.
Clary DO, Reichardt LF (1994) An alternatively spliced form of the nerve growth factor
receptor TrkA confers an enhanced response to neurotrophin 3. Proc Natl Acad Sci U S A
(UNITED STATES) 91:11133-11137.
Cook DI, Dinudom A, Komwatana P, Kumar S, Young JA (2002) Patch-clamp studies on
epithelial sodium channels in salivary duct cells. Cell Biochem Biophys (United States)
36:105-113.
2-59
Corbit KC, Foster DA, Rosner MR (1999) Protein kinase cdelta mediates neurogenic but
not mitogenic activation of mitogen-activated protein kinase in neuronal cells. Mol Cell
Biol (UNITED STATES) 19:4209-4218.
Dalva MB, Takasu MA, Lin MZ, Shamah SM, Hu L, Gale NW, Greenberg ME (2000)
EphB receptors interact with NMDA receptors and regulate excitatory synapse formation.
Cell (UNITED STATES) 103:945-956.
DeCaen PG, Yarov-Yarovoy V, Scheuer T, Catterall WA (2011) Gating charge
interactions with the S1 segment during activation of a na+ channel voltage sensor. Proc
Natl Acad Sci U S A (United States) 108:18825-18830.
DeCaen PG, Yarov-Yarovoy V, Sharp EM, Scheuer T, Catterall WA (2009) Sequential
formation of ion pairs during activation of a sodium channel voltage sensor. Proc Natl
Acad Sci U S A (United States) 106:22498-22503.
DeCaen PG, Yarov-Yarovoy V, Zhao Y, Scheuer T, Catterall WA (2008) Disulfide
locking a sodium channel voltage sensor reveals ion pair formation during activation.
Proc Natl Acad Sci U S A (United States) 105:15142-15147.
Dib-Hajj SD, Tyrrell L, Black JA, Waxman SG (1998) NaN, a novel voltage-gated na
channel, is expressed preferentially in peripheral sensory neurons and down-regulated
after axotomy. Proc Natl Acad Sci U S A (UNITED STATES) 95:8963-8968.
Dinudom A, Harvey KF, Komwatana P, Jolliffe CN, Young JA, Kumar S, Cook DI
(2001) Roles of the C termini of alpha -, beta -, and gamma -subunits of epithelial na+
channels (ENaC) in regulating ENaC and mediating its inhibition by cytosolic na+. J Biol
Chem (United States) 276:13744-13749.
2-60
Dryer SE (2003) Molecular identification of the na+-activated K+ channel. Neuron
(United States) 37:727-728.
Egan MF, Kojima M, Callicott JH, Goldberg TE, Kolachana BS, Bertolino A, Zaitsev E,
Gold B, Goldman D, Dean M, Lu B, Weinberger DR (2003) The BDNF val66met
polymorphism affects activity-dependent secretion of BDNF and human memory and
hippocampal function. Cell (United States) 112:257-269.
Eide FF, Vining ER, Eide BL, Zang K, Wang XY, Reichardt LF (1996) Naturally
occurring truncated trkB receptors have dominant inhibitory effects on brain-derived
neurotrophic factor signaling. J Neurosci (UNITED STATES) 16:3123-3129.
Ernfors P, Wetmore C, Olson L, Persson H (1990a) Identification of cells in rat brain and
peripheral tissues expressing mRNA for members of the nerve growth factor family.
Neuron (UNITED STATES) 5:511-526.
Ernfors P, Ibanez CF, Ebendal T, Olson L, Persson H (1990b) Molecular cloning and
neurotrophic activities of a protein with structural similarities to nerve growth factor:
Developmental and topographical expression in the brain. Proc Natl Acad Sci U S A
(UNITED STATES) 87:5454-5458.
Frade JM, Barde YA (1998) Nerve growth factor: Two receptors, multiple functions.
Bioessays (ENGLAND) 20:137-145.
George J, Dravid SM, Prakash A, Xie J, Peterson J, Jabba SV, Baden DG, Murray TF
(2009) Sodium channel activation augments NMDA receptor function and promotes
neurite outgrowth in immature cerebrocortical neurons. J Neurosci (United States)
29:3288-3301.
2-61
Ghosh A, Greenberg ME (1995) Calcium signaling in neurons: Molecular mechanisms
and cellular consequences. Science (UNITED STATES) 268:239-247.
Grunwald IC, Korte M, Wolfer D, Wilkinson GA, Unsicker K, Lipp HP, Bonhoeffer T,
Klein R (2001) Kinase-independent requirement of EphB2 receptors in hippocampal
synaptic plasticity. Neuron (United States) 32:1027-1040.
Guiton M, Gunn-Moore FJ, Glass DJ, Geis DR, Yancopoulos GD, Tavare JM (1995)
Naturally occurring tyrosine kinase inserts block high affinity binding of phospholipase C
gamma and shc to TrkC and neurotrophin-3 signaling. J Biol Chem (UNITED STATES)
270:20384-20390.
Guy HR, Seetharamulu P (1986) Molecular model of the action potential sodium channel.
Proc Natl Acad Sci U S A (UNITED STATES) 83:508-512.
Harrison SM, Jones ME, Uecker S, Albers KM, Kudrycki KE, Davis BM (2000) Levels
of nerve growth factor and neurotrophin-3 are affected differentially by the presence of
p75 in sympathetic neurons in vivo. J Comp Neurol (UNITED STATES) 424:99-110.
Hartshorne RP, Catterall WA (1981) Purification of the saxitoxin receptor of the sodium
channel from rat brain. Proc Natl Acad Sci U S A (UNITED STATES) 78:4620-4624.
Heinemann SH, Terlau H, Stuhmer W, Imoto K, Numa S (1992) Calcium channel
characteristics conferred on the sodium channel by single mutations. Nature
(ENGLAND) 356:441-443.
Henderson JT, Georgiou J, Jia Z, Robertson J, Elowe S, Roder JC, Pawson T (2001) The
receptor tyrosine kinase EphB2 regulates NMDA-dependent synaptic function. Neuron
(United States) 32:1041-1056.
2-62
HODGKIN AL, HUXLEY AF (1952a) Currents carried by sodium and potassium ions
through the membrane of the giant axon of loligo. J Physiol (Not Available) 116:449-
472.
HODGKIN AL, HUXLEY AF (1952b) The dual effect of membrane potential on sodium
conductance in the giant axon of loligo. J Physiol (Not Available) 116:497-506.
HODGKIN AL, HUXLEY AF (1952c) Movement of sodium and potassium ions during
nervous activity. Cold Spring Harb Symp Quant Biol (Not Available) 17:43-52.
Huang EJ, Reichardt LF (2003a) Trk receptors: Roles in neuronal signal transduction.
Annu Rev Biochem (United States) 72:609-642.
Huang EJ, Reichardt LF (2003b) Trk receptors: Roles in neuronal signal transduction.
Annu Rev Biochem (United States) 72:609-642.
Isom LL, Ragsdale DS, De Jongh KS, Westenbroek RE, Reber BF, Scheuer T, Catterall
WA (1995) Structure and function of the beta 2 subunit of brain sodium channels, a
transmembrane glycoprotein with a CAM motif. Cell (UNITED STATES) 83:433-442.
Isom LL, De Jongh KS, Patton DE, Reber BF, Offord J, Charbonneau H, Walsh K,
Goldin AL, Catterall WA (1992) Primary structure and functional expression of the beta
1 subunit of the rat brain sodium channel. Science (UNITED STATES) 256:839-842.
Jabba SV, Prakash A, Dravid SM, Gerwick WH, Murray TF (2010) Antillatoxin, a novel
lipopeptide, enhances neurite outgrowth in immature cerebrocortical neurons through
activation of voltage-gated sodium channels. J Pharmacol Exp Ther (United States)
332:698-709.
Jaworski J, Sheng M (2006) The growing role of mTOR in neuronal development and
plasticity. Mol Neurobiol (United States) 34:205-219.
2-63
Kaplan DR, Hempstead BL, Martin-Zanca D, Chao MV, Parada LF (1991) The trk proto-
oncogene product: A signal transducing receptor for nerve growth factor. Science
(UNITED STATES) 252:554-558.
Kelleher RJ,3rd, Govindarajan A, Jung HY, Kang H, Tonegawa S (2004) Translational
control by MAPK signaling in long-term synaptic plasticity and memory. Cell (United
States) 116:467-479.
Khodorov BI (1985) Batrachotoxin as a tool to study voltage-sensitive sodium channels
of excitable membranes. Prog Biophys Mol Biol (ENGLAND) 45:57-148.
Kidane AH, Heinrich G, Dirks RP, de Ruyck BA, Lubsen NH, Roubos EW, Jenks BG
(2009) Differential neuroendocrine expression of multiple brain-derived neurotrophic
factor transcripts. Endocrinology (United States) 150:1361-1368.
Klann E, Dever TE (2004) Biochemical mechanisms for translational regulation in
synaptic plasticity. Nat Rev Neurosci (England) 5:931-942.
Klein R, Jing SQ, Nanduri V, O'Rourke E, Barbacid M (1991a) The trk proto-oncogene
encodes a receptor for nerve growth factor. Cell (UNITED STATES) 65:189-197.
Klein R, Nanduri V, Jing SA, Lamballe F, Tapley P, Bryant S, Cordon-Cardo C, Jones
KR, Reichardt LF, Barbacid M (1991b) The trkB tyrosine protein kinase is a receptor for
brain-derived neurotrophic factor and neurotrophin-3. Cell (UNITED STATES) 66:395-
403.
Kolarow R, Brigadski T, Lessmann V (2007) Postsynaptic secretion of BDNF and NT-3
from hippocampal neurons depends on calcium calmodulin kinase II signaling and
proceeds via delayed fusion pore opening. J Neurosci (United States) 27:10350-10364.
2-64
Komwatana P, Dinudom A, Young JA, Cook DI (1996) Cytosolic na+ controls and
epithelial na+ channel via the go guanine nucleotide-binding regulatory protein. Proc Natl
Acad Sci U S A (UNITED STATES) 93:8107-8111.
Kontis KJ, Goldin AL (1997) Sodium channel inactivation is altered by substitution of
voltage sensor positive charges. J Gen Physiol (UNITED STATES) 110:403-413.
Kontis KJ, Rounaghi A, Goldin AL (1997) Sodium channel activation gating is affected
by substitutions of voltage sensor positive charges in all four domains. J Gen Physiol
(UNITED STATES) 110:391-401.
Lee CC, Huang CC, Wu MY, Hsu KS (2005) Insulin stimulates postsynaptic density-95
protein translation via the phosphoinositide 3-kinase-akt-mammalian target of rapamycin
signaling pathway. J Biol Chem (United States) 280:18543-18550.
Lee KC, Loh TP (2006) Total synthesis of antillatoxin. Chem Commun (Camb)
(England) (40):4209-4211.
Lee R, Kermani P, Teng KK, Hempstead BL (2001) Regulation of cell survival by
secreted proneurotrophins. Science (United States) 294:1945-1948.
Lessmann V, Gottmann K, Malcangio M (2003) Neurotrophin secretion: Current facts
and future prospects. Prog Neurobiol (England) 69:341-374.
Li WI, Marquez BL, Okino T, Yokokawa F, Shioiri T, Gerwick WH, Murray TF (2004)
Characterization of the preferred stereochemistry for the neuropharmacologic actions of
antillatoxin. J Nat Prod (United States) 67:559-568.
Li WI, Berman FW, Okino T, Yokokawa F, Shioiri T, Gerwick WH, Murray TF (2001)
Antillatoxin is a marine cyanobacterial toxin that potently activates voltage-gated sodium
channels. Proc Natl Acad Sci U S A (United States) 98:7599-7604.
2-65
Li Y, Jia YC, Cui K, Li N, Zheng ZY, Wang YZ, Yuan XB (2005) Essential role of
TRPC channels in the guidance of nerve growth cones by brain-derived neurotrophic
factor. Nature (England) 434:894-898.
Liepinsh E, Ilag LL, Otting G, Ibanez CF (1997) NMR structure of the death domain of
the p75 neurotrophin receptor. EMBO J (ENGLAND) 16:4999-5005.
Linden DJ, Smeyne M, Connor JA (1993) Induction of cerebellar long-term depression in
culture requires postsynaptic action of sodium ions. Neuron (UNITED STATES)
11:1093-1100.
Lindholm D, Castren E, Hengerer B, Zafra F, Berninger B, Thoenen H (1992)
Differential regulation of nerve growth factor (NGF) synthesis in neurons and astrocytes
by glucocorticoid hormones. Eur J Neurosci 4:404-410.
Linford NJ, Cantrell AR, Qu Y, Scheuer T, Catterall WA (1998) Interaction of
batrachotoxin with the local anesthetic receptor site in transmembrane segment IVS6 of
the voltage-gated sodium channel. Proc Natl Acad Sci U S A (UNITED STATES)
95:13947-13952.
Majdan M, Shatz CJ (2006) Effects of visual experience on activity-dependent gene
regulation in cortex. Nat Neurosci (United States) 9:650-659.
Makkerh JP, Ceni C, Auld DS, Vaillancourt F, Dorval G, Barker PA (2005) p75
neurotrophin receptor reduces ligand-induced trk receptor ubiquitination and delays trk
receptor internalization and degradation. EMBO Rep (England) 6:936-941.
Mayer ML, Westbrook GL, Guthrie PB (1984) Voltage-dependent block by Mg2+ of
NMDA responses in spinal cord neurones. Nature (ENGLAND) 309:261-263.
2-66
McAllister AK, Katz LC, Lo DC (1996) Neurotrophin regulation of cortical dendritic
growth requires activity. Neuron (UNITED STATES) 17:1057-1064.
Meakin SO, Gryz EA, MacDonald JI (1997) A kinase insert isoform of rat TrkA supports
nerve growth factor-dependent cell survival but not neurite outgrowth. J Neurochem
(UNITED STATES) 69:954-967.
Meakin SO, Suter U, Drinkwater CC, Welcher AA, Shooter EM (1992) The rat trk
protooncogene product exhibits properties characteristic of the slow nerve growth factor
receptor. Proc Natl Acad Sci U S A (UNITED STATES) 89:2374-2378.
Miller FD, Kaplan DR (2003) Signaling mechanisms underlying dendrite formation. Curr
Opin Neurobiol (England) 13:391-398.
Monyer H, Burnashev N, Laurie DJ, Sakmann B, Seeburg PH (1994) Developmental and
regional expression in the rat brain and functional properties of four NMDA receptors.
Neuron (UNITED STATES) 12:529-540.
Monyer H, Sprengel R, Schoepfer R, Herb A, Higuchi M, Lomeli H, Burnashev N,
Sakmann B, Seeburg PH (1992) Heteromeric NMDA receptors: Molecular and functional
distinction of subtypes. Science (UNITED STATES) 256:1217-1221.
Moolenaar WH, Defize LH, De Laat SW (1986) Ionic signalling by growth factor
receptors. J Exp Biol (ENGLAND) 124:359-373.
Moolenaar WH, Tsien RY, van der Saag PT, de Laat SW (1983) Na+/H+ exchange and
cytoplasmic pH in the action of growth factors in human fibroblasts. Nature
(ENGLAND) 304:645-648.
Mori H, Mishina M (1995) Structure and function of the NMDA receptor channel.
Neuropharmacology (ENGLAND) 34:1219-1237.
2-67
Moriyoshi K, Masu M, Ishii T, Shigemoto R, Mizuno N, Nakanishi S (1991) Molecular
cloning and characterization of the rat NMDA receptor. Nature (ENGLAND) 354:31-37.
Mowla SJ, Pareek S, Farhadi HF, Petrecca K, Fawcett JP, Seidah NG, Morris SJ, Sossin
WS, Murphy RA (1999) Differential sorting of nerve growth factor and brain-derived
neurotrophic factor in hippocampal neurons. J Neurosci (UNITED STATES) 19:2069-
2080.
Nakata H, Nakamura S (2007) Brain-derived neurotrophic factor regulates AMPA
receptor trafficking to post-synaptic densities via IP3R and TRPC calcium signaling.
FEBS Lett (Netherlands) 581:2047-2054.
Nicholls D, Attwell D (1990) The release and uptake of excitatory amino acids. Trends
Pharmacol Sci (ENGLAND) 11:462-468.
Noda M, Suzuki H, Numa S, Stuhmer W (1989) A single point mutation confers
tetrodotoxin and saxitoxin insensitivity on the sodium channel II. FEBS Lett
(NETHERLANDS) 259:213-216.
Noda M, Ikeda T, Suzuki H, Takeshima H, Takahashi T, Kuno M, Numa S (1986)
Expression of functional sodium channels from cloned cDNA. Nature (ENGLAND)
322:826-828.
Orjala J, Nagle D, Hsu V, Gerwick WH (1995) Antillatoxin: An exceptionally
ichthyotoxic cyclic lipopeptide from the tropical cyanobacterium lyngbya majuscula. J
Am Chem Soc 117:8281-8282.
Pang PT, Lu B (2004) Regulation of late-phase LTP and long-term memory in normal
and aging hippocampus: Role of secreted proteins tPA and BDNF. Ageing Res Rev
(England) 3:407-430.
2-68
Paoletti P, Neyton J (2007) NMDA receptor subunits: Function and pharmacology. Curr
Opin Pharmacol (England) 7:39-47.
Patino GA, Brackenbury WJ, Bao Y, Lopez-Santiago LF, O'Malley HA, Chen C,
Calhoun JD, Lafreniere RG, Cossette P, Rouleau GA, Isom LL (2011) Voltage-gated na+
channel beta1B: A secreted cell adhesion molecule involved in human epilepsy. J
Neurosci (United States) 31:14577-14591.
Payandeh J, Scheuer T, Zheng N, Catterall WA (2011) The crystal structure of a voltage-
gated sodium channel. Nature (England) 475:353-358.
Pert CB, Snowman AM, Snyder SH (1974a) Localization of opiate receptor binding in
synaptic membranes of rat brain. Brain Res (NETHERLANDS) 70:184-188.
Pert CB, Aposhian D, Snyder SH (1974b) Phylogenetic distribution of opiate receptor
binding. Brain Res (NETHERLANDS) 75:356-361.
Prescott ED, Julius D (2003) A modular PIP2 binding site as a determinant of capsaicin
receptor sensitivity. Science (United States) 300:1284-1288.
Puttfarcken P, Werling LL, Brown SR, Cote TE, Cox BM (1986) Sodium regulation of
agonist binding at opioid receptors. I. effects of sodium replacement on binding at mu-
and delta-type receptors in 7315c and NG108-15 cells and cell membranes. Mol
Pharmacol (UNITED STATES) 30:81-89.
