ARHGAP4 IS a SPATIALLY REGULATED RHOGAP THAT INHIBITS NIH/3T3 CELL MIGRATION and DENTATE GRANULE CELL AXON OUTGROWTH by DANIEL L
ARHGAP4 IS A SPATIALLY REGULATED RHOGAP THAT INHIBITS
NIH/3T3 CELL MIGRATION AND DENTATE GRANULE CELL AXON
OUTGROWTH
By
DANIEL LEE VOGT
Submitted in partial fulfillment of the requirements
for the degree of Doctor of Philosophy
Department of Neuroscience
CASE WESTERN RESERVE UNIVERSITY
August, 2007
CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
Daniel Lee Vogt ______candidate for the Ph.D. degree *.
(signed) (chair of the committee)______Stefan Herlitze
______Alfred Malouf
Robert Miller ______
______Thomas Egelhoff
______Susann Brady-Kalnay
______
(date) ______6-21-2007
*We also certify that written approval has been obtained for any proprietary material contained therein.
ii
Copyright © 2007 by Daniel Lee Vogt
All rights reserved
iii Table of contents
Page #
Title page i
Table of contents iv
List of figures vii
Abstract 1
Chapter one: General introduction 2
Hippocampal axon pathways and development 3
Guidance cues in hippocampal axon outgrowth 6
Slit/Robo 7
Semaphorins, plexins and neuropilins 8
Ephrins and ephs 11
Other guidance cues in the hippocampus 13
GTPases: structure and function of ras superfamily members 15
Ras GTPases 17
Ran GTPases 18
Arf GTPases 18
Rab GTPases 19
Rho GTPases 20
GTPase regulatory proteins and the role of GAPs 23
RasGAP family functions, localizations and binding partners 24
RanGAP family 25
ArfGAP family 26
iv RabGAP family 26
RhoGAP family 27
GAP protein mutations and developmental consequences 29
Regulation of GAP proteins 29
Phosphorylation 30
Protein-protein interactions 31
Other types of regulation 32
Actin and microtubule dynamics in growth cone and fibroblast motility 33
General actin dynamics 33
General microtubule dynamics 36
Coordinated signaling regulated by Rho-family GTPases 40
Cell migration and functional parallels to growth cone guidance 41
Previous work on ARHGAP4 45
Conserved domains of ARHGAP4 48
FCH domain, extended FCH domain and ARNEY domain 48
RhoGAP domain 51
SH3 domain 52
Proline rich (PxxP) domains 52
Research goals 53
Figures 55
Chapter two: ARHGAP4 is a spatially regulated GTPase activating protein
(GAP) expressed in NIH/3T3 fibroblasts and dentate granule neurons 71
Summary 72
v Introduction 72
Materials and methods 74
Results 82
Discussion 89
Figures 97
Chapter three: ARHGAP4 is an inhibitor of NIH/3T3 cell motility and
Dentate granule cell axon outgrowth 123
Summary 124
Introduction 124
Materials and methods 126
Results 130
Discussion 133
Figures 139
Chapter four: Mechanistic insights into the function of ARHGAP4 155
Summary 156
Introduction 156
Materials and methods 158
Results 161
Discussion 165
Figures 169
Chapter five: General Discussion 179
Chapter six: Bibliography 195
vi List of figures
Figure Page #
1.1 Hippocampal anatomy and pathways 55
1.2 Common guidance cues and receptors found in the hippocampus utilize various
GTPases to effect growth cone guidance 57
1.3 The GTPase cycle and role of GAPs, GEFs and GDIs 59
1.4 Rho-family GAP proteins and their domain assortments 61
1.5 Actin and microtubule cytoskeletons in migrating cells and growth cones 63
1.6 ARHGAP4’s amino-terminus contains FCH, extended FCH and ARNEY domains 65
1.7 ARHGAP4 GAP domain alignment 67
1.8 ARHGAP4 SH3 domain alignment 69
2.1 ARHGAP4 constructs used in transfections and bacterial protein expression 97
2.2 In vitro GAP assay of WT and R562A ARHGAP4 GAP domains 99
2.3 Endogenous ARHGAP4 localizes to the leading edge of NIH/3T3 cells 101
2.4 Amino acids 1-71 are necessary and sufficient to target ARHGAP4 to the tips of NIH/3T3 cell cytoplasmic extensions 103
2.5 ARHGAP4 is enriched in the mossy fiber axons of the hippocampus 105
2.6 Endogenous ARHGAP4 is enriched in mossy fiber growth cones 107
2.7 ARHGAP4 expressed proteins localize to growth cones via amino acids
1-71 109
2.8 Amino acids 1-71 are necessary and sufficient to target ARHGAP4 to growth cones 111
vii 2.9 Amino acids 1-71 can associate indirectly but do not bind microtubules
in an in vitro microtubule cosedimentation assay 113
2.10 Leading edge distribution of microtubules and F-actin in the presence of
nocodazole or cytochalasin-D 115
2.11 Full length ARHGAP4 (1-965) leading edge distribution in the presence of
nocodazole or cytochalasin-D 117
2.12 1-770 leading edge distribution in the presence of nocodazole or
cytochalasin-D 119
2.13 72-965 leading edge distribution in the presence of nocodazole or
cytochalasin-D 121
3.1 Wound assay model system and quantification of cell migration in
individually transfected cells 139
3.2 siRNA mediated knockdown of ARHGAP4 increases cell migration in
NIH/3T3 cells 141
3.3 ARHGAP4 inhibits NIH/3T3 cell migration via its GAP domain 143
3.4 ARHGAP4 inhibits axon outgrowth from dentate explant cultures through its
FCH, GAP and SH3 domains 145
3.5 Dentate explant axon outgrowth is inhibited by ARHGAP4 in an FCH, GAP and SH3 domain manner 147
3.6 Dentate explant astrocyte outgrowth is not altered by ARHGAP4 149
3.7 ARHGAP4 inhibits axon outgrowth in dissociated granule cells in a GAP
dependent manner 151
3.8 ARHGAP4 significantly inhibits axon outgrowth in dissociated granule cells
viii in a GAP dependent manner 153
4.1 ARHGAP4 amino-terminal protein design and expression 169
4.2 Amino acids 1-289 are the minimal unit required to inhibit NIH/3T3 cell migration 171