Rattiner LM, Davis M, Ressler KJ (2005) Brain-derived neurotrophic factor in amygdala-
dependent learning. Neuroscientist (United States) 11:323-333.
Read DE, Gorman AM (2009) Involvement of akt in neurite outgrowth. Cell Mol Life Sci
(Switzerland) 66:2975-2984.
2-69
Redmond L, Kashani AH, Ghosh A (2002) Calcium regulation of dendritic growth via
CaM kinase IV and CREB-mediated transcription. Neuron (United States) 34:999-1010.
Reichardt LF (2006) Neurotrophin-regulated signalling pathways. Philos Trans R Soc
Lond B Biol Sci (England) 361:1545-1564.
Rodriguez-Tebar A, Dechant G, Barde YA (1990) Binding of brain-derived neurotrophic
factor to the nerve growth factor receptor. Neuron (UNITED STATES) 4:487-492.
Rose CR, Konnerth A (2001) NMDA receptor-mediated na+ signals in spines and
dendrites. J Neurosci (United States) 21:4207-4214.
Rose CR, Kovalchuk Y, Eilers J, Konnerth A (1999) Two-photon na+ imaging in spines
and fine dendrites of central neurons. Pflugers Arch (GERMANY) 439:201-207.
Rose CR, Blum R, Pichler B, Lepier A, Kafitz KW, Konnerth A (2003) Truncated TrkB-
T1 mediates neurotrophin-evoked calcium signalling in glia cells. Nature (England)
426:74-78.
Salter MW, Kalia LV (2004) Src kinases: A hub for NMDA receptor regulation. Nat Rev
Neurosci (England) 5:317-328.
Saneyoshi T, Wayman G, Fortin D, Davare M, Hoshi N, Nozaki N, Natsume T,
Soderling TR (2008) Activity-dependent synaptogenesis: Regulation by a CaM-kinase
kinase/CaM-kinase I/betaPIX signaling complex. Neuron (United States) 57:94-107.
Sarbassov DD, Ali SM, Sabatini DM (2005a) Growing roles for the mTOR pathway.
Curr Opin Cell Biol (United States) 17:596-603.
Sarbassov DD, Guertin DA, Ali SM, Sabatini DM (2005b) Phosphorylation and
regulation of Akt/PKB by the rictor-mTOR complex. Science (United States) 307:1098-
1101.
2-70
Schlief T, Schonherr R, Imoto K, Heinemann SH (1996) Pore properties of rat brain II
sodium channels mutated in the selectivity filter domain. Eur Biophys J (GERMANY)
25:75-91.
Seidah NG, Benjannet S, Pareek S, Chretien M, Murphy RA (1996) Cellular processing
of the neurotrophin precursors of NT3 and BDNF by the mammalian proprotein
convertases. FEBS Lett (NETHERLANDS) 379:247-250.
Shaywitz AJ, Greenberg ME (1999) CREB: A stimulus-induced transcription factor
activated by a diverse array of extracellular signals. Annu Rev Biochem (UNITED
STATES) 68:821-861.
Strohmaier C, Carter BD, Urfer R, Barde YA, Dechant G (1996) A splice variant of the
neurotrophin receptor trkB with increased specificity for brain-derived neurotrophic
factor. EMBO J (ENGLAND) 15:3332-3337.
Takasu MA, Dalva MB, Zigmond RE, Greenberg ME (2002) Modulation of NMDA
receptor-dependent calcium influx and gene expression through EphB receptors. Science
(United States) 295:491-495.
Tanaka N, Fujisawa T, Daimon T, Fujiwara K, Yamamoto M, Abe T (1999) The effect of
electrolyzed strong acid aqueous solution on hemodialysis equipment. Artif Organs
(UNITED STATES) 23:1055-1062.
Terlau H, Heinemann SH, Stuhmer W, Pusch M, Conti F, Imoto K, Numa S (1991)
Mapping the site of block by tetrodotoxin and saxitoxin of sodium channel II. FEBS Lett
(NETHERLANDS) 293:93-96.
2-71
Trainer VL, Brown GB, Catterall WA (1996) Site of covalent labeling by a photoreactive
batrachotoxin derivative near transmembrane segment IS6 of the sodium channel alpha
subunit. J Biol Chem (UNITED STATES) 271:11261-11267.
Traynelis SF, Wollmuth LP, McBain CJ, Menniti FS, Vance KM, Ogden KK, Hansen
KB, Yuan H, Myers SJ, Dingledine R (2010) Glutamate receptor ion channels: Structure,
regulation, and function. Pharmacol Rev (United States) 62:405-496.
Ulbricht W (1969) The effect of veratridine on excitable membranes of nerve and muscle.
Ergeb Physiol (GERMANY, WEST) 61:18-71.
Ultanir SK, Kim JE, Hall BJ, Deerinck T, Ellisman M, Ghosh A (2007) Regulation of
spine morphology and spine density by NMDA receptor signaling in vivo. Proc Natl
Acad Sci U S A (United States) 104:19553-19558.
Wang J, Yarov-Yarovoy V, Kahn R, Gordon D, Gurevitz M, Scheuer T, Catterall WA
(2011) Mapping the receptor site for alpha-scorpion toxins on a na+ channel voltage
sensor. Proc Natl Acad Sci U S A (United States) 108:15426-15431.
Watanabe M, Inoue Y, Sakimura K, Mishina M (1992) Developmental changes in
distribution of NMDA receptor channel subunit mRNAs. Neuroreport (ENGLAND)
3:1138-1140.
Wayman GA, Impey S, Marks D, Saneyoshi T, Grant WF, Derkach V, Soderling TR
(2006) Activity-dependent dendritic arborization mediated by CaM-kinase I activation
and enhanced CREB-dependent transcription of wnt-2. Neuron (United States) 50:897-
909.
2-72
Werling LL, Brown SR, Puttfarcken P, Cox BM (1986) Sodium regulation of agonist
binding at opioid receptors. II. effects of sodium replacement on opioid binding in guinea
pig cortical membranes. Mol Pharmacol (UNITED STATES) 30:90-95.
West AE, Griffith EC, Greenberg ME (2002) Regulation of transcription factors by
neuronal activity. Nat Rev Neurosci (England) 3:921-931.
West JW, Patton DE, Scheuer T, Wang Y, Goldin AL, Catterall WA (1992) A cluster of
hydrophobic amino acid residues required for fast na(+)-channel inactivation. Proc Natl
Acad Sci U S A (UNITED STATES) 89:10910-10914.
Wong RO, Ghosh A (2002) Activity-dependent regulation of dendritic growth and
patterning. Nat Rev Neurosci (England) 3:803-812.
Yamane M, Yamashita N, Yamamoto H, Iizuka A, Shouji M, Usui H, Goshima Y (2012)
Semaphorin3A facilitates axonal transport through a local calcium signaling and
tetrodotoxin-sensitive voltage-gated sodium channels. Biochem Biophys Res Commun
(United States) 422:333-338.
Yarov-Yarovoy V, DeCaen PG, Westenbroek RE, Pan CY, Scheuer T, Baker D, Catterall
WA (2012) Structural basis for gating charge movement in the voltage sensor of a
sodium channel. Proc Natl Acad Sci U S A (United States) 109:E93-102.
Yoshii A, Constantine-Paton M (2007) BDNF induces transport of PSD-95 to dendrites
through PI3K-AKT signaling after NMDA receptor activation. Nat Neurosci (United
States) 10:702-711.
Yu XM, Salter MW (1998) Gain control of NMDA-receptor currents by intracellular
sodium. Nature (ENGLAND) 396:469-474.
2-73
Zhang JZ, Yarov-Yarovoy V, Scheuer T, Karbat I, Cohen L, Gordon D, Gurevitz M,
Catterall WA (2011) Structure-function map of the receptor site for beta-scorpion toxins
in domain II of voltage-gated sodium channels. J Biol Chem (United States) 286:33641-
33651.
3-74
3 CHAPTER 3 - Antillatoxin, a Novel Lipopeptide, Enhances
Neurite Outgrowth in Immature Cerebrocortical Neurons
Through Activation of Voltage–Gated Sodium Channels
3-75
3.1 Abstract
Antillatoxin (ATX) is a structurally novel lipopeptide that activates voltage-gated
sodium channels (VGSC) leading to sodium influx in cerebellar granule neurons and DIV
8-9 cerebrocortical neurons (Cao et al., 2008; Li et al., 2001). The precise recognition site
for ATX on the VGSC, however, remains to be defined. Inasmuch as elevation of
cytoplasmic sodium may increase N-methyl-D-aspartate receptor (NMDAR) mediated
Ca2+ influx, intracellular sodium [Na+]i may function as a signaling molecule. Here we
hypothesized that ATX may enhance neurite outgrowth in cerebrocortical neurons by
elevating [Na+]i and augumenting NMDAR function. ATX concentrations of 30-100 nM
robustly stimulated neurite outgrowth and this enhancement was sensitive to the VGSC
antagonist, tetrodotoxin. To unambiguously demonstrate the enhancement of NMDA
receptor function by ATX, we recorded single-channel currents from cell-attached
patches. ATX was found to increase the open probability of NMDA receptors. Na+
dependent upregulation of NMDAR function has been shown to be regulated by Src
family kinase (SFK) (Yu and Salter, 1998). The Src kinase inhibitor PP2 abrogated
ATX-enhanced neurite outgrowth suggesting a SFK involvement in this response. ATX-
enhanced neurite outgrowth was also inhibited by the NMDAR antagonist, MK-801, and
the calmodulin dependent kinase kinase (CaMKK) inhibitor, STO-609, demonstrating the
requirement for NMDAR activation with subsequent downstream engagement of the Ca2+
dependent CaMKK pathway. These results with the structurally and mechanistically
novel natural product, ATX, confirm and generalize our earlier results with a neurotoxin
site 5 ligand. These data suggest that VGSC activators may represent a novel
3-76
pharmacological strategy to regulate neuronal plasticity through NMDAR-dependent
mechanisms.
Antillatoxin (ATX) is a structurally novel lipopeptide with an exceptionally high
degree of methylation unlike any known natural product (Lee and Loh, 2006). Isolated
from the cyanobacterium Lyngbya majuscula, this compound is also distinguished by
multiple stereocenters (Orjala et al., 1995). The essential role of the asymmetric carbon
atoms in ATX is reflected in the stereoselective effects of ATX enantiomers (Li et al.,
2004). ATX is considered to be the second most potent ichthyotoxic compound obtained
from marine sources following only brevetoxin (PbTx-1) (Perez-Otano and Ehlers, 2005).
Exposure to L. majuscula blooms are associated with adverse human health effects,
including respiratory irritation, eye inflammation and severe contact dermatitis. Previous
work has further demonstrated that ATX is a potent activator of voltage-gated sodium
channels (VGSCs) that elevates intracellular Na+ concentration ([Na+]i) in intact neurons
(Cao et al., 2008; Li et al., 2001). ATX has moreover been shown to be neurotoxic in
cerebellar granule cells (CGCs) through an indirect activation of N-methyl-D-aspartate
receptors (NMDARs) as a consequence of glutamate release (Li et al., 2001; Li et al.,
2004).
Regulation of [Na+]i plays a critical role in the nervous system, not only because
Na+ influx through VGSCs is responsible for the initiation (the rising phase) and
propagation of action potentials, but also because various neuronal cell functions such as
intracellular pH, Ca2+ homeostasis and reuptake of neurotransmitters are dependent on
the Na+ gradient. Previous studies have further indicated that intracellular Na+ can also
3-77
act as a signaling molecule to modulate cell functions including, but not restricted to, cell
proliferation, ion channel permeability, G-protein function and opioid ligand-receptor
interactions (Yu, 2006). Recent studies have moreover demonstrated that neuronal
activity mediated increases in [Na+]i in structures including soma, dendrites and spines
may act as a signaling molecule and contribute to activity-dependent synaptic plasticity
(Rose and Konnerth, 2001). In cerebellar Purkinje neurons, AMPA receptor mediated
Na+ influx was shown to be required for induction of LTD (Linden et al., 1993). In both
hippocampal and immature cerebrocortical neurons an elevation in intracellular Na+ was
found to increase NMDAR-mediated whole-cell currents and NMDAR single channel
activity by increasing both channel open probability and mean open time (George et al.,
2009; Yu and Salter, 1998). This [Na+]i mediated upregulation of NMDAR function has
been shown to require Src kinase activation (George et al., 2009; Yu and Salter, 1998).
Src family kinases act as a crucial point of convergence for signaling pathways that
enhance NMDAR activity, and, by upregulating the function of NMDARs, Src gates the
production of NMDAR-dependent synaptic potentiation and plasticity (Salter and Kalia,
2004).
Neuronal activity has a major role in the development of dendritic complexity and
neuronal circuits. The mechanisms by which neuronal activity translate into
morphological changes are complex. Numerous studies have shown that activity-
dependent neuronal development involves various calcium influx pathways mediated by
ionotropic glutamate receptors (mainly NMDAR) and voltage-gated Ca2+ channels
(VGCCs) (Ghosh and Greenberg, 1995; West et al., 2002). Intracellular calcium acts as a
signaling molecule largely through the binding to calmodulin, a calcium binding protein
3-78
that engages downstream Ca2+/calmodulin dependent protein kinase (CaMK) and
mitogen activated protein kinase (MAPK) signaling pathways (Ghosh and Greenberg,
1995; West et al., 2002). CaMK kinase (CaMKK) has been demonstrated to be an
upstream regulator of both CaMK- and MAPK-signaling pathways. Moreover, previous
studies have demonstrated that activity-dependent neurite outgrowth (Wayman et al.,
2006) and synaptogenesis (Saneyoshi et al., 2008) are regulated by NMDAR-dependent
CaMKK/calmodulin kinase I-signaling cascades. NMDARs therefore play a critical role
in activity-dependent development and plasticity (Ghosh and Greenberg, 1995), dendritic
arborization (Miller and Kaplan, 2003; Wayman et al., 2006; Wong and Ghosh, 2002),
spine morphogenesis (Ultanir et al., 2007) and synapse formation (Saneyoshi et al., 2008)
by stimulating these calcium-dependent signaling pathways.
Inasmuch as neuronal activity induced increments in cytoplasmic sodium may
augment NMDAR mediated currents, we reasoned that intracellular Na+ may function as
a signaling molecule and regulate neuritogenesis in immature cerebrocortical neurons.
We have recently demonstrated that brevetoxin (PbTx-2), a VGSC activator, enhanced
NMDAR function and augumented neurite outgrowth (George et al., 2009). In the
present study we extend our earlier work to demonstrate that these pharmacologic actions
of the neurotoxin site 5 ligand brevetoxin generalize to the structurally and
mechanistically novel VGSC activator, ATX. We found that ATX promoted
neuritogenisis by elevating [Na+]i which in turn augumented NMDAR function leading to
Ca2+ influx and engagement of a CaMKK pathway. These data provide further support
for the hypothesis that sodium channel activators appear capable of mimicking activity-
3-79
dependent neuronal development through potentiation of NMDAR signaling pathways
that influence neuronal plasticity.
3.2 Experimental Procedures
Cerebrocortical neuron culture- Primary cultures of cerebrocortical neurons were
harvested from Swiss–Webster mice on embryonic day 16 and cultured as described
previously (Cao et al., 2008). Cells were plated onto poly-L-lysine -coated (Sigma) 96-
well (9 mm), clear-bottomed, black-well culture plates (Costar) at a density of 1.8 x 106
cells per mL (150 µL per well) , 24-well (15.6 mm) culture plates at a density of 0.05 x
106 cells per ml (0.5 mL per well), or 6-well (35 mm) culture dishes at a density of 2.25 x
106 cell per ml (2 ml per well), respectively, and incubated at 37°C in a 5% CO2 and 95%
humidity atmosphere. All animal use protocols were approved by the Creighton
University Institutional Animal Care and Use Committee.
Immunocytochemistry and determination of total neurite length- Cells were plated
on poly-lysine coated 12 or 15 mm glass coverslips (Fisher Scientific), and placed inside
of 24-well culture plates at a low density of 0.05 x 106 cells per ml (0.5 mL per well).
To assess the influence of ATX on neuritogenesis, primary cultures of immature
cerebrocortical neurons were exposed to various concentrations of ATX ranging from 1-
1,000 nM for 24 h beginning three hours post plating and total neurite outgrowth was
measured. In some experiments these concentrations of ATX were coincubated with
tetrodotoxin (TTX) (1 µM) (Biomol), MK-801 (1 µM) (Sigma), nifedipine (1 µM)
(Sigma), 1,8-naphthoylene benzimidazole-3-carboxylic acid (STO-609) (2.6 µM)
(Calbiochem), 4-Amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolo[3, 4-d] pyrimidine (PP2)
3-80
or 4-amino-7-phenylpyrazol [3, 4-d] pyrimidine (PP3) (Calbiochem). At 24 h post
plating, cultures were fixed at room temperature for 20 min using 4% paraformaldehyde
in PBS. Post-fixation, neurons were blocked and permeabilized by incubating for 30 min
with PBS containing 2% fetal bovine serum (Atlanta Biologicals) and 0.15% Triton X-
100 (Sigma). The coverslips were incubated overnight at 4°C with protein gene product
9.5 (anti-PGP 9.5) primary antibody (AbD SeroTec). After washing 3X in blocking
buffer, coverslips were incubated with a secondary antibody [FITC (anti-rabbit IgG)]
(Jackson ImmunoResearch Laboratories) for 60 min at room temperature. Coverslips
were washed and mounted on microscope slides and analyzed by fluorescence
microscopy on an Olympus IX 71 inverted microscope with a Nikon camera. Digital
images of individual neurons were captured and total neurite length quantified using
Image Pro plus (Media Cybernetics). To reduce the effect of paracrine neurotrophic
factors on neurite growth, only those neurons that were separated from surrounding cells
by approximately 150 µm were digitally acquired and analysed. At least 25 randomly
chosen neurons from different cultures were evaluated for each treatment group.
Diolistic labeling- The Helios Gene Gun System (Bio-Rad, Hercules, CA) was used to
deliver DiI coated tungsten particles (1.3 µM) (Bio-Rad) into paraformaldehyde fixed
DIV-1 cerebrocortical neurons. Diolistic bullet preparation was based on the method of
O`Brien and Lummis (O'Brien and Lummis, 2006). Briefly, 2.5-3.5 mg of DiI
(Invitrogen) was suspended in 200 µl of dichloromethane (Sigma). The dissolved dye
was added over evenly spread 35 mg tungsten particles placed on a clean glass slide, and
3-81
then allowed to dry. The dye coated particles were scraped onto another clean glass slide
and chopped to fine particles using a clean razor blade and later re-suspended in 3 ml
deionized water. This dye slurry was sonicated for 10 minutes and then vortex briefly to
form a uniform suspension. After adding 100 µl of polyvinylpyrrolidone (PVP) stock
solution (0.96 % PVP in ethanol) to the dye slurry, it was drawn into a PVP pre-coated
tefzel tubing mounted on a preparation station (Bio-Rad) using a 5-10 ml syringe. The
dye particles were allowed to settle for 20-30 minutes and then the supernatant water was
carefully withdrawn from tefzel tubing using a syringe. The tubing was rotated for 1-2
min to uniformly spread the particles. The tubing was then allowed to dry for 5 min
before cutting into bullets using a tube cutter. The DIV-1 cerebrocortical neurons grown
on cover slips were shot post-fixation (1.5 % paraformaldehyde) using DiI bullets loaded
onto a Helios gene gun at 140-160 psi of helium pressure from a distance of 2.5 cm. The
dye particles were allowed to spread across the neuronal membrane overnight and cover
slips were then mounted for imaging.
Intracellular sodium concentration ([Na+]i) measurement- [Na+]i measurement and
full in situ calibration of SBFI fluorescence ratio were performed as described previously
(Cao et al., 2008). Cells grown in 96-well plates were washed four times with Locke's
buffer (in mM: 8.6 HEPES, 5.6 KCl, 154 NaCl, 5.6 glucose, 1.0 MgCl2, 2.3 CaCl2, 0.1
glycine, pH 7.4) using an automated microplate washer (Bio-Tek Instruments). After
measuring the background fluorescence of each well, cells were incubated for 1 h at 37°C
with dye-loading buffer (100 µl/well) containing 10 µM SBFI-AM (Invitrogen) and
0.02% pluronic F-127 (Invitrogen). Cells were then washed 5X with Locke's buffer
leaving a final volume of 120 µl in each well. The plate was then transferred back to the
3-82
incubator for 15 min to allow the cells to equilibrate following washing and then placed
in a FlexStation II (Molecular Devices) chamber to detect Na+- bound SBFI emission at
505 nm (cells were excited at 340 and 380 nm). Fluorescence readings were taken once
every 5 s for 60 s to establish the baseline, and then 40 µl ATX was added to each well
from the compound plate at a rate of 26 µl/s, yielding a final volume of 160 µl/well. After
correcting for background fluorescence, SBFI fluorescence ratios (340/380) versus time
were analyzed, and time- or concentration-response graphs were generated using
GraphPad Prism (GraphPad Software).
Full in situ calibration of the SBFI fluorescence ratio was performed using
calibration media containing the following (in mM): 0.6 MgCl2, 0.5 CaCl2, 10 HEPES,
Na+ and K+ such that [Na+] plus [K+] = 130, 100 gluconate, and 30 Cl- (titrated with 10
mol/l KOH to pH 7.4). Gramicidin D (5 µM) (Na+ ionophore), monensin (10 µM)
(Na+/H+ carrier), and ouabain (100 µM) (Na+/K+-ATPase inhibitor) to equilibrate the
intracellular and extracellular sodium concentration. After five washes, Locke's buffer
was replaced by 150 µl sodium-containing calibration solution (0-130 mM). The plate
was then loaded onto the FlexStation chamber for recording of emitted fluorescence
during excitation at 340 and 380 nm. Fluorescence data were converted to a ratio
(340/380) after background correction. To convert the fluorescence ratio of emitted SBFI
signals into a [Na+]i value, the following equation was used: [Na+] = βKd [(R – Rmin)/(Rmax
– R)] (equation 1), where β is the ratio of the fluorescence of the free (unbound) dye to
bound dye at the second excitation wavelength (380 nm), Kd is the apparent dissociation
constant of SBFI for Na+, R is the background-subtracted SBFI fluorescence ratio, and
Rmin and Rmax are, respectively, the minimum and maximum fluorescence values. Data
3-83
relating [Na+]i to R were fitted by a three-parameter hyperbolic equation having the
following form: R = Rmin + [a ([Na+])/(b + [Na+])] (equation 2), where a and b are
constants and equal to Rmax – Rmin and βKd, respectively (2, 33). These data relating
[Na+]i to R (see Fig. 6B) were well described (r2 = 0.98) by Equation 2. The derived
parameters were Rmin = 1.47 ± 0.03, a = 3.541 ± 0.11, and b = 63.30 ± 4.93. The value for
Rmin obtained by this method was similar to the value of Rmin derived experimentally at
[Na+] = 0 mM. The corresponding values for Rmax and βKd were, therefore, Rmax = 5.01 ±
0.13 and βKd = 63.30 ± 4.93 mM. We compared the values of Rmax and βKd obtained from
a Hanes plot (Cao et al., 2008) to those derived from the three-parameter hyperbolic fit.
The equation was rearranged to generate a Hanes plot such that [Na+]/(R – Rmin) =
[Na+]/(Rmax – Rmin) + [βKd/(Rmax – Rmin)] (equation 3).
The plotting of [Na+]/(R – Rmin) versus [Na+]i as a Hanes function yielded a
straight line (r2 = 1) (data not shown). The slope 1/(Rmax – Rmin) of this regression
provides a means to estimate of Rmax, whereas the intercept on the abscissa is equal to –
βKd. The value for Rmin was obtained from the experimental data. The values of Rmax and
βKd calculated from Hanes plot were 4.97 ± 0.10 and 63.3 ± 1.9 mM, respectively, and
were therefore not significantly different from the values derived from the three-
parameter hyperbolic fit which were 5.01 ± 0.13 (Rmax) and 63.3 ± 4.93 mM (βKd).
Intracellular Ca2+ monitoring- DIV-1 cerebrocortical neurons grown in 96-well plates
were used for intracellular Ca2+ concentration ([Ca2+]i) measurements as described
previously (George et al., 2009). Briefly, the growth medium was removed and replaced
with dye-loading medium (100 µl per well) containing 8 µM fluo-3 AM (Invitrogen) and
3-84
0.04% pluronic acid in Locke's buffer. After 1 h of incubation in dye-loading medium, the
neurons were washed four times in fresh Locke's buffer (200 µl per well, 22°C) using an
automated microplate washer (Bio-Tek Instruments) and transferred to a FlexStation II
benchtop scanning fluorometer chamber. The final volume of Locke's buffer in each well
was 120 µl. Fluorescence measurements were performed at 37°C. The neurons were
excited at 488 nm and Ca2+-bound fluo-3 emission was recorded at 538 nm at 1.2 s
intervals. After recording baseline fluorescence for 27 s, 40 µl of a 4X concentration of
ATX in the presence or absence of either PP2, PP3, nifedipine or MK 801 were added to
the cells at a rate of 26 µl/s yielding a final volume of 160 µl/well; the fluorescence was
monitored for an additional 220–270 s. The fluo-3 fluorescence was expressed as (Fmax –
Fmin)/Fmin where Fmax is the maximum, and Fmin the minimum fluorescence measured in
each well.
Western blotting- Western blot analysis was performed using cells grown in 6-well
plates. Three hours post platting, cells exposed to 30 nM ATX for time periods ranging
from 0 to 24 h at 37ºC. At the end of each time period cultures were transferred onto an
ice slurry to terminate drug exposure and washed 3X with ice-cold PBS. Cells were
lysed using ice-cold lysis buffer (50 mM Tris, 50 mM NaCl, 2 mM EDTA, 2 mM EGTA,
1% NP-40, 0.1% SDS, 2.5 mM sodium pyrophosphate, and 1 mM sodium
orthovanadate). Phenylmethylsulfonyl fluoride (1 mM) and 1X protease inhibitor
mixture (Sigma) were then added and the lysate incubated for 30 min at 4°C. Cell lysates
were sonicated and then centrifuged at 13,000 x g for 15 min at 4°C. The supernatant was
assayed by the Bradford method to determine protein content. Equal amounts of protein
were mixed with the Laemmli sample buffer and heated for 5 min at 75°C. The samples
3-85
were loaded onto a 10% SDS-PAGE gel and transferred to a nitrocellulose membrane and
immunoblotted with anti-phospho Src (416) and total Src antibodies (Cell Signaling
Technology). Blots were developed with ECL Plus kit (GE Healthcare) for 3 min. Blots
were subsequently stripped (63 mM Tris base, 70 mM SDS, 0.0007% 2-mercaptoethanol,
pH 6.8) and reprobed for further use. Western blot densitometry data were obtained using
MCID Basic 7.0 software (Imaging Research).
Membrane potential assay fluorescence monitoring- Membrane potential in the
cerebrocortical neuron cultures was determined using the FLIPR membrane potential
(FMP) assay (Molecular Devices) as previously described (George et al., 2009). FMP
blue dye was used to assess the membrane potential of neurons in culture. Quantification
of changes in membrane potential was derived using KCl as a reference. In preliminary
experiments we determined that, with cerebrocortical neurons in culture, the optimum dye
concentration was one-eighth of that suggested by the manufacturer. After removing the
culture medium, 180 µl of assay buffer was added to the neurons, and the plate was
incubated at 37°C in a 5% CO2 and 95% humidity atmosphere for 30 min. For KCl
calibration measurements, varying concentrations of 10X KCl standard solutions in assay
buffer were prepared. After 30 min equilibration incubation, the plate was transferred to a
FlexStation II chamber, and the fluorescence measurements were performed at 37°C.
Neurons were excited at 530 nm, and emission was recorded at 565 nm at 2 s intervals.
After recording the baseline for 60 s, either 20 µl KCl or ATX was added to a final
volume of 200 µl at a rate of 26 µl/s, and the fluorescence was monitored for an
additional 240 s.
3-86
A linear regression analysis of the log [K+] versus FMP blue fluorescence change
(F-F0) was generated. We used the Goldman–Hodgkin–Katz equation to generate a
standard curve for the estimation of membrane potential (EM) at various concentrations of
extracellular K+:
, ,
[ ] [ ] [ ], ln( )
[ ] [ ] [ ]out out inNa K Cl
m K Na Clin in outNa K Cl
P Na P K P ClRTEF P Na P K P Cl
+ + −
+ + −
+ + −+
+ + −
+=
+ + (equation 4)
where EM is membrane potential, R is universal gas constant, T is temperature using the
Kelvin scale, and PK, PNa, and PCl are permeabilities for K+, Na+, and Cl–, respectively.
[K+]out, [Na+]out, and [Cl–]out, and [K+]in, [Na+]in, and [Cl–]in are the respective extracellular
and intracellular concentrations of K+, Na+, and Cl–. A 1 d in vitro (DIV-1) neuronal [Cl–
]in value of 140 mM was used for these calculations(35). The regression for the [K+]out
versus Δfluorescence and EM was used for estimating ATX-induced change in membrane
potential.
Electrophysiology- Single-channel currents were recorded at 23°C in the cell-attached
configuration (Hamill et al., 1981). Patch pipettes were pulled from borosilicate glass
capillaries (Warner Instruments), coated with Sylgard 184 (Dow Corning) and fire-
polished to a resistance of 10–15 MΩ when filled with the pipette solution. The external
recording solution consisted of Mg2+-free Locke's buffer with 20 µM EDTA to chelate
trace amounts of divalent cations. ATX was always bath-applied. The patch pipette
solution consisted of extracellular Locke's buffer without MgCl2 and with 100 µM
NMDA and 100 µM glycine. In some experiments, 10 µM strychnine, 10 µM bicuculline
methiodide and 10 µM DNQX were included in the external solution to block nonspecific
3-87
components. All recordings were done from DIV-1 cerebrocortical neurons. Cell-attached
patch recordings were done using an Axopatch 200B amplifier (Molecular Devices),
filtered at 8 kHz (–3 dB, 8-pole Bessel), and digitized at 40 kHz digitized with Axon
pClamp 10.2 software. The pipette potential was +60 mV. Records were idealized with a
segmental k-means algorithm (Qin, 2004) using QUB software (www.qub.buffalo.edu).
All conductance levels were assumed to be equal for the analysis. Dwell-time histograms
were generated and fitted using Channelab (Synaptosoft) with an imposed dead time of 50
µs. The open probability (Po), mean open time, and amplitude were compared by paired t
test. For representation in figures, the Po and mean open time were normalized to the
average of respective control values. The corresponding ATX-treated values were
normalized to their paired control values.
3.3 Results
Antillatoxin is a VGSC activator in immature cerebrocortical neurons. In previous
reports we demonstrated that ATX is an activator of VGSCs in cerebellar granule
neurons (Li et al., 2001) and mature (DIV-9) cerebrocortical neurons (Cao et al., 2008).
We therefore sought to determine whether immature cerebrocortical neurons were also
sensitive to ATX-induced elevation of [Na+]i. We assessed ATX-induced elevation of
[Na+]i in DIV-1 cerebrocortical neurons loaded with SBFI. As shown in Figure 1A & B,
30 nM ATX elevated [Na+]i in DIV-1 cerebrocortical neurons. To confirm that the
observed Na+ influx was mediated by VGSCs, we tested the influence of TTX, a selective
antagonist of VGSCs, on the response to ATX. Pretreatment of DIV-1 cerebrocortical
3-88
neurons with TTX (1 μM) abolished ATX-induced Na+ influx. These results indicate that
ATX is an activator of VGSCs in DIV-1 cerebrocortical neurons.
Antillatoxin enhances neurite outgrowth in immature cerebrocortical neurons. We next
wanted to determine the influence of ATX on neuritogenesis in immature cerebrocortical
neurons. Three hours post plating, primary cultures of immature cerebrocortical neurons
were exposed to various concentrations of ATX ranging from 1-1,000 nM for 24 h and
total neurite outgrowth was then assessed. Either immunostaining of PGP 9.5 or diolistic
labeling were used to visualize neurons and determine the influence of ATX on neurite
outgrowth (Fig. 2A). ATX significantly enhanced total neurite outgrowth in immature
cerebrocortical neurons with concentrations of 30 and 100 nM producing a robust >2-fold
increase in total neurite length. As previously observed with PbTx-2, the ATX
concentration-response profile was bidirectional, or hormetic (Fig 2B).
Antillatoxin-induced neurite outgrowth is mediated by VGSCs. Given that ATX is a
VGSC activator with the ability to augment neurite outgrowth, we wanted to confirm the
involvement of VGSCs in the latter functional response. DIV-1 cerebrocortical neurons
were coincubated in the presence or absence of TTX (1 µM) and 30 nM ATX for 24 h
and total neurite length was determined. Consistent with the involvement of VGSCs,
TTX completely abolished ATX- induced neurite outgrowth (control, 109.5 ± 22.5 μm;
TTX, 153.23 ± 8.4 μm; ATX, 236.65 ± 17.9 μm; TTX plus ATX, 115.3 ± 11.8 μm) (Fig
3A, B).
Antillatoxin-induced neurite outgrowth involves NMDARs, VGCCs and the Ca2+
dependent CaMKK pathway. Inasmuch as previous studies have indicated that activity-
3-89
dependent neuritogenesis and neuronal development involve Ca2+ influx pathways
through NMDAR and voltage-gated calcium channels (VGCC) with subsequent
engagement of a CaMKK pathway (Konur and Ghosh, 2005; Wayman et al., 2006), we
assessed the role of this signaling cascade in ATX-induced neurite outgrowth. Co-
incubation of MK-801 (1 µM), an uncompetitive antagonist of NMDAR with 30 nM
ATX abrogated ATX-enhanced neurite outgrowth in immature cerebrocortical neurons
(Fig 4A,B) (control, 109.5 ± 22.5 µm; ATX, 236.65 ± 17.9 µm; ATX plus MK-801, 113.8
± 9.8 µm) demonstrating that ATX-enhanced neurite outgrowth involves NMDARs. To
investigate the role of VGCCs in the response to ATX, we used the L-type calcium
channel blocker, nifedipine (1 µM). Nifedipine pretreatment partially reduced ATX-
stimulated neurite outgrowth (Fig 4A,B) (control, 109.5 ± 22.5 µm; ATX, 236.65 ± 17.9
µm; ATX plus nifedipine, 166.3 ± 13.45 µm) suggesting that VGCCs may play a role in
the response to ATX. Next we investigated the involvement of a downstream Ca2+
dependent CaMKK in ATX-induced stimulation of neurite outgrowth. CaMKK is an
important upstream activator of essential signaling mediators of activity-dependent
neurite outgrowth such as CaMK1, CaMKIV, and MAPKs. STO-609 (2.6 µM), a
selective CaMKK inhibitor (Tokumitsu et al., 2002), eliminated the stimulatory effect of
ATX on neurite outgrowth in immature cerebrocortical neurons (Fig 4A,B) (control,
109.5 ± 22.5 µm; ATX, 236.65 ± 17.9 µm; ATX plus STO-609, 120.4 ± 9.02 µm). This
observation suggests that a Ca2+-dependent CaMKK pathway contributes to the
stimulatory effects of ATX on neuritogenesis.
Antillatoxin-induced neurite outgrowth is mediated by Src family tyrosine kinase
activation. Activity-dependent neurite outgrowth involves upregulation of NMDAR
3-90
function. As earlier studies (Salter and Kalia, 2004; Yu et al., 1997; Yu and Salter, 1998)
have shown that [Na+]i and activated Src family kinases (SFKs) upregulate NMDAR
function, we reasoned that Src family kinases may participate in ATX-enhanced neurite
outgrowth. Exposure of neurons to the Src family kinase inhibitor PP2 (2 µM), but not its
inactive congener PP3 (2 µM), eliminated the stimulatory effect of ATX on neurite
outgrowth. These findings establish a role for Src family kinases in ATX-induced
stimulation of neurite outgrowth (Fig 5A,B) (control, 118.8 ± 12.9 µm; ATX, 194.4 ±
17.2 µm; ATX plus PP2, 83.9 ± 9.7 µm; ATX plus PP3, 181.3 ± 15.3 µm). The catalytic
activity of Src kinase is controlled by phosphorylation and dephosphorylation events,
primarly that of Y416. Intermolecular autophosphorylation of Y416 stimulates Src
kinase activity by permitting access to its substrates and ligands (Yu et al., 1997). To
assess the ability of ATX to activate Src, we determined the phosphorylation of the Y416
residue using an anti-phospho-Y416 Src antibody. Immature cerebrocortical neurons
were exposed to 30 nM of ATX and cell lysates were collected at various time periods for
western blot analysis. These results revealed that 30 nM ATX produced a robust
activation of Src kinase as reflected in the sustained increase in the phosphorylation of
tyrosine 416 (Fig 5B,C). These findings indicate that ATX exposure produces an
activation of Src kinase that is temporally correlated with the stimulation of neurite
outgrowth.