4.3 Amino acids 1-71 and 1-289 of ARHGAP4 result in different actin phenotypes in migrating NIH/3T3 cells 173
4.4 Full length ARHGAP4 overexpression decreases levels of GTP-bound RhoA in NIH/3T3 cells 175
4.5 Trends in axon outgrowth and branching are altered in DRGs expressing the R562A mutant on laminin and aggrecan 177
5.1 Model of ARHGAP4 activity 193
ix ARHGAP4 is a spatially regulated RhoGAP that inhibits NIH/3T3 cell migration
and dentate granule cell axon outgrowth
Abstract
By
DANIEL LEE VOGT
Cell migration and axonal growth cone guidance are tightly regulated events that
share many similarities. While some factors are not always shared between migrating
cells and growth cones, the signaling events that are required for cell migration and
growth guidance each utilize GTPases to regulate the actin and microtubule
cytoskeletons. GTPases are expressed ubiquitously but signal in discrete subcellular
locales to control directed cell migration and growth cone guidance. This strict control of
GTPase signaling is controlled by GAPs, GEFs and GDIs, which can assemble signaling
complexes and localize to unique regions due to an array of conserved functional
domains. The Rho-family GAP ARHGAP4 contains FCH and SH3 domains, as well as
other conserved domains of unknown function. The FCH domain has been attributed to
binding actin, microtubules and lipids, but there is no consensus on its function. Here we
show that ARHGAP4 localizes to the leading edges of NIH/3T3 fibroblasts and to growth
cones of dentate granule neurons, via its FCH domain. Overall, ARHGAP4 inhibits
NIH/3T3 cell migration and dentate granule cell axon outgrowth in a GAP dependent
manner, and this inhibition is temporally and spatially regulated by ARHGAP4’s conserved functional domains.
1 Chapter 1
General Introduction
2
Unraveling the intricacies of neural networks and their functions during development and in the adult has been an ongoing task since the infancy of developmental biology. Deciphering how neurons migrate, send axons and dendrites to their proper targets and communicate with the vast array of non-neuronal cells is still not well understood. Although many signaling events have been characterized and candidate proteins identified, the detailed mechanisms whereby all the players in neuronal development and maturation are brought together is still far from complete. The mechanisms that regulate axon outgrowth and guidance require tight regulation of cytoskeletal elements, and these mechanisms are shared among many migratory cells as well. Conserved families of proteins that include guidance and growth cues with their respective receptors, actin and microtubule regulatory proteins, GTPases and their regulators the GEFs, GAPs and GDIs, are all involved in guiding an axon or migrating cell to its determined location. These signaling cascades require spatial and temporal control to utilize the potential of the GTPases, and regulatory proteins must have functional domains that allow for spatial control and assembly with signaling components. The RhoGAP family of proteins have diverse tissue expression, subcellular location and signaling partners, and achieve this through a myriad of functional domains.
ARHGAP4 is a RhoGAP that inhibits axon outgrowth and cell motility, and whose actions are temporally and spatially regulated by its conserved functional domains.
Hippocampal axon pathways and development
One of the classic areas studied for axon outgrowth is the hippocampus. With its stereotyped pattern of axonal fibers, adult neurogenesis and axon outgrowth, and rich
3 history of electrophysiological mapping of circuits, the hippocampus has been an excellent model to study axon outgrowth and regeneration. The hippocampus arises from the invaginating dorsal midline of the telencephalon, and its initial formation is dependent on the patterning of the dorsal telencephalon (Theil et al., 1999). The mature hippocampus has a recognizable morphology with distinguishable dentate gyrus and cornu ammon horn areas (CA) 1 and 3. Both the CA areas and dentate gyrus can be subdivided into specific layers, that are specified locations for dendrites and axon pathways, (figure 1.1). The cells that will become dentate granule cells are derived from the neuroepithelium after E16 in the rat, and migrate in two distinct waves to form the dentate gyrus (Altman and Bayer, 1990). The mature dentate gyrus is divided into a molecular layer (ML) which contains dentate granule cell dendrites, the dentate granule cell layer (DGL) which contains the dentate granule cell somas, and the hilus (H) which is the area where dentate granule cell axons, mossy fibers (MFs), project through (Figure
1.1). The earliest born cells are closest to the pia (and nearer to the molecular layer), while the later born cells are added on the side facing the forming hilus. The granule cells start to send out their axons into the hilus as early as P0, and start to invade the stratum lucidum (SL) layer by P3 (Amaral and Dent, 1981). MFs send out several collateral into the hilus, but compress into a narrow tract when they reach the SL (Amaral and Dent, 1981; Claiborne et al., 1986). As the mossy fibers traverse through the SL they come into contact with CA3 pyramidal cell dendrites. Pyramidal cells start to show spines on their proximal dendrites between P9-P11, which interact with the MF axons to form synapses called thorny excrescenses (Amaral and Dent, 1981).