Antillatoxin increases intracellular sodium levels in immature cerebrocortical neurons.
Given that the earlier studies of Yu and Salter (Yu et al., 1997; Yu and Salter, 1998; Yu,
2006) demonstrated that [Na+]i is a regulator of NMDAR-mediated signaling, it was
important to quantify the magnitude of ATX-induced elevation of [Na+]i in immature
3-91
cerebrocortical neurons. SBFI, a sodium-sensitive fluorescent indicator, was used to
determine the influence of ATX on [Na+]i in DIV-1 cerebrocortical neurons. Full in situ
calibration was performed in DIV-1 cerebrocortical neurons to determine the relationship
between the ratiometric SBFI signal and [Na+]i (Cao et al., 2008; George et al., 2009).
Cells loaded with the SBFI were excited at 340 and 380 and the emitted fluorescence was
recorded at 505. The 340/380 emission ratio was calculated after background correction
(Fig. 6A). A three-parameter hyperbolic function adequately fit the calibration data
relating SBFI fluorescence ratio to [Na+]i (Fig. 6B). ATX produced a concentration-
dependent increase in [Na+]i (Fig. 6C) with an EC50 value of 114.2 nM (70.8 to 184.2
nM, 95% CI). The in situ SBFI calibration showed that basal [Na+]i in DIV-1
cerebrocortical neurons was 17.3 ± 0.37 mM, and ATX produced a maximum elevation
of 78.6 ± 6.9 mM (Fig. 6D). Since a 30 nM concentration of ATX was sufficient to
produce a robust increase in neurite outgrowth, it was important to quantify the [Na+]i
increment associated with this treatment. The 30 nM ATX treatment produced a
maximum [Na+]i of 26.1 ± 0.4 mM, representing an increment of 8.8 mM over basal.
Previous reports in hippocampal neurons suggested that an increment of [Na+]i of 10 mM
was sufficient to produce significant increases in NMDAR channel activity (Yu and
Salter, 1998; Yu, 2006). It has moreover been reported that increments of [Na+]i of >5
mM may represent a critical threshold required to regulate NMDAR-mediated Ca2+ influx
in primary cultures of hippocampal neurons (Xin et al., 2005). Consistent with these
findings, the increment of [Na+]i detected in immature cerebrocortical neurons appears
sufficient to upregulate NMDAR function.
3-92
Antillatoxin-evoked change in membrane potential is inadequate to relieve the Mg2+
blockade of NMDARs. Given the evidence in support of the involvement of NMDARs in
ATX-induced stimulation of neuritogenesis, we considered mechanisms apart from the
increment of [Na+]i. The ability of ATX to engage NMDARs could be a consequence of
either the elevation of [Na+]i or neuronal depolarization with attendant relief of the Mg2+
block of NMDAR. To ascertain the magnitude of ATX-induced membrane
depolarization, we assessed membrane potential changes in DIV-1 cerebrocortical
neurons using the membrane-potential sensitive fluorescence dye, FMP blue.
As previously reported (George et al., 2009), FMP blue behaved as a Nernstian
fluorescent indicator of membrane potential in DIV-1 cerebrocortical neurons. This was
demonstrated by assessing the relationship between extracellular K+ concentration and
fluorescence intensity. Extracellular K+ produced a concentration-dependent increase in
maximum FMP blue fluorescence consistent with a depolarization-induced redistribution
of the lipophillic anion dye and attendant increase in fluorescence quantum efficiency
(Fig. 7A). As depicted in Fig. 7B, the regression analysis for K+ concentration-dependent
changes FMP blue fluorescence showed marked linear correlation (r2= 0.99). For a
Nernstian fluorescent indicator of membrane potential, the ratio of fluorescence inside to
the outside of the cell should be related to the membrane potential as described by the
Nernst equation (Ehrenberg et al., 1988). This prediction is based on the principal that the
membrane potential of isolated neurons is largely the result of the K+ diffusion potential
(Hille, 1992). We therefore used the Goldman-Hodgkin-Katz equation to generate a
standard curve for the estimation of membrane potential (EM) at various concentrations of
extracellular K+. The membrane potential of cerebrocortical neurons was dependent on
3-93
the external concentration of K+ (Hille, 1992). The concordance of the [K+]out versus
membrane fluorescence and [K+]out versus EM regressions indicates that changes in
cerebrocortical neuron FMP blue fluorescence can be used to estimate membrane
potential. Therefore, the relationship between fluorescence change and EM depicted in
Fig. 7B was generated to determine ATX-induced changes in membrane potential of
cerebrocortical neurons. The resting membrane potential of DIV-1 cerebrocortical
neurons was found to be –29.6 mV. This is consistent with previous demonstrations of a
relatively depolarized resting membrane potential of immature neurons that later becomes
more hyperpolarized as neurons mature (Kim et al., 1995; Ramoa and McCormick,
1994). As shown in Figure 7C, ATX produced a rapid and concentration-dependent
increment in FMP blue fluorescence in DIV-1 cerebrocortical neurons. Nonlinear
regression analysis of the ATX concentration–response relationship yielded an EC50
value of 92.3 nM (63.6–136.8 nM, 95% CI) (Fig. 7D). Because the 30 nM concentration
of ATX was sufficient to elevate [Na+]i, it was important to assess the membrane potential
changes associated with this treatment. The 30 nM ATX treatment produced a transient
increase in FMP blue fluorescence that was equivalent to the fluorescence change
produced by an extracellular K+ concentration of 7.6 mM. The corresponding membrane
potential change was accordingly found to be negligible, representing only a 0.9 mV
depolarization (from -29.6 ± 0.01 to 28.7 ± 0.15 mV). This change in membrane potential
would, therefore, not be sufficient to influence the voltage-dependent Mg2+ block of
NMDARs (Mayer et al., 1984).
3-94
Antillatoxin increases intracellular calcium levels ([Ca2+]i) in DIV 1 cerebrocortical
neurons. Previous studies have suggested that activity-dependent neuritogenesis and
neuronal development involves Ca2+- dependent signaling pathways through NMDAR
and VGCCs. Due to the finding that ATX-induced neurite outgrowth involved NMDARs
and VGCCs, we hypothesized that ATX exposure would produce Ca2+ influx in these
immature cerebrocortical neurons. To investigate this, cells loaded with fluo-3 were
exposed to various concentrations of ATX and [Ca2+]i was monitored. ATX produced
rapid and concentration-dependent increases in [Ca2+]i with even 30 nM ATX producing
a significant increase in calcium influx (Fig 8A).
To delineate the Ca2+ influx pathways triggered by ATX, the role of VGSCs, NMDARs
and VGCCs in DIV-1 cerebrocortical neurons were investigated. A pharmacologic
evaluation of the [Ca2+]i response to 100 nM ATX was performed. Cells were pretreated
with specific antagonists: TTX (VGSCs), MK-801 (NMDARs) or nifedipine (VGCCs) to
evaluate the role of these channels in ATX induced Ca2+ influx. TTX (1 µM) completely
blocked the response to ATX (data not shown), while MK-801 (1 µM) and nifedipine (1
µM) both significantly reduced ATX-induced Ca2+ influx (Fig 8B,C). Given the
previously demonstrated role of SFK activation in the upregulation of NMDAR function
and in ATX-induced neurite outgrowth, we examined the role of SFKs in ATX-induced
Ca2+ influx. PP2 (2 µM), a specific SFK family inhibitor, but not PP3 (2 µM), blocked
ATX stimulation of Ca2+ influx consistent with the involvement of a SFK in this response
(Fig 8D,E).
3-95
Antillatoxin increases NMDA receptor single-channel open probability but not the mean
open time. To gain insight into the effect of ATX on single-channel properties of NMDA
receptors, unitary currents were recorded from DIV-1 cerebrocortical neurons. Cell-
attached patch recording was performed with 100 µM NMDA and 100 µM glycine in the
patch pipette at a patch potential of +60 mV. Experiments were performed in the nominal
absence of extracellular Mg2+ in the recording buffer supplemented with 20 µM EDTA to
chelate trace amounts of divalent ions. In the majority of patches, only single openings
were observed with no apparent simultaneous double openings. The absence of double
openings can be presumably attributed to the supposedly low expression of NMDA
receptors in immature cerebrocortical neurons. Patches in which we observed
simultaneous double openings were not further analyzed. Single-channel recordings were
idealized using the QUB and analyzed using ChanneLab with an imposed resolution of
50 µs. Bath application of 100 nM ATX significantly increased the open probability (Po)
of NMDA receptors from 0.0053 ± 0.002 under control conditions to 0.012 ± 0.004 (206
± 46% of control) after ATX (n = 6, p < 0.05, paired t test) (Fig. 9A,B) The mean open
time was not affected by ATX (1.862 ± 0.38 ms without ATX; 1.90 ± 0.38 ms, with
ATX-101 ± 4.5% of control) (n = 8, p < 0.05, paired t test) (Fig. 9B). ATX similarly did
not affect the amplitude of single-channel currents (data not shown). The composite open
and shut dwell-time histograms were generated and fitted using Channelab. The open
time histogram could be fitted by the sum of three exponential components with time
constants of 0.127 (27%), 1.584 (53%) and 3.5 (20%). The time constants after ATX
application were 0.091 (19%), 1.409 (70%) and 4.67 (11%). The composite shut time
histograms could be fitted by sum of five exponential components with time constants of
3-96
0.7 (34%), 0.095 (17%), 20.8 (14%), 240 (22%), and 1090 (12%). The time constants
were similar after ATX application 0.78 (35%), 0.169 (23%), 11.0 (10%), 84 (16%), and
580 (16%), except that the duration of the longer shut time constants were reduced (Fig.
9C).
3.4 Discussion
ATX is a novel activator of VGSC; however, its precise recognition site on the channel
protein remains to be defined. The structure of ATX includes asymmetric carbon atoms
and the (4R,5R)-isomer is the naturally occurring compound. The (4R,5R)-isomer appears
in profile as an “L” shape with a hydrophobic interior and a cluster of hydrophilic groups
on the exterior of the macrocycle (Li et al., 2004). Thus the (4R,5R)-configuration is
important for creating a molecular topology that is recognized by the acceptor site on the
voltage-gated sodium channel alpha subunit.
We have previously shown that ATX allosterically enhances the specific binding
of [3H]batrachotoxin to intact cerebellar granule cells (Li et al., 2001) This effect of ATX
on [3H]batrachotoxin binding was synergistically augmented by brevetoxin. The strong
synergistic interaction of the ATX recognition site with neurotoxin site 5 suggests that
these sites may be topologically close and/or conformationally coupled. The results
obtained using [3H]batrachotoxin as a probe for sodium channel conformation allowed us
to exclude the interaction of ATX with neurotoxin sites 1, 2, 3, and 5 on VGSCs. Site 1
was ruled out because tetrodotoxin and saxitoxin bind to the outer vestibule of the pore of
the ion channel and allosterically inhibit the binding of [3H]batrachotoxin; this is an
effect that is antipodal to that of ATX. We were able to rule out sites 2 and 5 inasmuch as
3-97
these sites display positive allosteric coupling to the ATX site. Neurotoxin receptor site 3,
the target for α-scorpion toxins and sea-anemone toxins, was excluded because ATX
enhanced [3H]batrachotoxin binding in the presence of a maximally effective
concentration of sea-anemone toxin. Although we cannot exclude an interaction of ATX
with neurotoxin site 4, the target for β-scorpion toxin, it is reasonable to posit that ATX
binds to a novel recognition domain on the α-subunit of the VGSC. The relatively small
lipotripeptide structure of ATX would not be restricted to an extracellular target, as is the
case for the scorpion toxins, which are composed of 60-65 amino acids. Given the unique
structure and mechanism of action of ATX, we sought to further characterize its
pharmacologic actions in cerebrocortical neurons.
We have earlier demonstrated that NMDA receptor function may be increased
through activation of VGSCs with attendant elevation of intracellular sodium in
cerebrocortical neurons (George et al., 2009). VGSC activators function as gating
modifiers that elevate [Na+]i in the absence of substantial depolarization of neurons (Cao
et al., 2008; George et al., 2009). These findings have been confirmed and extended in
the present study by demonstrating that the structurally novel lipopeptide, ATX, elevates
intracellular Na+, increases NMDAR function and enhances neurite outgrowth in DIV-1
cerebrocortical neurons. These findings in DIV-1 murine cerebrocortical cultures provide
compelling evidence in support of a role for [Na+]i in activity-dependent processes of
neuronal development.
Antillatoxin enhances neurite outgrowth- Here we found that ATX enhanced
neurite outgrowth in DIV-1 cerebrocortical neurons as a result of elevation of
3-98
cytoplasmic [Na+], potentiation of NMDAR function and stimulation of calcium influx.
ATX enhanced total neurite outgrowth in immature cerebrocortical neurons in a
bidirectional, or hormetic, concentration-response relationship with 30-100 nM producing
a robust increases of more than 2-fold (Fig. 2). Thus, the ability of ATX to augment
NMDAR channel activity translated into an enhancement of the trophic influence of
NMDAR on developing cerebrocortical neurons. Based on the premise that the effects of
neuronal activity on dendritic arbor growth and structural plasticity are primarily
mediated by engagement of NMDA receptors (Tolias et al., 2005), our results suggest
that ATX activation of sodium channels with attendant enhancement of NMDA receptor
signaling mimics the response to neuronal activity.
Antillatoxin concentration- response for neurite growth is an inverted-U- An
inverted-U model describes the relationship between NMDA receptor activity and
neuronal survival and growth (Lipton and Nakanishi, 1999). This inverted-U
concentration–response relationship has primarily, but not exclusively, been attributed to
[Ca2+]i regulation. An optimal window for [Ca2+]i is required for activity-dependent
neurite extension and branching, with lower levels stabilizing growth cones and higher
levels stalling them, in both cases preventing extension (Gomez and Spitzer, 2000; Hui et
al., 2007). Although the precise mechanism for the ATX bidirectional concentration-
response curve is not known, one plausible explanation is therefore related to the
involvement of NMDA receptors in the trophic response to ATX. Other potential
explanations for the inverted-U response include the possibility that high concentrations
of ATX might promote slow inactivation of VGSCs with attendant reduction in sodium
influx (Mitrovic et al., 2000). Alternatively, high concentrations of ATX could increase
3-99
VGSC internalization, which has been shown to be a consequence of Na+ influx in
immature neuronal tissue. These results with ATX concur with those recently reported
for PbTx-2-induced stimulation of neuritogenesis in DIV-2 cerebrocortical neurons
(George et al., 2009). Although PbTx-2 is known to activate neurotoxin site 5 on VGSC
α-subunits, the molecular determinants for ATX on the VGSC remain to be defined (Li et
al., 2001).
ATX stimulated Ca2+ influx in cerebrocortical neurons through both NMDARs and
VGCCs. Ca2+-signaling pathways initiated by Ca2+ entry through L-type Ca2+ channels
and NMDA receptors have been shown to differ (Bading et al., 1993). Although MK-801
and nifedipine produced comparable reductions in ATX-induced Ca2+ influx, we found
that the NMDAR antagonist MK-801 completely blocked ATX-enhanced neurite
outgrowth whereas the L-type calcium channel blocker nifedipine produced only a partial
block of the stimulation of neurite outgrowth. These results suggest that Ca2+ which
enters neurons through NMDA receptors may have privileged access to the CaMKK and
CaMKI signaling elements that drive neuritogenesis.
In mature neurons a strong depolarizing stimulus (50 mM KCl) is required for the
engagement of L-type Ca2+ channels in dendritic growth and arborization, whereas a
smaller depolarizing stimulus (16 mM KCl) induced neurite outgrowth preferentially due
to Ca2+ influx through NMDARs (Redmond et al., 2002; Wayman et al., 2006). Our
observation that ATX concentrations of 30-100 nM provided a sufficient stimulus to
produce Ca2+ influx through VGCCs may be explained by the relatively depolarized
resting membrane potential of immature cerebrocortical neurons. The resting membrane
3-100
potential of DIV-1 cerebrocortical neurons was found to be –29.6 mV and ATX (30-300
nM) produced modest changes of 1-5 mV. These modest changes in membrane potential
produced by ATX may however be sufficient to activate L-type Ca2+ channels given the
relatively depolarized resting membrane potential (Nowycky et al., 1985).
Regulatory influence of Na+ on NMDAR activity- Recent studies have shown that
intracellular Na+ might act as a signaling molecule. Based on the original work of
Hodgkin and Huxley (HODGKIN and HUXLEY, 1952) with squid axons, a single action
potential was calculated to minimally change the Na+ electrochemical gradient (Hille,
1992). The situation in mammalian neurons with fine axons, dendrites, and spines is,
however, much different, due to greater surface-to-volume ratios. Thus, a single action
potential may elevate [Na+]i substantially (Hille, 1992). Yu and Salter (Yu et al., 1997;
Yu and Salter, 1998) previously demonstrated that elevation of intracellular Na+ increases
NMDA receptor mediated whole-cell currents and NMDAR single channel current by
increasing the open probability and mean open time of the channel. An increment of
[Na+]i of 10 mM was sufficient to produce significant increases in NMDA receptor
single-channel activity. They used veratridine, a VGSC modulator to demonstrate that
Na+ influx through TTX-sensitive VGSC was sufficient to upregulate NMDAR activity.
Moreover, this Na+-dependent regulation of NMDA receptor function was shown to be
controlled by Src-induced phosphorylation of the receptor (Yu et al., 1997; Yu and Salter,
1998). These results were extended in the present study using the novel sodium channel
activator ATX as a probe to elevate intracellular Na+. Single channel current recording in
presence of ATX directly demonstrated the enhancement of NMDA receptor function. An
increase in intracellular Na+ and Src activation following exposure to ATX increased the
3-101
open probability of the NMDAR. The shut time histogram with slow time constants
resemble NR2B-containing receptors (Erreger et al., 2005), consistent with the
expression of NR1/NR2B-containing receptors in immature neurons (Williams et al.,
1993). Given that the single-channel recordings were done in the absence of extracellular
Mg2+, these results additionally argue against relief of the voltage-dependent Mg2+ block
of the NMDAR in the actions of ATX. These data, therefore, confirm the regulatory
influence of Na+ on NMDAR channel activity in hippocampal neurons described
previously (Yu and Salter, 1998) and extend this relationship between [Na+]i and NMDA
receptor function to cerebrocortical neurons.
ATX represents a structurally and mechanistically novel activator of VGSCs whose
recognition domain on the α-subunit remains to be established (Li et al., 2001). Here we
found that ATX was capable of mimicking activity-dependent neuronal development by
upregulating NMDAR function. We propose a model for ATX-induced neuritogenisis
(Fig. 10) which involves direct activation of TTX-sensitive VGSCs, elevation of [Na+]i,
activation of a Src family kinase, potentiation of NMDAR function leading to Ca2+ influx
and engagement of a CaMKK pathway. We have recently reported another activator of
VGSCs, brevetoxin 2, is capable of upregulating NMDAR function and stimulating
neuritogenesis (George et al., 2009).
Structurally dissimilar sodium channel activators therefore appear capable of mimicking
activity-dependent structural plasticity by upregulating NMDA receptor signaling
pathways.
3-102
3-103
Figure 3-1: ATX increases intracellular sodium levels in DIV-1 cerebrocortical neurons. A. Time-response curve for ATX stimulation of Na+ influx. This ATX-induced stimulation of Na+ influx was prevented by coapplication of 1 µM TTX. B. Histogram representing SBFI fluorescence ratio (340/380) values following the indicated treatments. Data shown are from an experiment performed in octuplicates. ***p < 0.001, unpaired t test.
3-104
Figure 3-2: Effect of ATX on neurite outgrowth. A.Representative images of DiI-loaded immature cerebrocortical neurons at 24 h post plating (scale bar, 10 µm). Various concentrations of ATX were added to the culture medium at 3 h after plating. Depicted neurons were visualized by diolistic loading with DiI. B. Quantification of concentration-response effects of ATX on neurite outgrowth at 24 h post plating. ATX-enhanced neurite outgrowth displayed a hormetic concentration-response relationship with maximum enhancement seen at 30-100 nM ATX. Quantification of total neurite length was performed with Image Pro Plus.
3-105
Experiment was performed twice and each point represents the mean value derived from analysis of 25-30 neurons. ***p < 0.001, unpaired t test.
3-106
Figure 3-3. Effect of TTX on ATX-induced neurite outgrowth in immature cerebrocortical neurons. A. Representative images (scale bar, 10 µm) and (B) quantification of the effects of TTX on ATX-enhanced neurite outgrowth at 24 h post plating. Neurons were treated with 30 nM ATX in the presence and absence of 1 µM TTX beginning at 3 h post plating. Experiment was repeated twice and 25-30 neurons were quantified for each exposure condition. *p < 0.05, unpaired t test.
3-107
Figure 3-4:Pharmacological evaluation of signaling pathways involved in ATX-enhanced neurite outgrowth. A. Representative images (scale bar, 10 µm) and (B) quantification of neurite extension at 24 h. The 30 nM ATX exposure was examined in the presence or absence of MK-801 (1 µM), nifedipine (1 µM) or STO-609 (2.6 µM) beginning at 3 h after plating. Experiment was repeated twice and 25-30 neurons were quantified for each exposure condition. ***p < 0.001, *p < 0.05, unpaired t test.
3-108
Figure 3-5: ATX-induced neurite extension involves a Src Family kinase.
A. Representative images (scale bar, 10 µm) and (B) quantification of neurite extension at 24 h. Cerebrocortical neurons were treated with 30 nM ATX in the presence or absence of either 2 µM PP2 or PP3 beginning at 3 h after plating. Experiment was repeated four times and 20-30 neurons neurons were quantified for each exposure condition. ***p < 0.001, *p < 0.05, unpaired t test. C.Tyrosine phosphorylation (Y416) of Src kinase determined by immunoblotting. Cerebrocortical neurons were treated with 30 nM ATX beginning at 3 h post plating and P-Src (Y416) assessed at the indicated times. A representative blot is shown. The experiment was performed twice with independent cultures. Also depicted is the quantitative analysis of the relative band densities of immunoblots. Each bar represents mean ± SEM of two values.
3-109
Figure 3-6: Quantification of ATX-induced increase of intracellular sodium levels in DIV-1 cerebrocortical neurons.
A. In situ calibration of SBFI fluorescence ratio (340/380). Time-response data show stepwise changes in SBFI fluorescence ratio values evoked by successive increments in extracellular sodium concentration. B. Three parameter hyperbolic fit adequately describes calibration data. C. Nonlinear regression analysis of the ATX concentration-response data (EC50 =114.2 nM; 70.8-184.1 nM 95% CI). Data represent the mean ± SEM of 2 separate experiments each with 2-5 replicates. The scale on the left ordinate represents the SBFI fluorescence ratio, while that on the right ordinate depicts the [Na+]i determined from the calibration curve shown in C. Addition of 30 nM ATX produced an 8.8 mM increment in [Na+]i over basal.
3-110
Figure 3-7:ATX evoked change in membrane potential in DIV-1 cereobrocortical neurons.
A. Concentration-response profile for KCl-evoked FMP blue fluorescence change as a function of time. Each point represents the mean ± SEM of 3-9 values. B. The integrated time-response data for the increment in FMP blue fluorescence (Fmax-F0) plotted as a function of K+ concentration. The displayed regression and correlation coefficient (r2 = 0.995) were derived from linear regression analysis. The right ordinate scale shows membrane potential for each [K+] which was calculated using the Goldman-Hodgkin-Katz equation as described in materials and methods. The resting membrane potential was -29.6 mV. C. Time-response profiles for ATX-
3-111
induced changes in membrane potential as determined by changes in FMP blue fluorescence. D. Nonlinear regression analysis of the integrated time-response data for the increment in FMP blue fluorescence (Fmax-F0) as a function of ATX concentration. The membrane potential values were determined by performing K+ calibration regressions in the same culture plate. The membrane potential change evoked by 30 nM ATX was 0.9 mV.
3-112
Figure 3-8: ATX-induced Ca2+ influx and pharmacological evaluation in DIV-1 cerebrocortical neurons. A. Time-response profile of ATX-induced Ca2+ influx in fluo-3 loaded cerebrocortical neurons. Data shown are from a representative experiment performed with 2-5 replicates per point and repeated twice. ATX (30 nM) produced a significant increase in Ca2+
influx in these DIV-1 cerebrocortical neurons (inset). B. Pharmacological evaluation of ATX (100 nM) induced Ca2+ influx. Data are from representative experiment performed in triplicate and repeated twice. Cerebrocortical neurons were treated with either 1 µM MK-801 or 1 µM nifedipine before the addition of 100 nM ATX. C. Histogram representing quantification of the data shown in B. MK-801 and nifedipine significantly blocked ATX-induced calcium influx. D. Involvement of Src family kinase in ATX-induced Ca2+ influx. Data are from representative experiment performed in triplicate and repeated twice. Cerebrocortical neurons were treated with
3-113
either 2 µM PP2 or 2 µM PP3 before the addition of 100 nM ATX. E, Histogram representing quantification of the data shown in D. **p < 0.005, *p < 0.05, unpaired t test
3-114
Figure 3-9:. Increase in NMDA receptor channel open probability by ATX. A. Cell-attached patch recording from DIV-1 cerebrocortical neurons. NMDA receptor unitary currents were evoked by100 µM NMDA and 100 µM glycine in the patch pipette (pipette potential = +60 m V, filtered at 5 kHz for representation, digitized at 40 kHz). Enhancement of NMDA receptor activity by bath application of 100 nM ATX. B. Bath application of 100 nM ATX increased NMDA receptor channel open probability (Po), but not the mean open time (MOT) (n=6, *p<0.05, paired t test). C. Pooled dwell-time histograms were fitted using Channel lab. The open time histogram was fitted by the sum of three Gaussian components, and the shut time histogram was fitted by the sum of five Gaussian components. The time constants and area are described in Results.
3-115
Figure 3-10:Schematic diagram of the pathways involved in ATX-induced neurite outgrowth
3-116
3.5 References
Bading H, Ginty DD, Greenberg ME. (1993) Regulation of gene expression in
hippocampal neurons by distinct calcium signaling pathways. Science 260:181-186.
Cao Z, George J, Gerwick WH, Baden DG, Rainier JD, Murray TF. (2008) Influence of
lipid-soluble gating modifier toxins on sodium influx in neocortical neurons. J
Pharmacol Exp Ther 326:604-613.
Ehrenberg B, Montana V, Wei MD, Wuskell JP, Loew LM. (1988) Membrane potential
can be determined in individual cells from the nernstian distribution of cationic dyes.
Biophys J 53:785-794.
Erreger K, Dravid SM, Banke TG, Wyllie DJ, Traynelis SF. (2005) Subunit-specific
gating controls rat NR1/NR2A and NR1/NR2B NMDA channel kinetics and synaptic
signalling profiles. J Physiol 563:345-358.
George J, Dravid SM, Prakash A, Xie J, Peterson J, Jabba SV, Baden DG, Murray TF.
(2009) Sodium channel activation augments NMDA receptor function and promotes
neurite outgrowth in immature cerebrocortical neurons. J Neurosci 29:3288-3301.
Ghosh A and Greenberg ME. (1995) Calcium signaling in neurons: Molecular
mechanisms and cellular consequences. Science 268:239-247.
Gomez TM and Spitzer NC. (2000) Regulation of growth cone behavior by calcium: New
dynamics to earlier perspectives. J Neurobiol 44:174-183.
3-117
Hamill OP, Marty A, Neher E, Sakmann B, Sigworth FJ. (1981) Improved patch-clamp
techniques for high-resolution current recording from cells and cell-free membrane
patches. Pflugers Arch 391:85-100.
Hille B. (1992) Ionic channels of excitable membranes, in (Anonymous ) pp 403-411,
Sinauer Associates; Sunderland, MA.
HODGKIN AL and HUXLEY AF. (1952) Currents carried by sodium and potassium
ions through the membrane of the giant axon of loligo. J Physiol 116:449-472.
Hui K, Fei GH, Saab BJ, Su J, Roder JC, Feng ZP. (2007) Neuronal calcium sensor-1
modulation of optimal calcium level for neurite outgrowth. Development 134:4479-4489.
Kim HG, Fox K, Connors BW. (1995) Properties of excitatory synaptic events in neurons
of primary somatosensory cortex of neonatal rats. Cereb Cortex 5:148-157.
Konur S and Ghosh A. (2005) Calcium signaling and the control of dendritic
development. Neuron 46:401-405.
Lee KC and Loh TP. (2006) Total synthesis of antillatoxin. Chem Commun (Camb)
(40):4209-4211.
Li WI, Berman FW, Okino T, Yokokawa F, Shioiri T, Gerwick WH, Murray TF. (2001)
Antillatoxin is a marine cyanobacterial toxin that potently activates voltage-gated sodium
channels. Proc Natl Acad Sci U S A 98:7599-7604.
Li WI, Marquez BL, Okino T, Yokokawa F, Shioiri T, Gerwick WH, Murray TF. (2004)
Characterization of the preferred stereochemistry for the neuropharmacologic actions of
antillatoxin. J Nat Prod 67:559-568.
3-118
Linden DJ, Smeyne M, Connor JA. (1993) Induction of cerebellar long-term depression
in culture requires postsynaptic action of sodium ions. Neuron 11:1093-1100.
Lipton SA and Nakanishi N. (1999) Shakespeare in love--with NMDA receptors? Nat
Med 5:270-271.
Mayer ML, Westbrook GL, Guthrie PB. (1984) Voltage-dependent block by Mg2+ of
NMDA responses in spinal cord neurones. Nature 309:261-263.
Miller FD and Kaplan DR. (2003) Signaling mechanisms underlying dendrite formation.
Curr Opin Neurobiol 13:391-398.
Mitrovic N, George AL,Jr, Horn R. (2000) Role of domain 4 in sodium channel slow
inactivation. J Gen Physiol 115:707-718.
Nowycky MC, Fox AP, Tsien RW. (1985) Three types of neuronal calcium channel with
different calcium agonist sensitivity. Nature 316:440-443.
O'Brien JA and Lummis SC. (2006) Diolistic labeling of neuronal cultures and intact
tissue using a hand-held gene gun. Nat Protoc 1:1517-1521.
Orjala J, Nagle D, Gerwick WH. (1995) Malyngamide H, an ichthyotoxic amide
possessing a new carbon skeleton from the caribbean cyanobacterium lyngbya majuscula.
J Nat Prod 58:764-768.
Perez-Otano I and Ehlers MD. (2005) Homeostatic plasticity and NMDA receptor
trafficking. Trends Neurosci 28:229-238.
3-119
Qin F. (2004) Restoration of single-channel currents using the segmental k-means
method based on hidden markov modeling. Biophys J 86:1488-1501.
Ramoa AS and McCormick DA. (1994) Developmental changes in electrophysiological
properties of LGNd neurons during reorganization of retinogeniculate connections. J
Neurosci 14:2089-2097.
Redmond L, Kashani AH, Ghosh A. (2002) Calcium regulation of dendritic growth via
CaM kinase IV and CREB-mediated transcription. Neuron 34:999-1010.
Rose CR and Konnerth A. (2001) NMDA receptor-mediated na+ signals in spines and
dendrites. J Neurosci 21:4207-4214.
Salter MW and Kalia LV. (2004) Src kinases: A hub for NMDA receptor regulation. Nat
Rev Neurosci 5:317-328.
Saneyoshi T, Wayman G, Fortin D, Davare M, Hoshi N, Nozaki N, Natsume T,
Soderling TR. (2008) Activity-dependent synaptogenesis: Regulation by a CaM-kinase
kinase/CaM-kinase I/betaPIX signaling complex. Neuron 57:94-107.
Tolias KF, Bikoff JB, Burette A, Paradis S, Harrar D, Tavazoie S, Weinberg RJ,
Greenberg ME. (2005) The Rac1-GEF Tiam1 couples the NMDA receptor to the
activity-dependent development of dendritic arbors and spines. Neuron 45:525-538.
Ultanir SK, Kim JE, Hall BJ, Deerinck T, Ellisman M, Ghosh A. (2007) Regulation of
spine morphology and spine density by NMDA receptor signaling in vivo. Proc Natl
Acad Sci U S A 104:19553-19558.
3-120
Wayman GA, Impey S, Marks D, Saneyoshi T, Grant WF, Derkach V, Soderling TR.
(2006) Activity-dependent dendritic arborization mediated by CaM-kinase I activation
and enhanced CREB-dependent transcription of wnt-2. Neuron 50:897-909.
West AE, Griffith EC, Greenberg ME. (2002) Regulation of transcription factors by
neuronal activity. Nat Rev Neurosci 3:921-931.
Williams K, Russell SL, Shen YM, Molinoff PB. (1993) Developmental switch in the
expression of NMDA receptors occurs in vivo and in vitro. Neuron 10:267-278.
Wong RO and Ghosh A. (2002) Activity-dependent regulation of dendritic growth and
patterning. Nat Rev Neurosci 3:803-812.
Xin WK, Kwan CL, Zhao XH, Xu J, Ellen RP, McCulloch CA, Yu XM. (2005) A
functional interaction of sodium and calcium in the regulation of NMDA receptor activity
by remote NMDA receptors. J Neurosci 25:139-148.
Yu XM. (2006) The role of intracellular sodium in the regulation of NMDA-receptor-
mediated channel activity and toxicity. Mol Neurobiol 33:63-80.
Yu XM, Askalan R, Keil GJ,2nd, Salter MW. (1997) NMDA channel regulation by
channel-associated protein tyrosine kinase src. Science 275:674-678.
Yu XM and Salter MW. (1998) Gain control of NMDA-receptor currents by intracellular
sodium. Nature 396:469-474.
4-121
4 CHAPTER 4 – Sodium channel
activator-stimulated neuronal
development involves BDNF-TrkB
signaling pathway.
4-122
4.1 Abstract
N-methyl D-aspartate receptor (NMDAR) activation directly stimulates calcium
influx and leads to an increase in brain-derived neurotrophic factor (BDNF) – TrkB
signaling. This pathway is implicated in activity-dependent neuronal development and
synaptic plasticity. Voltage-gated sodium channel (VGSC) activators promote neuronal
development by increasing [Na+]i and upregulating NMDAR function. Here we tested the
effect of the VGSC activator veratridine (VRT) on neurite outgrowth (NOG), synthesis
and release of BDNF and TrkB activation in DIV1 cerebrocortical neurons. Primary
cultures of murine cerebrocortical neurons were prepared from E16-17 Swiss-Webster
mice. Diolistic loading of DiI and confocal imaging were performed to visualize and
image neurons respectively to assess total neurite outgrowth was determined. BDNF
ELISA was performed to determine BDNF synthesis and release (BDNF ELISA in situ).
Activation of various signaling molecules was determined by western blotting. VRT
enhanced NOG in a hormetic concentration-response manner, and this response was
dependent on NMDAR and TrkB signaling. Inhibitors of NMDAR, TrkB, PI3K, and PLC
inhibited VRT-enhanced NOG. Acute treatment with VRT stimulated phosphorylation
of TrkB and its downstream effectors Akt, mTOR, PLC, ERK1/2 and CREB. VRT
increased BDNF synthesis and release in a concentration dependent manner; however,
VRT stimulation of TrkB phosphorylation displayed a biphasic concentration-response
curve. VRT stimulation of BDNF synthesis required VGSCs and NMDARs. These data
suggest the influence of VGSC activators on neurite outgrowth may involve BDBF
synthesis and release.
4-123
4.2 Introduction
Events that occur during neuronal development play a critical role in establishing
the morphological diversity and neuronal connectivity in brain. Dendritic arbor shape
determines the extent of neuronal connectivity and integration of synaptic signals: both
essential to the proper formation of neural circuits and proper function of nervous system.
In addition to intrinsic genetic programs, neuronal activity signals regulate neuronal
developmental events including neurogenesis, neurite outgrowth, dendritic arborization,
spinogenesis and synaptogenesis 452 Ben-Ari,Y. 2001; 206 McAllister,A.K. 2000; 1
Wong,R.O. 2002. Activity-dependent control of neuronal development primarily
involves calcium-dependent signaling and neurotrophins signaling. Activity-dependent
calcium signaling involves various calcium influx pathways including ionotropic
glutamate receptors (NMDAR) and voltage-gated Ca2+ channels (VGCCs) 254
Ghosh,A. 1995; 446 West,A.E. 2001; 32 West,A.E. 2002. Intracellular calcium acts as
a signaling molecule largely through the binding to calmodulin, a calcium-binding
protein that engages downstream Ca2+/calmodulin-dependent protein kinase (CaMK).