4 The MF to CA3 projection is one component of a well studied circuit involved in learning and memory, that also includes projections from the entorhinal cortex and CA3 to CA1 projections. Axons from the entorhinal cortex terminate onto dentate granule cell dendrites and constitute the perforant pathway, and are the major input of information to the hippocampus (Dolorfo and Amaral, 1998b, a). The dentate gyrus to CA3 constitutes the MF pathway (Henze et al., 2000), while the CA3 to CA1 constitutes the Schaffer collateral pathway, and both of these pathways are involved in the induction of long term potentiation (LTP) associated with memory formation (Lopez-Garcia, 1998). The MF pathway is mostly mature by P21 in the rat and new granule cells are continuously born and send out axons through this postnatal and adult environment (Lledo et al., 2006). In the adult animal, new granule cells are continuously generated in the subgranular zone, which faces the hilus (Kaplan and Hinds, 1977; Kaplan and Bell, 1984), and these neurons send out axons that incorporate into the established network and function like neighboring granule cells (Laplagne et al., 2006). In addition, the MFs are capable of regenerating after injury , supporting the hypothesis that there are signaling events in MF growth cones that are able to overcome inhibitory cues in the adult (Butler et al., 2004;
Emery et al., 2005). While this regeneration is a special feature of these axons, it is not surprising considering that dentate granule cells are continuously born in the adult and must send out their axons in this adult landscape of inhibitory cues.
The signaling events that guide MFs to their targets during development and in the adult are not well worked out, but an understanding of these events and the proteins involved would greatly advanced the ability to develop treatments for diseases and traumatic injuries that inhibit axon outgrowth or have led to a loss of neurons needed for
5 normal cognitive function. These include but are not limited to alzheimer’s disease,
stroke, epilepsy, mental retardation, and depression. While there are many holes to be
filled in understanding how MFs navigate the hippocampus, many guidance cue families
have emerged as important regulators of specific axon pathways in the hippocampus, and
have elucidated several signaling events that may serve as theraputic tools to treat
ailments that lead to a loss of axon outgrowth or neuronal damage. In addition, many of
these guidance cues utilize similar GTPase signaling pathways to regulate cytoskeletal
elements during axon guidance.
Guidance cues in hippocampal axon outgrowth
The axons and dendrites that constitute the hippocampal areas are formed by a
combination of attractive and repulsive cues which are expressed in a restricted spatial
and temporal pattern. The postnatal environment of the brain is highly restrictive to
unwarranted growth and migration, yet the MF axons of the dentate granule cells are
capable of navigating the hippocampus in a stereotyped pattern to reach their appropriate targets throughout adulthood (Stanfield and Trice, 1988). How these particular axons are capable of navigating this environment is still not well understood. There have been a number of repulsive and attractive guidance cues identified in the hippocampus during early postnatal stages when the first MFs grow out, and present at later stages into adulthood when new dentate granule cells send out MFs. Several guidance cues are now known to regulate various axons and dendrite trajectories in the developing and postnatal hippocampus. These cues include, but are not limited to, the slits, semaphorins, ephrins,
6 proteoglycans and, and cell adhesion molecules (Milner and Campbell, 2002; Huber et al., 2003).
Slit/ROBO
The Slit proteins are one the most well studied guidance cue families and are ligands for the roundabout (ROBO) receptors (Brose et al., 1999; Yuan et al., 1999).
While the Slits have roles in the cell migration of both neurons (Nguyen-Ba-Charvet et al., 2004) and leukocytes (Wu et al., 2001), they are better known for their repulsive actions on axons. The Slit/ROBO interaction was first characterized in Drosophila midline axon crossing, where high levels of slit in the ventral midline glia (Rothberg et al., 1988; Rothberg et al., 1990) repulse the axons that present the ROBO receptor after crossing the midline (Kidd et al., 1998). Slit/ROBO interactions also influence several axons in mammals, including those of the olfactory bulb (Li et al., 1999) and hippocampus (Nguyen Ba-Charvet et al., 1999). Mammalian slit1, slit2 and slit3 are enriched in different tissues, with expression in the brain, spinal cord and thyroid, respectively (Itoh et al., 1998). Brain-enriched slit1 is expressed in the amygdala, caudate nucleus, hippocampus, hypothalamus, and cerebral cortex at E18, and is expressed in the cerebral cortex, hippocampus, olfactory bulb and amygdala in the adult
(Itoh et al., 1998).
Within the hippocampus Slit1 is expressed in the CA3 and CA1 pyramidal neurons as well as the dentate granule neurons (Itoh et al., 1998). Although slit1 is enriched in the brain (Itoh et al., 1998), all three slit mRNAs have been detected in the hippocampus (Marillat et al., 2002). The Slit receptors ROBO1 and ROBO2 have
7 differential expression patterns in the hippocampus. ROBO1 is expressed as early as E15 in the CA1 and CA3 regions and turns on at P0 in the dentate gyrus, while ROBO2 begins expressing at E18 in the subiculum and at P5 in the dentate gyrus, with both of these expression patterns continuing into adulthood (Marillat et al., 2002). While studies have demonstrated a role for Slit/ROBO signaling in hippocampal axon repulsion in vitro
(Nguyen Ba-Charvet et al., 1999), there have been no studies looking at the role of
Slit/ROBOs in terms of axon guidance in the postnatal hippocampus. Most work in the brain has focused on olfactory bulb projections (Nguyen-Ba-Charvet et al., 2002), retinal ganglion axon projections (Plump et al., 2002), as well as gross guidance defects in the corpus callosum, corticothalamic and thalamocortical axon tracts (Bagri et al., 2002).
Activated ROBO can recruit several proteins to its cytoplasmic domains to transduce its repulsive cue, including the Slit/ROBO (sr)GAPs (figure 1.2), and the Abl tyrosine kinase. Once bound to the ROBO receptor, Slit ROBO (sr)GAP1 has been shown to locally inactivate the GTPase Cdc42 (Wong et al., 2001), which can locally destabilize the actin network and cause growth cone turning. The ROBO receptor can also inhibit the attractive cues transduced by the netrin receptor frazzled in the
Drosophila midline (Bhat, 2005), and potentially regulate microtubules (MTs) in growth cones through its regulation of the kinase Abl (Lee et al., 2004).