One such important CaMK is CaMKII, an important downstream regulator of dendritic
remodeling and synaptic activity 467 Fink,C.C. 2003; 468 Vaillant,A.R. 2002; 471
Zou,D.J. 1999; 469 Shen,K. 1998; 470 Wu,G.Y. 1998. Moreover, previous studies
have demonstrated that activity-dependent neurite outgrowth 99 Wayman,G.A. 2006
and synaptogenesis 98 Saneyoshi,T. 2008 are regulated by NMDAR-dependent
CaMKK/calmodulin kinase I-signaling cascades. Therefore, NMDARs play a critical role
in activity-dependent development and plasticity 254 Ghosh,A. 1995, dendritic
arborization 1 Wong,R.O. 2002; 84 Miller,F.D. 2003; 99 Wayman,G.A. 2006, spine
4-124
morphogenesis 36 Ultanir,S.K. 2007, and synapse formation 98 Saneyoshi,T.
2008 by stimulating these calcium-dependent signaling pathways.
In addition to neuronal activity, numerous studies have implicated a role for
neurotrophins in dendritic development. Brain-derived neurotrophic factor (BDNF)
increase dendritic complexity of cortical pyramidal neurons by increasing total dendritic
length, the number of branch points and the number of primary dendrites 460
McAllister,A.K. 1995; 461 Dijkhuizen,P.A. 2005. Also, activity-dependent increases
in [Ca2+]i trigger a release of BDNF through the regulated pathway of BDNF release
456 Hartmann,M. 2001; 455 Goodman,L.J. 1996; 457 Nakajima,T. 2008; 331
Brigadski,T. 2005; 323 Kolarow,R. 2007; 458 Balkowiec,A. 2002 and mediates
activity-dependent dendritic development and synaptic plasticity455 Goodman,L.J.
1996; 456 Hartmann,M. 2001; 464 Ghosh,A. 1994; 463 Kohara,K. 2001; 462 Kohara,K.
2003. Changes in neuronal [Ca2+]i are due to influx either through NMDAR or VGCCs
or due to release from intracellular Ca2+ stores. The calcium activates CamKII leading to
the fusion of BDNF containing secretory granules with the plasma membrane and slow
release of BDNF into the extracellular milieu 331 Brigadski,T. 2005; 323 Kolarow,R.
2007. The effects of BDNF on dendritic morphology are due to the activation of
signaling mechanisms downstream of TrkB that influence neuronal development. Three
major intracellular signaling pathways are activated by BDNF binding to TrkB receptor.
They are: 1) the PI3K-Akt pathway, 2) the Ras-MAPK pathway and 3) the PLCγ-Ca2+
pathway 267 Reichardt,L.F. 2006.
Recent studies have demonstrated that neuronal activity-mediated increases in neuronal
[Na+]i augment NMDAR function and may contribute to activity-dependent synaptic
4-125
plasticity 91 Rose,C.R. 2001; 96 Yu,X.M. 1998. Inasmuch as neuronal activity-
induced increments in cytoplasmic sodium may augment NMDAR-mediated currents, we
reasoned that intracellular Na+ may function as a signaling molecule to positively
regulate neuronal development in immature cerebrocortical neurons. In the present study,
we used veratridine (VRT), a VGSC gating modifier to manipulate [Na+]i in immature
cerebrocortical neurons. We have previously demonstrated that in cerebrocortical neurons
VGSC activators, brevetoxin (PbTx-2) and antillatoxin (ATX) elevated [Na+]i and
augmented NMDAR function 296 Jabba,S.V. 2010; 6 George,J. 2009. These VGSC
activators also enhanced neurite outgrowth in a hormetic concentration-relationship. The
inverted-U response to ATX and PbTx-2 on neurite outgrowth is similar to that of
NMDA (unpublished). Moreover, BDNF also display an inverted-U concentration-
response for retinal ganglion survival following optic nerve transaction 466 Klocker,N.
1998, and in promoting serotonergic axonal growth and remodeling in the adult brain
447 Mamounas,L.A. 2000
We therefore hypothesized that sodium channel activators stimulate neuronal
development by elevating [Na+]i , augmenting NMDAR function and enhancing BDNF
release with activation of downstream BDNF-TrkB signaling pathways. Here, we
demonstrate that VRT enhances NOG with a hormetic concentration-response
relationship, and this response is dependent on TrkB receptor signaling. We also show
that acute exposure to VRT caused increases in [Na+]i and [Ca2+]i with the [Na+]i
increment being sufficient to upregulate NMDAR function. Further, inhibition of TrkB
receptors and its downstream signaling molecules, PI3K, and PLC inhibited VRT-
enhanced NOG. Acute treatment with VRT stimulated phosphorylation of TrkB and its
4-126
downstream effectors Akt, mTOR, PLC, ERK1/2 and CREB. More importantly, VRT
increased BDNF synthesis and release in a concentration dependent manner. Veratridine
stimulation of TrkB phosphorylation displayed a biphasic concentration-response curve.
Taken together, these data suggest that VRT activates VGSCs and elevates [Na+]i , which
in turn augments NMDAR function leading to increased BDNF synthesis and release.
Released BDNF binds to the TrkB receptor causing activation of BDNF-TrkB receptor
signaling, thereby mediating VGSC activator-enhanced neurite outgrowth. These data
provide further support for the hypothesis that sodium channel activators are capable of
mimicking activity-dependent neuronal development through potentiation of NMDAR
and neurotrophin signaling pathways.
4.3 Materials and Methods
Cerebrocortical Neuron Culture.
Primary cultures of cerebrocortical neurons were harvested from Swiss Webster mice on
embryonic day 16 and cultured as described previously (Cao et al., 2008). Cells were
plated onto poly-l-lysine-coated (Sigma-Aldrich, St. Louis, MO) 96-well (9 mm), clear-
bottomed, black-well culture plates (Corning Life Sciences, Lowell, MA) at a density of
1.8 × 106 cells/ml (150 μl/well), 24-well (15.6 mm) culture plates at a density of 0.05 ×
106 cells/ml (0.5 ml/well), 12-well (22 mm) culture lates at a density of 1.8 × 106 cells/ml
(1.0 ml/well), or 6-well (35 mm) culture dishes at a density of 2.25 × 106 cells/ml (2
ml/well), respectively, and incubated at 37°C in a 5% CO2 and 95% humid atmosphere.
4-127
All animal use protocols were approved by the Creighton University Institutional Animal
Care and Use Committee.
Determination of Total Neurite Length and Diolistic Labeling.
Cells were plated on poly-lysine-coated 12- or 15-mm glass coverslips (Thermo Fisher
Scientific, Waltham, MA) and placed inside of 24-well culture plates at a low density of
0.05 × 106 cells/ml (0.5 ml/well). To assess the influence of VRT on neuritogenesis,
primary cultures of immature cerebrocortical neurons were exposed to various
concentrations of VRT ranging from 10 to 10000 nM for 24 h beginning 3 h after plating,
and total neurite outgrowth was measured. In some experiments, these concentrations of
VRT were coincubated with MK-801 (1 μM) (Sigma-Aldrich), LY29002 (10 μM)
(Sigma-Aldrich), U37122 (2 μM) (Calbiochem)K-252a (200 nM) (Calbiochem, San
Diego, CA), U0126 (10 μM) (Calbiochem) Rapamycin (1 μM) (Calbiochem). At 24 h
after plating, cultures were fixed at room temperature for 20 min using 1.5%
paraformaldehyde in phosphate-buffered saline (PBS). After fixation, neurons were
diolistically labeled with DiI. The Helios Gene Gun System (Bio-Rad Laboratories,
Hercules, CA) was used to deliver DiI-coated tungsten particles (1.3 μM) (Bio-Rad
Laboratories) into paraformaldehyde-fixed cerebrocortical neurons 1 day in vitro (DIV).
Diolistic bullet preparation was based on the method of O’Brien and Lummis (2006). In
brief, 2.5 to 3.5 mg of DiI (Invitrogen, Carlsbad, CA) was suspended in 200 μl of
dichloromethane (Sigma-Aldrich). The dissolved dye was added over evenly spread
tungsten particles (35 mg) placed on a clean glass slide and then allowed to dry. The dye-
coated particles were scraped onto another clean glass slide and chopped to fine particles
using a clean razor blade and later resuspended in 3 ml of deionized water. This dye
4-128
slurry was sonicated for 10 min and then vortex briefly to form a uniform suspension.
After adding 100 μl of polyvinylpyrrolidone (PVP) (Bio-Rad Laboratories) stock solution
(0.96% PVP in ethanol) to the dye slurry, it was drawn into a PVP-precoated Tefzel
tubing mounted on a preparation station (Bio-Rad Laboratories) using a 5- to 10-ml
syringe. The dye particles were allowed to settle for 20 to 30 min, and then the
supernatant water was carefully withdrawn from Tefzel tubing (Bio-Rad Laboratories)
using a syringe. The tubing was rotated for 1 to 2 min to uniformly spread the particles.
The tubing was then allowed to dry for 5 min before cutting into bullets using a tube
cutter. The DIV-1 cerebrocortical neurons grown on coverslips were shot postfixation
(1.5% paraformaldehyde) using DiI bullets loaded onto a Helios gene gun at 140 to 160
psi of helium pressure from a distance of 2.5 cm. The dye particles were allowed to
spread across the neuronal membrane overnight, and coverslips were then mounted for
imaging on an Olympus IX 71 inverted microscope with a Himamatsu camera. Digital
images of individual neurons were captured, and total neurite length was quantified. To
reduce the effect of paracrine neurotrophic factors on neurite growth, only those neurons
that were separated from surrounding cells by approximately 150 μm were digitally
acquired and analyzed. Digital images of individual neurons were captured and exported
as 16-bit images. All neurites in a single neuron including those from secondary branches
were semi-automatically traced, and the length was measured by using the using
FilamentTracer module of Imaris 6.4.0 software. At least 25 randomly chosen neurons
from two different cultures were evaluated for each treatment group.
Intracellular Sodium Concentration Measurement.
4-129
[Na+]i measurement and full in situ calibration of sodium-binding benzofuran
isophthalate (SBFI) fluorescence ratio were performed as described previously (Cao et
al., 2008). Cells grown in 96-well plates were washed four times with Locke's buffer (8.6
mM HEPES, 5.6 mM KCl, 154 mM NaCl, 5.6 mM glucose, 1.0 mM MgCl2, 2.3 mM
CaCl2, 0.1 mM glycine, pH 7.4) using an automated microplate washer (BioTek
Instruments, Winooski, VT). After measuring the background fluorescence of each well,
cells were incubated for 1 h at 37°C with dye-loading buffer (100 μl/well) containing 10
μM SBFI-AM (Invitrogen) and 0.02% Pluronic F-127 (Invitrogen). Cells were then
washed five times with Locke's buffer, leaving a final volume of 120 μl in each well. The
plate was then transferred back to the incubator for 15 min to allow the cells to
equilibrate after washing and then placed in a FlexStation II (Molecular Devices,
Sunnyvale, CA) chamber to detect Na+-bound SBFI emission at 505 nm (cells were
excited at 340 and 380 nm). Fluorescence readings were taken once every 5 s for 60 s to
establish the baseline, and then 40 μl of VRT was added to each well from the compound
plate at a rate of 26 μl/s, yielding a final volume of 160 μl/well. After correcting for
background fluorescence, SBFI fluorescence ratios (340/380) versus time were analyzed,
and time- or concentration-response graphs were generated using GraphPad Prism
(GraphPad Software Inc., San Diego, CA). Full in situ calibration of the SBFI
fluorescence ratio was performed as described previously (Cao et al., 2008, Jabba et al.,
2010)
Intracellular Ca2+ Monitoring.
DIV-1 cerebrocortical neurons grown in 96-well plates were used for intracellular Ca2+
concentration ([Ca2+]i) measurements as described previously (George et al., 2009, Cao et
4-130
al., 2008, Jabba et al., 2010. In brief, the growth medium was removed and replaced with
dye-loading medium (100 μl/well) containing 8 μM fluo-3 AM (Invitrogen) and 0.04%
Pluronic acid in Locke's buffer. After 1-h incubation in dye-loading medium, the neurons
were washed four times in fresh Locke's buffer (200 μl/well) using an automated
microplate washer (BioTek Instruments) and transferred to a FlexStation II benchtop
scanning fluorometer chamber. The final volume of Locke's buffer in each well was 120
μl. Fluorescence measurements were performed at 37°C. The neurons were excited at 488
nm, and Ca2+-bound fluo-3 emission was recorded at 538 nm at 2-s intervals. After
recording baseline fluorescence for 27 s, 40 μl of a 4× concentration of VRT in the
presence or absence of specific agonists like TTX or MK-801 were added to the cells at a
rate of 26 μl/s, yielding a final volume of 160 μl/well; the fluorescence was monitored for
an additional 220 to 270 s. The fluo-3 fluorescence was expressed as (Fmax – F0)/F0,
where Fmax is the maximum and F0 is the fluorescence measured in each well at time
zero. In some experiments the fluo-3 fluorescence was expressed as area under the curve
(AUC).
Western Blotting.
Western blot analysis was performed by using cells grown in either six-well or 12-well
plates. DIV-1 cells were exposed to 1000 nM VRT for various time periods at 37°C. For
pharmacological experiments, along with 1000 nM VRT, cultures were co-incubated
either in the presence or absence of specific antagonists. At the end of each time period,
cultures were transferred onto an ice slurry to terminate drug exposure and washed three
times with ice-cold PBS. Cells were lysed using ice-cold lysis buffer (50 mM Tris, 50
mM NaCl, 2 mM EDTA, 2 mM EGTA, 1% Nonidet P40, 0.1% SDS, 2.5 mM sodium
4-131
pyrophosphate, and 1 mM sodium orthovanadate). Phenylmethylsulfonyl fluoride (1
mM) and 1× protease inhibitor mixture (Sigma-Aldrich) were then added, and the lysate
was incubated for 30 min at 4°C. Cell lysates were sonicated and then centrifuged at
13,000g for 15 min at 4°C. The supernatant was assayed by the Bradford method
(Bradford, 1976) to determine protein content. Equal amounts of protein were mixed with
the Laemmli sample buffer and heated for 5 min at 75°C. The samples were loaded onto
a 10% SDS-polyacrylamide gel electrophoresis gel and transferred to a nitrocellulose
membrane and immunoblotted with specific antibodies. Blots were developed with ECL
Plus kit (GE Healthcare, Chalfont St. Giles, UK) for 3 min. Blots were subsequently
stripped (63 mM Tris base, 70 mM SDS, 0.0007% 2-mercaptoethanol, pH 6.8) and
reprobed for further use. Western blot densitometry data were obtained by using MCID
Basic 7.0 software (Imaging Research, St. Catharines, ON, Canada).
4.3.1.1 BDNF immunoassays.
Sandwich BDNF ELISA and BDNF ELISA in situ (Balkoweic and Katz, 2000
J.Neurosci.) were performed to measure BDNF protein using a BDNF Emax immunoassay
System (Promega, Madison, WI) according to manufacturer specifications, except that
the concentrations of the anti-BDNF monoclonal antibody and anti-human BDNF
polyclonal antibody were two-fold of the recommended concentrations for BDNF ELISA
in situ. Also, the dilution of the anti IgY-HRP antibody was 1:50 in BDNF ELISA in situ.
Sandwich BDNF ELISA: Protein lysates were made from primary neuronal cultures
grown in 6 or 12-well plates. Cultures were treated with 1000 nM VRT for specific times,
either in presence or absence of various antagonists and changes in total BDNF protein
was measured by sandwich BDNF ELISA. Breifly, 96-well ELISA plates (Nunc
4-132
maxisorp) were incubated overnight at 40C with anti-BDNF monoclonal antibody diluted
in carbonate coating buffer. The plates were washed 1X with TBST and blocked for 1h at
RT. Protein lysates were applied along with BDNF standards and incubated with shaking
(400 rpm) at RT for 2h. Subsequently plates were washed 5X and anti-human BDNF
polyclonal antibody was added and incubated with shaking at RT for 2h. After washing
5X, anti IgY-HRP antibody was added and with shaking at RT for 1h. The plate is
washed 5X and incubated with TMB solution for color development. After 10 minutes
reactions are stopped using1N HCl. Absorbance values were measured at 450 nm in a
plate reader (name the instrument).
BDNF ELISA in situ: This method was performed according to Balkoweic and Katz,
2000 J.Neurosci. Breifly, 96-well ELISA plates (Nunc maxisorp) were UV-sterilized for
30 minutes and then incubated overnight at 40C with anti-BDNF monoclonal antibody
diluted in carbonate coating buffer. Next, plates were blocked for 1h and then washed 2X
with plating media. Dissociated immature cerebrocortical neurons were plated and
cultured for 4 days. 3h post plating cells were treated to various concentrations of VRT
and 50 mM KCl. BDNF standards were also added to the plate at the start of the culture.
After 3-4d, the plates were washed vigorously with TBST to remove the cells and debris.
Subsequently anti-human BDNF polyclonal antibody was added and incubated with
shaking at RT for 2h. The remaining of the procedure was similar to that is described
above.
Plasmids and Nucleofection: FL-TrkB and DN TrkB (truncated TrkB) in pBluescript
sk-/- vector were a generous gift from Tony Hunter (Salk Institute, San Diego).
Dissociated cortical neurons obtained from E16-17 pups were re-suspended in
4-133
Nucleofector solution (Mouse Neuron Kit, Amaxa Biosystems) and transfected according
to the manufacturer's directions with plasmids containing the genes of interest, using an
Amaxa Nucleofector (program O-005). Five million and 1 million neurons were used per
reaction for western blot and neurite outgrowth experiments, respectively. One
microgram of plasmid containing the gene of interest was used per nucleofection
reaction. Control neurons were transfected with the empty vector. Also, all nucleofection
reactions contained GFP plasmid (1 µg) bringing the total amount of plasmids to 2 µg per
reaction. Transfected neurons were plated at 2 X 106 neurons/well and 0.5 X 105
neurons/well for western blot experiments and neurite outgrowth assays, respectively. In
order to give more time for the expression of genes of interest, DIV-2 neurons were
utilized in experiments involving nucleofection.
4.4 Results
Veratridine Enhances Neurite Outgrowth in Immature Cerebrocortical Neurons.
In previous reports, we demonstrated that the VGSC activators brevetoxin (PbTx-2) and
antillatoxin (ATX) stimulated neurite out growth. Here, we wanted to determine whether
these transactions generalized to the VGSC site 2ligand, veratridine(VRT) . Three hours
after plating, primary cultures of immature cerebrocortical neurons were exposed to
various concentrations of VRT ranging from 10 to 10,000 nM for 24 h, and total neurite
outgrowth and dendritic branch points were assessed. Diolistic labeling of DiI was used
to visualize neurons and determine the influence of VRT on neurite outgrowth (Fig. 1A)
and dendritic branch points. Veratridine significantly enhanced total neurite outgrowth in
immature cerebrocortical neurons with concentrations of 300 and 1,000 nM producing a
4-134
robust >2-fold increase in total neurite length (***, p < 0.001) (Fig 1B). As previously
observed with PbTx-2 and ATX, the VRT concentration-response profile was
bidirectional, or hormetic (Fig 1B). Veratridine also significantly enhanced dendritic
branch points in these neurons with concentrations of 300 and 1,000 nM producing a
robust 4- (***, p < 0.001) and 3-fold (**, p < 0.01) increases respectively.
Veratridine Increases Intracellular sodium ([Na+]i) Levels in Immature Cerebrocortical
Neurons.
Given that the earlier studies (Yu et al., 1997, Yu and Salter 1998, Yu et al 2006, George
et al 2009) demonstrated that [Na+]i is a regulator of NMDAR-mediated signaling, it was
important to quantify the magnitude of VRT-induced elevation of [Na+]i in immature
cerebrocortical neurons. SBFI, a sodium-sensitive fluorescent indicator, was used to
determine the influence of VRT on [Na+]i in DIV-1 cerebrocortical neurons. Full in situ
calibration was performed in DIV-1 cerebrocortical neurons to determine the relationship
between the ratiometric SBFI fluorescence signal and [Na+]i (Cao et al 2008, George et al
2009, Jabba et al 2010). VRT produced a concentration-dependent increase in [Na+]i (Fig
2 A,B) with an EC50 value of 3.12 µM (1.42-6.88 µM, 95% CI). The in situ SBFI
calibration showed that basal [Na+]i in DIV-1 cerebrocortical neurons was 16.1 ± 0.51
mM. Inasmuch as the VRT concentrations of 300 and 1,000 nM were sufficient to
produce a robust increase in neurite outgrowth, it was important to quantify the [Na+]i
increment associated with these treatments. The 300 nM VRT treatment produced a
maximal [Na+]i of 23.1± 0.3 mM and 1,000 nM produced 26.5 ± 0.36 mM, representing
increments of 7.0 and 10.4 mM over basal. Previous reports in hippocampal neurons
suggested that an increment of [Na+]i of 10 mM was sufficient to produce significant
4-135
increases in NMDAR channel activity (Yi and Salter 1998, Yan et al 2006). Moreover, it
has been reported that increments of [Na+]i >5 mM represent a critical threshold required
to regulate NMDAR-mediated Ca2+ influx in primary cultures of hippocampal neurons
(Xin et al., 2005). Consistent with these findings, the increment of [Na+]i detected in
immature cerebrocortical neurons appears sufficient to up-regulate NMDAR function.
Veratridine Increases Intracellular Calcium Levels ([Ca2+]i) in DIV-1 Cerebrocortical Neurons.
Activity-dependent neuritogenesis and neuronal development involve Ca2+-influx and to
a large extent this Ca2+-influx is NMDAR dependent. Previous studies have also
suggested that sodium channel activator-induced neurite outgrowth and Ca2+-influx
involves NMDARs and VGSCs. Hence, we hypothesized that VRT exposure would
produce Ca2+ influx in these immature cerebrocortical neurons. To investigate this theory,
cells loaded with fluo-3 were exposed to various concentrations of VRT, and [Ca2+]i was
monitored. VRT produced rapid and concentration-dependent increases in [Ca2+]i, with
an EC50 value of 2.56 µM (1.19-16.56 µM, 95% CI).
To investigate the role of VGSCs and NMDARs in VRT-induced Ca2+-influx in DIV-1
cerebrocortical neurons, a pharmacological evaluation was performed using TTX
(VGSCs) or MK-801 (NMDARs), prior to 1,000 nM VRT exposure and changes in
[Ca2+]i were monitored. TTX (1 μM) completely blocked the response to VRT (Fig 2E,
F), whereas MK-801 (1 μM) significantly reduced VRT-induced Ca2+ influx (**, p <
0.01) (Fig 2G, H).
4-136
Veratridine-induced neurite outgrowth involves TrkB receptors.
Inasmuch as previous studies have indicated that activity-dependent neuritogenesis and
neuronal development involve Ca2+ influx through NMDAR with subsequent
engagement of BDNF-TrkB signaling, we assessed the role of TrkB receptor in sodium
channel activator-induced neurite outgrowth. Coincubation of K-252a (200 nM), a TrkB
inhibitor, with 300 nM VRT inhibited VRT-stimulated neurite outgrowth in immature
cerebrocortical neurons (***, p < 0.001) (Fig. 3A, B and C), demonstrating the
requirement for TrkB receptor activation. These pharmacological data were confirmed
with a genetic approach in which DIV-1 neurons were transfected with either dominant
negative isoform of TrkB (truncated TrkB), the full length isoform of TrkB (TrkB.FL) or
the empty vector (back bone). Consistent with the involvement of TrkB receptors,
dominant negative TrkB completely abolished VRT-induced neurite outgrowth, whereas
neurons expressing TrkB.FL showed robust increase in neurite length similar to those
expressing empty back bone vector when exposed to VRT (Fig.3D and E).
Veratridine enhances BDNF release and synthesis in immature cerebrocortical neurons
Neurotrophins are expressed and released from neurons in an activity-dependent manner
and act in an autocrine/paracrine mode to induce morphological and functional changes
in neurons. Activation of NMDARs and elevation of [Ca2+]i are critical for the expression
and release of neurotrophins. Inasmuch as sodium channel activators mimic activity-
dependent neuronal development with attendant stimulation of NMDARs and increase in
intracellular Ca2+, we predicted that sodium channel activation will increase the synthesis
and release of BDNF with attendent activation of TrkB receptors. The influence of VRT
on release of endogenous BDNF was quantified using an in situ BDNF ELISA (Figure
4-137
4A). Three hour post-plating, neurons were exposed to various concentrations of VRT
ranging from 100 to 10,000 nM for 72 h. As a positive control, neurons were exposed to
50 mM KCl. VRT produced a concentration-dependent increase in BDNF release with 10
µM VRT producing comparable increases in BDNF release to that of 50 mM KCl (Figure
4 B). VRT in concentrations of 1 (*, p < 0.05), 3 (**, p < 0.01) and 10 µM (***, p <
0.001) produced significant increases in BDNF release compared to control. We further
investigated BDNF synthesis in immature cerebrocortical neurons following VRT
exposure. Neurons were exposed to 1,000 nM VRT and cell lysates were collected for
various time points starting at 3 h post plating to 24 h (corresponding to the time of
neurite outgrowth assay). Results revealed that 1,000 nM VRT exposure increased BDNF
synthesis at 1, 6 and 12 h of exposure but showed reduced amount of BDNF after 24 h
VRT exposure. (Figure 5A, B). We also characterized for the increase in BDNF synthesis
on VRT exposure using BDNF sandwich ELISA (Figure 5C). Three hours post plating
neurons were exposed to 1000 nM VRT either in the presence or absence of various
specific antagonists for 12 h. Cell lysates (using ELISA lysate buffer) were collected and
stored for BDNF sandwich ELISA assay. These results revealed that TTX, a VGSC
blocker, completely inhibited VRT-enhanced BDNF synthesis. BDNF synthesis was also
partially blocked (75 %) by MK 801, a NMDAR blocker. The VGCC blocker nifedipine
(1 µM) and PLCγ blocker U73122 were without effect on VRT-stimulated BDNF
synthesis. These results demonstrate that VRT enhances BDNF synthesis in immature
cerebrocortical neurons and this requires VGSC and NMDARs activation (Figure 5C).
We also demonstrated using BDNF sandwich ELISA assay and western blot analysis,
4-138
that VRT increased BDNF synthesis in a concentration- dependent manner.(Figure 6A,
B, C)
Veratridine-induced neurite outgrowth involves PI3-kinase activity
The PI3K-Akt signaling pathway is downstream of BDNF-TrkB receptor and plays an
important in neurotrophin-stimulated neurite outgrowth. Inasmuch as VRT stimulated the
synthesis and release of BDNF, we determined whether PI3K is involved in VRT-
induced neurite outgrowth. Coincubation of LY29002 (10 µM), a PI3K inhibitor, with
300 nM VRT completely inhibited VRT-stimulated neurite outgrowth in immature
cerebrocortical neurons (***, p < 0.001) (Figure 7 A, B and C), suggesting a PI3K
involvement in VRT-enhanced neurite outgrowth. Similarly, wortmanin, another PI3K
blocker, also inhibited VRT-stimulated neurite outgrowth (data not shown). The major
affecter downstream of PI3K that modulates neurite outgrowth is Akt. To assess the
ability of VRT to activate Akt, we determined the phosphorylation of the Ser473 residue
on Akt by using an anti-phospho-Ser473 Akt antibody. DIV-1 cerebrocortical neurons
were exposed to 1,000 nM VRT, and cell lysates were collected at various time periods
for western blot analysis. The results revealed that 1,000 nM VRT produced a robust
activation of Akt as reflected by the increase in the phosphorylation of Ser473 at 5
minutes after exposure (~2 fold). Akt activation peaked at 10-15 minutes of VRT
exposure (~3 fold) (Figure 7 C, D). Prior incubation with VGSC blocker TTX for 15
minutes attenuated VRT-stimulated Akt activation indicating the requirement for VGSC
activation (Figure 7 E,F). These findings establish a role for PI3K-Akt signaling in the
stimulatory effect of VRT on neurite outgrowth.
4-139
Veratridine stimulated Akt phosphorylation involves TrkB receptors
To determine the involvement of TrkB receptors in Veratridine stimulated Akt
phosphorylation, DIV-1 cultures were \ pre-incubated with K252a (200 nM) for 30
minutes and then exposed to 1,000 nM VRT for 5 minutes. Cell lysates were collected
and investigated for activation of Akt (Ser473) by immunoblotting. Western blot analysis
revealed that K252a blockade completely inhibited VRT-stimulated Akt activation
indicating the requirement of TrkB receptors (Figure 8C, D). To further confirm these
pharmacologic results, we utilized a genetic approach by nucleofecting the neurons with
either DN-TrkB or FL-TrkB. Neurons nucleofected with an empty back-bone vector were
used as a control. All nucleofected neurons were exposed to 1,000 nM VRT for 5
minutes. Subsequently cell lysates were collected and investigated for activation of Akt
(Ser473) by immunoblotting. Nucleofection with DN TrkB completely blocked the VRT-
stimulated Akt activation, but neither the FL-TrkB (~4 fold) nor the empty vector (~6
fold) had any effect on VRT-stimulated Akt activation (Figure 8A,B). The basal Akt
activity was also higher in neurons nucleofected with FL TrkB. These data suggest the
involvement of TrkB receptors in Veratridine stimulated Akt phosphorylation.
Veratridine-induced neurite outgrowth involves the PI3K-Akt-mTOR pathway
Downstream of TrkB is the mTOR signaling complex which is critical for protein
synthesis in dendrites and participates in activity–dependent dendritic arborization. PI3K-
Akt signaling acting through, or in coordination with mTOR, is involved in the regulation
of dendritic morphogenesis and synaptic plasticity. To determine the involvement of
mTOR signaling in VRT-stimulated neurite outgrowth, cerebrocortical neurons were co-
incubated with VRT in the presence or absence of rapamycin, specific inhibitor of
4-140
mTOR, for 24 h and total neurite length was determined. Consistent with the involvement
of mTOR, rapamycin inhibited VRT-stimulated neurite outgrowth (Figure 9 A, B, C). To
assess the ability of VRT to activate mTOR, we determined the phosphorylation of the
Ser2448 residue on mTOR using an anti-phospho-Ser2488 mTOR antibody. DIV-1
cerebrocortical neurons were exposed to 1,000 nM VRT, and cell lysates were collected
at various time periods for western blot analysis. The results revealed that 1,000 nM VRT
produced a ~2 fold activation of mTOR as reflected by an increase in the phosphorylation
of Ser2448 at 15 minutes post exposure. (Figure 9 D). Taken together, these results
indicate that mTOR activation participates in sodium channel activator-stimulated
neuronal neurite outgrowth.
Veratridine-induced Ca2+ influx and neurite outgrowth involves PLC mediated
release of Ca2+ from intracellular stores
Another important signaling pathway downstream of neurotrophin-TrkB pathway is the
PLCγ pathway. BDNF activation of TrkB receptors at Y816 recruits and activates PLCγ
which in turn hydrolyzes PIP2 to DAG and IP3, with IP3 triggering the release of Ca2+
from intracellular stores. Release of Ca2+ from intracellular calcium stores leads to release
of secretory growth factors vesicles. Here we explored the involvement of PLCγ in VRT-
induced changes in [Ca2+]i using the PLCγ inhibitor U73102. We also determined for the
role of PLCγ in VRT- induced neurite outgrowth pharmacologically using U73102. Fluo-
3 loaded DIV-1 neurons were pretreated with the PLCγ specific antagonist-U73102, prior
to 1,000 nM VRT exposure and changes in [Ca2+]i were monitored. U73102 (2 μM)
significantly reduced VRT-induced Ca2+ influx (**, p < 0.01) (Figure 10 A, B). We then
determined whether the attenuation in Ca2+ response was due to inhibition of release of
4-141
Ca2+ from intracellular Ca2+ stores. We pretreated neurons with 10 μM thapsigargin for
40 minutes to deplete the ER of calcium and then treated with 1,000 nM VRT. VRT-
induced change in intracellular was significantly attenuated in presence of thapsigargin,
suggesting that PLC mediated release of Ca2+ from intracellular stores contributes to
VRT-induced elevation of Ca2+ (Figure 10 C, D). Next, we investigated whether PLC is
involved in VRT-induced neurite outgrowth. Coincubation of U73102 (2 µM) with 300
nM VRT completely inhibited VRT-stimulated neurite outgrowth in immature
cerebrocortical neurons (***, p < 0.001) (Figure 11 A), indicating that VRT-enhanced
neurite outgrowth involves PLC signaling. We next determined whether phosphorylation
of the Y816 residue on TrkB following VRT exposure was involved in the recruitment of
downstream PLCγ signaling machinery to TrkB receptor. DIV-1 cerebrocortical neurons
were exposed to 1,000 nM VRT, and cell lysates were collected at various time periods
for western blot analysis. The results revealed that 1,000 nM VRT produced a robust
activation of TrkB Y816 residue as reflected by the increase in the phosphorylation at 2
minutes post exposure. (Figure 11 B) We also examined the ability of VRT to activate
PLCγ by determining the phophorylation of Y783 residue on PLCγ1. We found the Y783
residue on PLCγ1 is activated by VRT exposure at 2-5 minutes. (Figure 11 C). Taken
together these data indicate the involvement of a TrkB- PLCγ signaling in VRT-
stimulated neurite outgrowth.
MAPK pathway has a modest role in veratridine-induced neurite outgrowth.
The third signaling pathway downstream of neurotrophin-TrkB pathway is the MAPK
pathway. BDNF phosphorylation of the Tyr515 site on TrkB receptors activates
downstream MEK-MAPK/Erk signaling, which promotes neuronal differentiation and
4-142
growth. We investigated for involvement of the MAPK pathway in VRT-induced neurite
outgrowth using the MEK inhibitor U0126 (20 µM). Coincubation of U0126 with 300
nM VRT partially inhibited VRT-stimulated neurite outgrowth in immature
cerebrocortical neurons (*, p < 0.05) (Fig.12 A, B and C), indicating that VRT-enhanced
neurite outgrowth partially involves MEK-MAPK pathway. We further demonstrated
phosphorylation of ERK1/2 (T202/Y204) and CREB (S133) following VRT exposure.
DIV-1 cerebrocortical neurons were exposed to 1,000 nM VRT, and cell lysates were
collected at various time periods for western blot analysis. The results revealed that 1,000
nM VRT produced a robust activation of ERK1/2 ((T202/Y204) and CREB (S133) in
immature cerebrocortical neurons. (Figure 12 A, B and C).
4.5 Discussions
The electrical signals of neurons are fundamentally dependent on Na+ influx through
VGSCs. Sodium channels are primarily responsible for the rising phase of action
potential and hence supply the current that drives the membrane potential to peak
depolarization (Hille B, 2001). VGSCs activity has been shown to regulate
neurotransmitter release in developing cortex and also to mediate neuronal firing
dependent synaptic plasticity (Platel JC, 2005, Cantrell AR, 2001). In immature
cerebrocortical neurons, VGSCs activators enhanced neurite outgrowth through
potentiation of NMDAR signaling pathways that influence neuronal morphology 296
Jabba,S.V. 2010; 6 George,J. 2009, indicating that sodium channel activators appear
capable of mimicking activity-dependent neuronal development. Neuronal activity also
cooperates with neurotrophins to influence neuronal development and plasticity 1
Wong,R.O. 2002; 262 Yoshii,A. 2010; 267 Reichardt,L.F. 2006. Although activity-
4-143
dependent neurotrophins influences on neuronal development are well studied, little is
known regarding the influence of VGSCs activation on neurotrophin signaling. Towards
achieving this goal, we used a VGSC gating modifier and determined the effects of
VGSCs activation on BDNF synthesis, secretion and BDNF receptor (TrkB) dependent
signaling pathways, and their role in neurite outgrowth. Our findings provide one of the
first reports of the influence a VGSC gating modifier on neurotrophin signaling. We
found that VGSCs activator increased BDNF release, synthesis and activated BDNF-
TrkB signaling pathways, including the PI3K-Akt cascasde, the PLCγ pathway, and the
MAPK/Erk1/2 pathway. An interesting feature of our results is that the influence of VRT
on TrkB activation followed a bidirectional pattern similar to that of VRTon neurite
outgrowth.