Semaphorins, Plexins and Neuropilins
The Semaphorins compose another family of guidance cues that transduce their signals through the plexin receptor family or form a tertiary complex with the neuropilins in vertebrates (Tamagnone and Comoglio, 2000). The over 20 semaphorins share an
8 amino-terminal sema domain (Gherardi et al., 2004) and each have a variable carboxy-
terminus (Tamagnone and Comoglio, 2000), and are categorized into 8 classes reflecting their sequence similarity and organism they are derived from. They can be membrane bound (classes 1, 4, 5, and 6), GPI anchored (class 7), or secreted (classes 2, 3, or V)
(Nakamura et al., 2000). There are also several plexin receptor types, including plexin-
A1-4, plexin-B1-3, plexin-C1, and plexin-D1 (Tamagnone et al., 1999), as well as neuropilin receptors 1 and 2, which can bind the class 3 semaphorins and signal through a tertiary complex that includes a plexin receptor (Bagri and Tessier-Lavigne, 2002).
The class 3 semaphorins (sema 3A, B, C, D, E and F) have been most extensively characterized in hippocampal development and are the only semaphorins known to bind to the neuropilin receptors (Kolodkin et al., 1997). Sema 3A is expressed in the entorhinal cortex, while sema 3C is expressed in various interneurons, dentate granule neurons and in the CA3 and CA1 pyramidal neurons. Sema 3E is expressed in the dentate granule neurons, CA3 and CA1 pyramidal neurons and within the mossy cells that reside in the hilus. Lastly, sema 3F is expressed in dentate granule neurons, CA3 and
CA1 pyramidal neurons, and within various interneurons (Sahay et al., 2005). The neuropilins are expressed in the developing dentate gyrus at late embryonic stages and at
P0 in the mouse (Chedotal et al., 1998), and continue to be expressed into postnatal and adult stages (Pascual et al., 2005). Neuropilin1 can bind preferentially to sema 3A
(Kolodkin et al., 1997), while neuropilin2 can interact with sema E, IV and 3F (Chen et al., 1997; Gammill et al., 2006a). The neuropilins do not have intracellular domains and are thought to transduce signals by forming a ternary complex with the plexin receptors
(Puschel, 2002). The plexinA receptors also have unique expression patterns in the
9 hippocampus. At postnatal day 3, plexins A1, A2, and A3 are found in the subiculum
and in the CA1, and CA3 pyramidal cell layers, while only plexin A2 was expressed at high levels in the DGL (Murakami et al., 2001). Plexin-B1 and B2 receptors are
expressed in many areas of the brain from embryonic stages into adulthood, while plexin-
B3 turns on perinatally, peaks at P7, and declines thereafter (Worzfeld et al., 2004).
However, even though the plexin-B receptors have been implicated in collapsing
hippocampal growth cones (Oinuma et al., 2004a), there has not been an extensive
examination of their expression patterns in the postnatal hippocampus.
The semaphorins have the ability to collapse growth cones and repel hippocampal
axons (Luo et al., 1993; Chedotal et al., 1998), and transduce their signals through several
GTPase signaling networks to alter both the actin and microtubule cytoskeletons (Huber
et al., 2003; Togashi et al., 2006), (figure 1.2). The plexinB1 receptor can bind the ligand sema3A, and this interaction allows plexinB1 to bind and sequester Rac1 from its effectors, leading to growth cone collapse (Vikis et al., 2000). The plexin receptors are also unique in that they contain a region of homology with GTPase activating protein
(GAP) domains, and plexin-B1 can act as a GAP for R-Ras (Oinuma et al., 2004b;
Oinuma et al., 2004a). Interestingly, the GTPase Rnd1 binds the plexin-B1 receptor in a manner that activates its GAP activity for R-Ras leading to collapse of hippocampal growth cones (Oinuma et al., 2004a), and stimulates the interaction of PDZ-RhoGEF with the plexin-B1 receptor and subsequent RhoA activation in COS-7 cells (Oinuma et al., 2003). Rnd1 induced activation can also be antagonized by the actions of RhoD, which can compete for binding to the plexin receptor (Zanata et al., 2002). The GAP activity towards R-ras and the Guanine nucleotide exchange factor (GEF) activity
10 towards RhoA may both occur in neurons and COS-7 cells, or each event may be the
result of the plexin-B1 receptor utilizing local proteins specifically expressed in each cell
type.
While the aforementioned events mainly target the actin cytoskeleton, new target
proteins have been identified downstream of semaphorin signaling that modify MTs as
well. The Collapsin Response Mediator Protein-2 (CRMP-2) (Gu and Ihara, 2000) and
CRMP-associated molecule (CRAM) (Hotta et al., 2005) are MT associated proteins that
function downstream of semaphorin signaling to regulate MT dynamics involved in
growth cone collapse and guidance. Aside from their well known role in axon guidance
and growth cone collapse, the semaphorins and their receptors are involved in guiding the
migration of several different cell types (Cohen, 2005; Kerjan et al., 2005; Gammill et al.,
2006b). In addition, semaphorin signaling can be an attractive for some dendrites
(Polleux et al., 2000), and the neuropilin receptors can form complexes with other
factors, including the cell adhesion molecule L1 (Castellani et al., 2000) and VEGF
(Soker et al., 1998).