VRT enhanced total neurite outgrowth and dendritic arborization in immature
cerebrocortical neurons with a bidirectional, or hormetic, concentration-response
relationship with 300-1,000 nM producing robust increases of more than 2-fold. (Figure
1). This result is in accordance with previous reports using such as other VGSCs
activators such as PbTx-2 and ATX 6 George,J. 2009; 296 Jabba,S.V. 2010. PbTx-2
and ATX enhanced neurite outgrowth in DIV-1 cerebrocortical neurons as a result of
elevation of cytoplasmic [Na+], potentiation of NMDAR function and stimulation of
calcium influx. Yu and Salter 96 Yu,X.M. 1998first suggested that [Na+]i may act as
signaling molecule and augments NMDAR fuction. They used VRT to demonstrate that
Na+ influx through TTX-sensitive VGSC was sufficient to upregulate NMDAR activity
in hippocampal neurons. Similarly, previous reports 296 Jabba,S.V. 2010; 6 George,J.
2009 using single-channel current recordings from cell-attached patches
4-144
unambiguously confirmed that VGSCs activators like PbTx-2, ATX augmented NMDAR
function by increasing open probability or mean open time (or both) of NMDARs. Yu
and Salter have also determined that an increment in [Na+]i of 10 mM was sufficient to
potentiate NMDAR single-channel activity. Moreover, this Na+-dependent regulation of
NMDA receptor function was shown to be controlled by Src-induced phosphorylation of
the receptor. These results were extended in the present study using sodium channel
activator VRT as a probe to elevate intracellular Na+ in cerebrocortical neurons. VRT
treatment increased the release of glutamate and enhanced NMDA-induced Ca2+ influx in
immature cerebrocortical neurons (data not shown). Veratridine exposure also led to the
activation of Src (data not shown) 95 Yu,X.M. 1997. We have previously shown that
an increase in intracellular Na+ and Src activation following exposure to ATX increased
the open probability of the NMDAR. These data, therefore, confirm the regulatory
influence of Na+ on NMDAR channel activity in hippocampal neurons described
previously 96 Yu,X.M. 1998 and extend this relationship between [Na+]i and NMDA
receptor function to cerebrocortical neurons.
NMDAR activation promotes neurite growth and dendritic arborization, whereas
pharmacological blockade of NMDARs reduces it 43 Rajan,I. 1998. Stimulation of
NMDAR function also activates Ca2+ signaling pathways that regulate neurite outgrowth
and dendritic arborization 38 Konur,S. 2005. Consistent with these reports, we found
that exposure of immature cerebrocortical neurons to VRT increased [Ca2+]i with
significant dependence on NMDARs. Thus, the ability of VRT to augment NMDAR
channel activity translated into an enhancement of the trophic influence of NMDAR on
developing cerebrocortical neurons. Based on the premise that the effects of neuronal
4-145
activity on dendritic arbor growth and structural plasticity are primarily mediated by
engagement of NMDA receptors 106 Tolias,K.F. 2005, our results suggest that VRT
activation of sodium channels with attendant enhancement of NMDA receptor signaling
mimics the response to neuronal activity.
Another important finding of the study is that a VGSC activator increased BDNF
synthesis, release and activated BDNF-TrkB signaling pathways. BDNF plays an
important role in regulating neural survival, development, function and plasticity.
Veratridine-enhanced BDNF synthesis was TTX-sensitive, and pharmacological
blockade of NMDARs significantly attenuated the response. Nifedipine and U73122, a
VGCC and PLC inhibitor, respectively, were without effect on VRT-enhanced BDNF
synthesis (Figure 5). These data, therefore demonstrate that VGCCs and intracellular
Ca2+ stores are not required for VRT-induced BDNF synthesis. Transcription of Bdnf
from its promoters (I & IV) is highly regulated by neuronal activity-dependent Ca2+
influx 321 Kidane,A.H. 2009; 332 Rattiner,L.M. 2005, and is regulated by the route
of Ca2+ influx into the cell and by the pattern of phosphorylation induced on the
transcription factor CREB 446 West,A.E. 2001. In rat cerebrocortical cultures,
Ghosh et al., 445 Ghosh,A. 1994 demonstrated that transcription of Bdnf is
preferentially driven by Ca2+ influx through L-VGCCs, whereas it is poorly induced by
calcium coming through NMDARs. In that study they demonstrated that glutamate
increased Bdnf gene expression transiently, with peak expression around 1 h and with
reduced expression thereafter. The stimulation of Bdnf gene expression by depolarizing
levels of KCl was more delayed and peaked at approximately 3 h with a subsequent
sustained elevation. This difference could be due to inability of VRT to alter membrane
4-146
potential in cerebrocortical neurons, which would not provide the stimulus for VGCC
activation. Also, similar to glutamate increased Bdnf gene expression, VRT increased
BDNF protein expression peaked at approximately 1 h and was sustained for 12 h.
Exposure to VRT in immature cerebrocortical neurons moreover increased BDNF release
(secretion) in a concentration-dependent manner (0.1-10 µM VRT). The released BDNF
binds to its cognate receptor TrkB in an autocrine/paracrine fashion to activate
downstream signaling pathways and thereby influence neuronal development. Similarly,
a 12 h VRT exposure increased BDNF synthesis in a concentration-dependent manner
(0.1-10 µM VRT). Interestingly, the influence of VRT on TrkB activation (Y816)
followed a bidirectional concentration-relationship, similar to that of VRT-mediated
neurite outgrowth. Marini et al., 450 Marini,A.M. 1998 have demonstrated that
NMDA exertes a neuroprotective activity by increasing BDNF release (acute effect, 2-5
‘) and synthesis (chronic effect, ~3 h), leading to activation of TrkB signaling. They
proposed that there is an integral relationship between NMDAR activation and BDNF-
TrkB signaling in a bidirectional manner, consonant with the bidirectional pattern shown
by NMDA and VRT on neurite outgrowth. An inverted-U model describes the
relationship between NMDA receptor activity and neuronal survival and growth 80
Lipton,S.A. 1999. In both cases, this inverted-U concentration–response relationship
has primarily, but not exclusively, been attributed to [Ca2+]i regulation. An optimal
window for [Ca2+]i is required for activity-dependent neurite extension and branching,
with lower levels stabilizing growth cones and higher levels stalling them, in both cases
preventing extension 50 Gomez,T.M. 2000; 74 Hui,K. 2007. In regard to NMDAR-
BDNF-TrkB signaling, one possible explaination could be that , the TrkB receptor may
4-147
be desensitized at higher concentrations of VRT exposure, due to higher amounts of
BDNF being synthesized and released. Conversely, at lower concentrations of VRT
exposure the neurons are not able to synthesize and release sufficient BDNF to activate
TrkB receptors. The trophic role of BDNF in promoting serotonergic neuron axonal
growth and remodeling in the adult brain also displayed an inverted U-concentration-
response relationship 447 Mamounas,L.A. 2000 . In this study, cortical infusions of
various BDNF concentrations activated TrkB signaling locally, leading to highly
localized seratonin sprouting response, in which both the seratonin sprouting and Trk
receptor signaling displayed maximal responses at intermediate BDNF doses and are
depressed at higher doses. BDNF is a key mediator of bidirectional responses to exercise
and anti-depressants 448 Gomez-Pinilla,F. 2008. Although higher levels of BDNF
are generally beneficial to neuronal survival, excessive activation of its receptor, TrkB
can have adverse affects on neuronal survival and plasticity.
Sodium channels gating modifiers such as VRT appear capable of mimicking
neuronal activity and neurotrophin-dependent neurite outgrowth and hence may represent
a novel pharmacological strategy to regulate neuronal development through NMDAR-
BDNF-TrkB-dependent mechanisms. Recent studies have shown that neurotrophin
signaling plays an important role in various neurodevelopmental, neurodegenerative and
psychiatric disorders, and hence it is feasible that VGSC gating modifiers that augment
neuritogenesis and neurotrophin signaling may help in recovering from such disorders.
4-148
Figure 4-1: Veratridine stimulated neurite outgrowth and dendritic arborization in immature cerebrocortical neurons
4-149
Figure 4-2: Veratridine (VRT) increases intracellular Na+ and Ca2+ in DIV1 cerebrocortical neurons and this Ca2+ influx is TTX-sensitive and NMDAR dependent
4-150
Figure 4-3:TrkB is essential for veratridine-induced neurite outgrowthVeratridine
4-151
Figure 4-4: In situ BDNF ELISA: Veratridine enhances BDNF release in immature cerebrocortical neurons
4-152
Figure 4-5:Veratridine enhances BDNF synthesis in immature cerebrocortical neurons and this requires VGSCs and partially involves NMDARs
4-153
Figure 4-6: Veratridine enhances BDNF synthesis in immature cerebrocortical neurons and this requires VGSCs and partially involves NMDARs
4-154
Figure 4-7: - Veratridine-induced neurite outgrowth involves PI3-kinase activity
4-155
Figure 4-8- : Veratridine stimulated Akt phosphorylation involves TrkB receptors
4-156
Figure 4-9 - : Veratridine-induced neurite outgrowth involves the PI3K-Akt-mTOR pathway
4-157
Figure 4-10: - Veratridine-induced Ca2+ influx involves PLC mediated release of Ca2+ from intracellular
4-158
Figure 4-11: Veratridine-induced neurite outgrowth requires phospholipase C (PLC)
4-159
Figure 4-12:MAPK pathway has a modest role in veratridine-induced neurite outgrowth.
4-160
Figure 4-13:Pharmacological characterization of Akt activation by veratridine
4-161
Figure 4-14: Model 12
4-162
4.6 References
Balkowiec A and Katz DM. (2002) Cellular mechanisms regulating activity-dependent
release of native brain-derived neurotrophic factor from hippocampal neurons. J Neurosci
22:10399-10407.
Ben-Ari Y. (2001) Developing networks play a similar melody. Trends Neurosci 24:353-
360.
Brigadski T, Hartmann M, Lessmann V. (2005) Differential vesicular targeting and time
course of synaptic secretion of the mammalian neurotrophins. J Neurosci 25:7601-7614.
Dijkhuizen PA and Ghosh A. (2005) BDNF regulates primary dendrite formation in
cortical neurons via the PI3-kinase and MAP kinase signaling pathways. J Neurobiol
62:278-288.
Fink CC, Bayer KU, Myers JW, Ferrell JE,Jr, Schulman H, Meyer T. (2003) Selective
regulation of neurite extension and synapse formation by the beta but not the alpha
isoform of CaMKII. Neuron 39:283-297.
George J, Dravid SM, Prakash A, Xie J, Peterson J, Jabba SV, Baden DG, Murray TF.
(2009) Sodium channel activation augments NMDA receptor function and promotes
neurite outgrowth in immature cerebrocortical neurons. J Neurosci 29:3288-3301.
Ghosh A, Carnahan J, Greenberg ME. (1994a) Requirement for BDNF in activity-
dependent survival of cortical neurons. Science 263:1618-1623.
Ghosh A, Carnahan J, Greenberg ME. (1994b) Requirement for BDNF in activity-
dependent survival of cortical neurons. Science 263:1618-1623.
Ghosh A and Greenberg ME. (1995) Calcium signaling in neurons: Molecular
mechanisms and cellular consequences. Science 268:239-247.
4-163
Gomez TM and Spitzer NC. (2000) Regulation of growth cone behavior by calcium: New
dynamics to earlier perspectives. J Neurobiol 44:174-183.
Gomez-Pinilla F. (2008) The influences of diet and exercise on mental health through
hormesis. Ageing Res Rev 7:49-62.
Goodman LJ, Valverde J, Lim F, Geschwind MD, Federoff HJ, Geller AI, Hefti F. (1996)
Regulated release and polarized localization of brain-derived neurotrophic factor in
hippocampal neurons. Mol Cell Neurosci 7:222-238.
Hartmann M, Heumann R, Lessmann V. (2001) Synaptic secretion of BDNF after high-
frequency stimulation of glutamatergic synapses. EMBO J 20:5887-5897.
Hui K, Fei GH, Saab BJ, Su J, Roder JC, Feng ZP. (2007) Neuronal calcium sensor-1
modulation of optimal calcium level for neurite outgrowth. Development 134:4479-4489.
Jabba SV, Prakash A, Dravid SM, Gerwick WH, Murray TF. (2010) Antillatoxin, a novel
lipopeptide, enhances neurite outgrowth in immature cerebrocortical neurons through
activation of voltage-gated sodium channels. J Pharmacol Exp Ther 332:698-709.
Kidane AH, Heinrich G, Dirks RP, de Ruyck BA, Lubsen NH, Roubos EW, Jenks BG.
(2009) Differential neuroendocrine expression of multiple brain-derived neurotrophic
factor transcripts. Endocrinology 150:1361-1368.
Klocker N, Cellerino A, Bahr M. (1998) Free radical scavenging and inhibition of nitric
oxide synthase potentiates the neurotrophic effects of brain-derived neurotrophic factor
on axotomized retinal ganglion cells in vivo. J Neurosci 18:1038-1046.
Kohara K, Kitamura A, Adachi N, Nishida M, Itami C, Nakamura S, Tsumoto T. (2003)
Inhibitory but not excitatory cortical neurons require presynaptic brain-derived
4-164
neurotrophic factor for dendritic development, as revealed by chimera cell culture. J
Neurosci 23:6123-6131.
Kohara K, Kitamura A, Morishima M, Tsumoto T. (2001) Activity-dependent transfer of
brain-derived neurotrophic factor to postsynaptic neurons. Science 291:2419-2423.
Kolarow R, Brigadski T, Lessmann V. (2007) Postsynaptic secretion of BDNF and NT-3
from hippocampal neurons depends on calcium calmodulin kinase II signaling and
proceeds via delayed fusion pore opening. J Neurosci 27:10350-10364.
Konur S and Ghosh A. (2005) Calcium signaling and the control of dendritic
development. Neuron 46:401-405.
Lipton SA and Nakanishi N. (1999) Shakespeare in love--with NMDA receptors? Nat
Med 5:270-271.
Mamounas LA, Altar CA, Blue ME, Kaplan DR, Tessarollo L, Lyons WE. (2000) BDNF
promotes the regenerative sprouting, but not survival, of injured serotonergic axons in the
adult rat brain. J Neurosci 20:771-782.
Marini AM, Rabin SJ, Lipsky RH, Mocchetti I. (1998) Activity-dependent release of
brain-derived neurotrophic factor underlies the neuroprotective effect of N-methyl-D-
aspartate. J Biol Chem 273:29394-29399.
McAllister AK. (2000) Cellular and molecular mechanisms of dendrite growth. Cereb
Cortex 10:963-973.
McAllister AK, Lo DC, Katz LC. (1995) Neurotrophins regulate dendritic growth in
developing visual cortex. Neuron 15:791-803.
Miller FD and Kaplan DR. (2003) Signaling mechanisms underlying dendrite formation.
Curr Opin Neurobiol 13:391-398.
4-165
Nakajima T, Sato M, Akaza N, Umezawa Y. (2008) Cell-based fluorescent indicator to
visualize brain-derived neurotrophic factor secreted from living neurons. ACS Chem Biol
3:352-358.
Rajan I and Cline HT. (1998) Glutamate receptor activity is required for normal
development of tectal cell dendrites in vivo. J Neurosci 18:7836-7846.
Rattiner LM, Davis M, Ressler KJ. (2005) Brain-derived neurotrophic factor in
amygdala-dependent learning. Neuroscientist 11:323-333.
Reichardt LF. (2006) Neurotrophin-regulated signalling pathways. Philos Trans R Soc
Lond B Biol Sci 361:1545-1564.
Rose CR and Konnerth A. (2001) NMDA receptor-mediated na+ signals in spines and
dendrites. J Neurosci 21:4207-4214.
Saneyoshi T, Wayman G, Fortin D, Davare M, Hoshi N, Nozaki N, Natsume T,
Soderling TR. (2008) Activity-dependent synaptogenesis: Regulation by a CaM-kinase
kinase/CaM-kinase I/betaPIX signaling complex. Neuron 57:94-107.
Shen K and Meyer T. (1998) In vivo and in vitro characterization of the sequence
requirement for oligomer formation of Ca2+/calmodulin-dependent protein kinase
IIalpha. J Neurochem 70:96-104.
Tolias KF, Bikoff JB, Burette A, Paradis S, Harrar D, Tavazoie S, Weinberg RJ,
Greenberg ME. (2005) The Rac1-GEF Tiam1 couples the NMDA receptor to the
activity-dependent development of dendritic arbors and spines. Neuron 45:525-538.
Ultanir SK, Kim JE, Hall BJ, Deerinck T, Ellisman M, Ghosh A. (2007) Regulation of
spine morphology and spine density by NMDA receptor signaling in vivo. Proc Natl
Acad Sci U S A 104:19553-19558.
4-166
Vaillant AR, Zanassi P, Walsh GS, Aumont A, Alonso A, Miller FD. (2002) Signaling
mechanisms underlying reversible, activity-dependent dendrite formation. Neuron
34:985-998.
Wayman GA, Impey S, Marks D, Saneyoshi T, Grant WF, Derkach V, Soderling TR.
(2006) Activity-dependent dendritic arborization mediated by CaM-kinase I activation
and enhanced CREB-dependent transcription of wnt-2. Neuron 50:897-909.
West AE, Chen WG, Dalva MB, Dolmetsch RE, Kornhauser JM, Shaywitz AJ, Takasu
MA, Tao X, Greenberg ME. (2001) Calcium regulation of neuronal gene expression.
Proc Natl Acad Sci U S A 98:11024-11031.
West AE, Griffith EC, Greenberg ME. (2002) Regulation of transcription factors by
neuronal activity. Nat Rev Neurosci 3:921-931.
Wong RO and Ghosh A. (2002) Activity-dependent regulation of dendritic growth and
patterning. Nat Rev Neurosci 3:803-812.
Wu GY and Cline HT. (1998) Stabilization of dendritic arbor structure in vivo by
CaMKII. Science 279:222-226.
Yoshii A and Constantine-Paton M. (2010) Postsynaptic BDNF-TrkB signaling in
synapse maturation, plasticity, and disease. Dev Neurobiol 70:304-322.
Yu XM, Askalan R, Keil GJ,2nd, Salter MW. (1997) NMDA channel regulation by
channel-associated protein tyrosine kinase src. Science 275:674-678.
Yu XM and Salter MW. (1998) Gain control of NMDA-receptor currents by intracellular
sodium. Nature 396:469-474.
4-167
Zou DJ and Cline HT. (1999) Postsynaptic calcium/calmodulin-dependent protein kinase
II is required to limit elaboration of presynaptic and postsynaptic neuronal arbors. J
Neurosci 19:8909-8918.