Ephrins and Ephs
The ephrins are ligands that bind to the Eph receptors and are involved in cell migration (Poliakov et al., 2004), synapse plasticity and axon guidance (Martinez and
Soriano, 2005). Ephrins fall into two categories: ephrin-As that are GPI-anchored and
bind to the EphA receptors (EphA1-8), and ephrin-Bs that are transmembrane ligands and
bind to the EphB receptors (EphB1-6) as well as the EphA4 receptor (Himanen and
Nikolov, 2003; Blits-Huizinga et al., 2004). The Eph receptors are receptor tyrosine
11 kinases that dimerize via their fibronectin repeat domains when activated (Lackmann et al., 1998; Himanen et al., 2001). When ephrins bind to their Eph receptors, both bidirectional signaling can occur, and Eph receptor (forward) signaling and ephrin
(reverse) signaling are involved in defining cell boundaries (Sela-Donenfeld and
Wilkinson, 2005), synaptic formation and plasticity in the hippocampus (Henkemeyer et al., 2003; Armstrong et al., 2006; Moeller et al., 2006). The ability of ephrins and Eph gradients to control axon guidance was firmly established in the retino-tectal/superior colliculus topographic map, where a gradient of low to high ephrin expression dictates the final position of retinal axons in the tectum/superior colliculus (McLaughlin and
O'Leary, 2005), and the hippocampal to septal projection (Gao et al., 1996; Zhou, 1997).
Within the hippocampus, dentate granule cells express ephrinA3 and EphA5, while the CA1 and CA3 fields express ephrinsA3 and A5 as well as EphA5 (Skutella and
Nitsch, 2001). Other ephrins and Ephs have been implicated in hippocampal axon guidance, including the EphA receptors 3-7, which increase expression from the lateral to medial areas of the hippocampus, while the expression of ephrins A2, 3, and 5 increases from dorso-medial to ventro-lateral regions in the septum (Linke et al., 1995). Outgrowth from medial hippocampal neurons, but not lateral hippocampal neurons, is inhibited by ephrins A2, A3, and A5 (Brownlee et al., 2000). In mice expressing an EphA5 mutant receptor, hippocampal axons projecting to the septum from medial areas were highly disorganized, but axons from lateral regions were unaffected (Yue et al., 2002), demonstrating a functional role for the ephrin/Eph interaction in this pathway.
Interestingly, in EphB2 and B3 knockouts these projections become defasiculated when they reach their target area, indicating another level of regulation (Chen et al., 2004).
12 Additionally, the inner molecular layer of the dentate gyrus expresses Eph A5, which
excludes ephrin A3 commissural axons from this area (Otal et al., 2006), and is also
thought to keep the dentate granule cell axons from invading the ML.
Ephs and ephrins regulate members of the Rho and Ras families of GTPases to
mediate many of their effects (Klein, 2004; Dail et al., 2006), (figure 1.2). The EphA
receptors can phosphorylate the RhoA GEF ephexin, promoting its activation and the
subsequent upregulation of active RhoA (Sahin et al., 2005). The EphB2 receptor is
known to inhibit R-ras, by activating the RasGAP p120 (Dail et al., 2006). While the
actin cytoskeleton is a primary target, MTs can also be targeted by Eph/ephrin signaling.
CRMP-2 is known to bind tubulin heterodimers and microtubule polymers and facilitate polymerization (Fukata et al., 2002b). In the presence of ephrin-A5, the RhoA effector
ROCK is activated and subsequently phosphorylates CRMP-2, abolishing its binding to
MTs and contributing to growth cone collapse (Arimura et al., 2005). These events mediate the repulsion of axons, but other functions of Eph/ephrin and GTPase signaling have been discovered, including the maturation of dendritic spines and synapses (Kayser et al., 2006; Moeller et al., 2006).
Other guidance cues in the hippocampus
There are many other guidance entities in the hippocampus that have not received as much attention as the above guidance cue families. The proteoglycans are one such family that mediate repulsive axon guidance cues. The chrondroitin sulfate proteoglycans (CSPGs) and keratin sulfate proteoglycans (KSPGs) are two examples that have been implicated in sculpting MF axon tracts (Butler et al., 2004). Proteoglycan
13 levels can also increase in certain conditions that affect hippocampal activity, and have
been shown to increase in the dentate gyrus area in mice with temporal lobe epilepsy
(Heck et al., 2004), demonstrating that levels of guidance factors can differ over the
lifetime of an organism in response to environmental changes. Proteoglycan-mediated
repulsion is thought to occur by activating RhoA-mediated signaling events (Huber et al.,
2003; Schweigreiter et al., 2004).
Several attractive factors that shape the hippocampus have also been identified,
including Netrin/deleted in colon cancer (DCC), L1, and the neural cell adhesion
molecule (NCAM). Netrin is an attractive factor that is highly expressed in the
developing fimbria and at a lower level in the dentate gyrus, CA1 and CA3 regions at
mid to late embryonic periods, but decreases by p0 (Barallobre et al., 2000). Netrin is a ligand for the receptor DCC, which is also expressed in the developing and adult hippocampus (Volenec et al., 1997), and Netrin-1 knockouts show targeting defects that include loss of contralaterally projecting hippocampal axons to the septum and hippocampus, as well as multiple hippocampal projections targeting to the wrong layers within the hippocampus itself (Barallobre et al., 2000). The cell adhesion molecule L1 is also involved in hippocampal morphology. In L1 knockouts, there is loss of dentate granule cells and pyramidal cells, and those pyramidal cells still present have defects in their apical dendrites (Demyanenko et al., 1999). The modified version of NCAM, polysialylated (PSA)-NCAM, is highly expressed in the MF axons of the hippocampus and loss of this protein results in major mossy fiber guidance defects in both the developing and adult organism (Cremer et al., 1997).
14 Although each of the above guidance cue families utilize unique receptors, and
are expressed in differential spatial and temporal patterns, they all utilize the GTPases in
some way to transduce their instructions to migrating axons. This common feature
allows various signals to converge on targets such as the actin and MT cytoskeletons, as
well as other proteins that regulate membrane integrity, transcription and translation to
guide a growth cone through its environment. Understanding how these signals control
growth cone guidance requires an understanding of the GTPases involved, their
regulation of target proteins, and crosstalk between GTPase families.
GTPases: Structure and function of Ras superfamily members
GTPases are small molecular weight proteins that cycle between a GTP-bound and GDP-bound state, and control a diverse array of signaling events (Mitin et al., 2005),
(figure 1.3, borrowed from Bernards and Settleman, 2004). GTP-bound GTPases are considered active because many GTPases bind their effector proteins and stimulate signaling events in this state. In turn, when GTP is hydrolysed to GDP, the GTPase is considered inactive and unable to bind effector proteins (Takai et al., 2001). These events are regulated by GEFs and GAPs, which are structurally suited to enhance GDP for GTP exchange and GTP hydrolysis, respectively (Geyer and Wittinghofer, 1997). In addition GTPases can be sequestered in the cytoplasm by guanine dissociation inhibititors (GDIs) which prevent dissociation of GDP from the GTPase (Dransart et al.,
2005).
When the small GTPases RhoA and Rac1 were found to mediate the changes in the actin cytoskeleton in response to extracellular cues (Ridley and Hall, 1992; Ridley et
15 al., 1992), it brought the signaling role of these proteins into perspective. These changes were the alterations of well recognized structures (stress fibers, membrane ruffles, and filopodia) by specific the GTPases RhoA, Rac1 and Cdc42 respectively (Allen et al.,
1997). The GTPases are now recognized as key players in many events that regulate cytoskeletal dynamics. The Rho-family GTPases are part of a larger group of GTPases, called the Ras superfamily. This superfamily includes the Ras, Ran, Arf, Rab and Rho subfamilies, categorized by their respective homologies and by their abilities to regulate similar mechanisms in cells. (Burridge and Wennerberg, 2004; Wennerberg et al., 2005).
The GTPase family proteins share structural features that are critical for their function, including binding to their effectors. The GTPases have flexible regions, referred to as switch helix I and switch helix II, which undergo conformational changes when the GTPase is bound to GTP. These changes expose sites on the GTPases that are required for binding to effectors as well as to GAPs (Schaber et al., 1989). GTPases share several conserved residues that, when mutated, drastically alter the biological functions of the GTPases. Some of these mutations, first discovered and changed in several Ras proteins, have led to commonly used protein tools still utilized today. Ras
GTP hydrolysis requires a primed water molecule to hydrolyze the gamma phosphate from GTP, but when the invariant glycine 12 is mutated, Ras becomes constitutively active and can cause aberrant growth and is the cause of many cancer types (Taparowsky et al., 1982). From crystal structures, it is now known that any residue other than glycine at this position sterically hinders water from entering the catalytic site to hydrolyze GTP to GDP (Krengel et al., 1990). Conversely, mutation of serine 17 to asparagine was shown to block the transforming abilities of Ras by locking the GTPase in a GDP-bound
16 form (Kaplan, 1994). These constitutively active (CA) and dominant negative (DN) mutants have been widely used to understand the roles of GTPases from all the Ras superfamily members. Although the Rho-family of GTPases are well known for
mediating cytoskeleton dynamics, there is a great deal of cytoskeletal regulation by other
GTPase subfamiles, both directly and indirectly through crosstalk with the Rho-family of
GTPases.
Ras GTPases
The Ras family of GTPases includes the Ras GTPases (H-ras, N-ras, and K-ras),
as well as R-Ras, Rap and Ral GTPases, which are highly conserved except in the
carboxy-terminus within the hypervariable domain (Hancock, 2003; Mitin et al., 2005).
This hypervariable domain is thought to provide some differential functions between the
Ras family members, and also contains a CAAX motif that is modified so that the
GTPases can associate with membranes (Coats et al., 1999). Initially discovered to
transform fibroblasts (Stacey and Kung, 1984) and differentiate PC12 cells (Satoh et al.,
1987), the Ras family of GTPases is now linked to several signaling events including cell
migration, guidance and outgrowth in neurons, and cell growth (Mor and Philips, 2006).
The Ras GTPases can be activated downstream of growth factors, are capable of
activating Raf and the MAPK cascade, and are often associated with positive effects on
growth or migration (Giehl et al., 2000). The Ras GTPases are also regulated
downstream of guidance cue receptors (Huber et al., 2003), and in some cases the Ras
GTPase effector proteins can regulate other GTPase family members to control cell
migration, including the Rho-family of GTPases (Ehrenreiter et al., 2005).
17
Ran GTPases
The Ran GTPases (Clarke and Zhang, 2001) are involved in nuclear import/export
(Gorlich and Kutay, 1999; Dasso, 2001), metaphase mitotic spindle assembly (Kalab et
al., 1999; Heald, 2000) and in the assembly of the nuclear envelope after mitosis (Hughes
et al., 1998; Hetzer et al., 2000). Ran-GDP is highly concentrated in the cytoplasm while
Ran-GTP is enriched in the nucleus. This segregation of GTP- and GDP-bound Ran is
controlled by the localization of the nuclear RanGEF, Regulator of chromatin condensation 1 (RCC1) (Klebe et al., 1995), and the cytosolic RanGAP (RanGAP1)
(Bischoff et al., 1994). Although these GTPases have not been reported to be targets of
extracellular guidance cues, they do have roles in regulating kinetichore MTs (Sillje et
al., 2006).
Arf GTPases
The ADP ribosylation factor (ARF) GTPase subfamily is divided into three
classes: class I (Arf1-3), class II (Arf4, Arf5) and class III (Arf6) (Donaldson and Honda,
2005), and are loosely related to the Arf-like (Arl) GTPases (Van Valkenburgh et al.,
2001). Aside from possessing switch I and switch II regions, the ArfGTPases are very
different from other GTPases. Class I and class II ARF GTPases are targeted to the golgi
by a conserved MXXE motif (Honda et al., 2005). ArfGTPases are myristoylated at their
amino-terminus, and this modification regulates their association with membranes when
GTP-bound (Haun et al., 1993; Franco et al., 1996). Arf6, which lacks the MXXE motif,
localizes to the plasma membrane when GTP-bound and to recycling endosomes when
18 GDP-bound (D'Souza-Schorey et al., 1998). Arf6 alters actin dynamics at the plasma membrane and is necessary for Rac1 cycling to the plasma membrane (D'Souza-Schorey et al., 1997; Radhakrishna et al., 1999). In addition, Arf6 activity leads to an increase in neurite outgrowth that is mediated by Rac1b (Albertinazzi et al., 2003), indicating that the ArfGTPases can influence axon outgrowth through their regulation of other GTPase families.
Rab GTPases
The Rab family of GTPases is the largest family, with over 30 members in mammals (Chavrier and Goud, 1999), and is the least characterized family. The cycling of GDP-bound and GTP-bound Rab GTPases controls several aspects of vesicle and organelle trafficking (Grosshans et al., 2006). The Rab GTPases are prenylated on their carboxy-terminal CAAX motif (Anant et al., 1998), similar to Ras GTPases, but are still able to distinguish between different subcellular locales in a yet-undetermined way.
Aside from vesicle and organelle trafficking, some Rabs are known to interact with the molecular motors (Echard et al., 1998), while others are linked to actin stress fiber and focal adhesion dynamics (Imamura et al., 1998). Interestingly, the RabGTPases are now being thought of as important regulators of cell migration, due to their dynamic regulation of membrane cycling (Jones et al., 2006). Moreover, the Rho-family GTPase
Rac1 can regulate the actions of the Rab5 GTPase downstream of EGF signaling
(Lanzetti et al., 2000), indicating that these families crosstalk in processes that regulate growth and cell migration.
19 Rho GTPases
The Rho family of GTPases has been extensively characterized, and are intricately involved in the cytoskeletal dynamics of endocytosis, cell motility, polarity, as well as axon and dendrite growth and guidance (Hall, 2005; Jaffe and Hall, 2005; Ridley,
2006). The Rho family is composed of over 20 proteins in mammals (Wennerberg and
Der, 2004). The downstream signaling events that regulate actin dynamics have led to a wealth of knowledge concerning the mechanisms used by RhoA, Rac1 and Cdc42.
RhoA has been linked to major changes in cytoskeletal dynamics, first described as a mediator of stress fiber formation downstream of the extracellular cue LPA (Ridley and Hall, 1992). One of the primary ways that RhoA leads to changes in actin is through its effector ROCK (Amano et al., 2000). RhoA activates ROCK, a kinase which activates
LIM kinase, which in turn phosphorylates and inhibits cofilin (Bamburg and Wiggan,
2002). ROCK is also capable of phosphorylating the myosin-binding subunit (MBS) of myosin phosphatase at multiple sites (Kawano et al., 1999), leading to its inactivation and a subsequent rise in the levels of phosphorylated myosin light chain (MLC) (Kimura et al., 1996), which is enriched at the leading edge of motile cells (Matsumura et al., 1998).
These actions of ROCK have been attributed to the formation of actin stress fibers and focal adhesions, (Nakano et al., 1999). RhoA can also activate the formin protein, mammalian diaphanous (mDia), and this activation is associated with filopodia formation and cell polarization (Faix and Grosse, 2006). The mDia protein can bind to MTs and promote their stability when activated downstream of RhoA (Palazzo et al., 2001b).
Signaling via RhoA is primarily attributed to inhibition of both cell motility and axon outgrowth. In support of this, high levels of active RhoA have been attributed to
20 neuronal growth cone collapse (Sahin et al., 2005; Wu et al., 2005; Gallo, 2006) as well
as the inhibition of cell motility and migration (Mills et al., 2005). These experiments
have revealed a great deal about the molecular mechanisms of RhoA activation, but they
are often based on RhoA overexpression or the use of CA or DN mutants of RhoA, and not local activation in cells. In fact, there are now reports indicating that active RhoA is highly enriched in the peripheral domain of dynamic growth cones (Nakamura et al.,
2005) and at the leading edge of migrating cells (Pertz et al., 2006), supporting the idea that RhoA is necessary for proper cell and growth cone motility and not just collapse or repulsive signalling. One report has even shown that the RhoA effector protein ROCK acts in conjunction with Rac1 in growth cones to assemble and disassemble point contacts, which aid the growth cone in its ability to extend (Woo and Gomez, 2006).
At the same time that RhoA was reported as a mediator of stress fiber formation,
Rac1 was reported as the mediator of membrane ruffling downstream of platelet derived
growth factor (PDGF) and bombesin (Ridley et al., 1992). Rac1 can exert its effects on
the actin cytoskeleton through its effector P21-activated kinase (PAK) (Eby et al., 1998;
Bokoch, 2003), activation of LIM kinase-1 (LIMK-1) and subsequent phosphorylation
and inactivation of cofilin (Yang et al., 1998), an actin depolymerizing protein. GTP- bound Rac1 also leads to the release of WAVE from its inhibitory complex, so that it can bind to the ARP2/3 complex to promote actin nucleation (Soderling and Scott, 2006).
Rac1 is also involved in a positive feedback loop with PI3K and PI(3,4,5)-trisphosphate at the leading edges of polarized structures, which promotes sustained extension of the actin cytoskeleton (Fukata et al., 2003). In addition to its well documented roles with
21 actin, the Rac1 effector PAK1 can phosphorylate and inhibit the actions of
OP18/stathmin, which normally destabilizes MTs (Wittmann et al., 2004).
Similar to Rac1, the small GTPase Cdc42 primarily acts through PAK (Bokoch,
2003), which leads to enhanced actin polymerization. Cdc42 is distinguishable from
Rac1 through the morphological structures to which it is linked, including the formation of actin structures like filopodia through its association with the Wiskott-Aldrich syndrome protein (WASP) family (Miki et al., 1998), which enhances actin polymerization with the ARP2/3 complex when active (Takenawa and Suetsugu, 2007).
Cdc42 also plays a major role in cell polarization through its activation of IQGAP
(Watanabe et al., 2004), which is part of a complex that locally captures MTs at the leading edges of cells and links them to leading edge actin. The polarity proteins Par-3, and Par-6, in conjunction with atypical protein kinase C (aPKC), form a complex with activated Cdc42 that establishes polarity through reorienting both the golgi and centrosome in migrating cells (Cau and Hall, 2005). This complex can also activate Rac1 through recruitment of its GEF Tiam1 (Nishimura et al., 2005). Localized Cdc42 signaling at the leading edges of migrating cells helps maintain a sustained directed migration that is important for chemotaxis and wound healing.
No one GTPase is solely responsibible for the enhancement or inhibition of cell motility or axon outgrowth. Expressing constitutively active Rac1 (G12V) or dominant negative Rac1 (S17N) in neurons often leads to the same net result: decreased axon outgrowth compared to wildtype neurons (Woo and Gomez, 2006). It is now being appreciated that there must be a balance of many GTPases to precisely regulate cell motility and axon outgrowth, each doing their part at the right time in the right place.
22 This precision requires signaling networks that can rapidly activate and inactivate the
GTPases when needed. This regulation can be found in the GEFs, GAPs, and GDIs that
act as the spatial and temporal regulators.
GTPase regulatory proteins and the role of GAPs
Rho family GTPases are ubiquitously expressed and utilized by a variety of
pathways to regulate cytoskeletal dynamics. GTPases are tightly regulated by GEFs and
GAPs, which promote the active and inactive states, respectively, of GTPases (Geyer and
Wittinghofer, 1997). The GTPases can also be sequestered by GDIs, which bind GDP- bound GTPases and sequester them in the cytoplasm (Dransart et al., 2005), and comprise
the main cytoplasmic pool of GTPases in a cell. While GEFs enhance the levels of active
GTPases in a cell, GAPs increase the levels of inactive GTPases. These GEFs and GAPs
have more restrictive expression patterns and subcellular localizations than the highly
ubiquitous GTPases they target. Even though they are more restricted in terms of tissue
type and localization, both GEFs and GAPs far outnumber their substrates, leading to the
idea that each GEF and GAP has a spatial and temporal niche to regulate GTPase
signaling. Aside from different catalytic domains, GEFs and GAPs regulate two sides of
the same process, and both have a wide array of functional domains to localize them to
sites of action and bring together signaling proteins. To understand how both of these
regulatory proteins control signaling events, a detailed understanding of their functional
domains is necessary. While there are too many GEFs to characterize here, a survey of
functional domains on GAP proteins will shed light on the complexity required to
spatially and temporally regulate GTPase signaling events.
23 There are just over 20 mammalian Rho-family GTPases, and greater than 80 Rho- family GAP proteins which regulate the same GTPases (Moon and Zheng, 2003). GAP proteins are characterized by a conserved GAP domain that catalyzes the hydrolysis of
GTP to GDP on GTPases, and contain a finger loop region with a conserved arginine that confers enzymatic function (Scheffzek et al., 1998). This arginine finger helps stabilize the GTP to GDP transition state, promoting hydrolysis of GTP’s gamma phosphate.
Although GTPases can undergo GTP hydrolysis at a low basal rate, GAPs increase this rate by several fold and can lead to rapid inactivation of GTPase signaling cascades
(Moore et al., 1992).
Similar to the various GTPase families, there are subfamilies of GAP proteins that are characterized by homology and shared GTPase substrates. The various families of GAPs contain several functional domains that confer specific localization, protein- protein interactions and regulatory regions (Bernards, 2003). This variety of domains between GAPs has led to a new level of signaling regulation referred to as the “GAP- ome” (Bernards and Settleman, 2004). As members of the RasGAP, RanGAP, ArfGAP,
RabGAP and RhoGAP subfamilies are characterized, it is becoming obvious that understanding the function of these GAPs requires a thorough analysis of how their domains regulate, localize, bind protein partners and are themselves regulated in cells.
RasGAP family functions, localizations and binding partners
The RasGAP family is a well characterized group of proteins (Scheffzek and
Ahmadian, 2005) whose actions are linked to cancer, synaptic plasticity, differention and cell growth. Many RasGAPs control signaling events that regulate cell growth and
24 migration, and these events are the consequences of not only their GAP activity, but protein-protein interactions and localization. One example is RasGAP which can bind to the F-actin associated protein filamin via its SH3 domain, and this interaction mediates myocyte cell growth and protein levels of the ribosomal S6 proteins and the kinase cdk7
(Lypowy et al., 2005). SYNGAP acts as a GAP towards Ras in vitro and associates with the post synaptic density 95 (PSD-95) protein (Chen et al., 1998; Kim et al., 1998).
SYNGAP heterozygous mice showed decreased long term potentiation (LTP) and decreased spatial learning compared to wild type mice (Komiyama et al., 2002), demonstrating a role in synaptic plasticity.
RanGAP family
The RanGAP proteins differ from other Ras superfamily GAPs because they do not use an arginine finger, or any amino acid, to stabilize the catalytic site when bound to
RanGTPases. Instead, a conserved glutamate that is part of the RanGTPase inserts into the active site to stabilize the reaction when bound to the GAP (Seewald et al., 2003).
The RanGAPs are localized to the nuclear pore complex either by a WPP motif in plants
(Jeong et al., 2005) or by sumoylation of the carboxy-terminus in animals (Matunis et al.,
1996). They also possess a central span of leucine-rich repeats, and a carboxy-terminal acidic domain (Seewald et al., 2003), which are implicated in protein-protein interactions.
This unique domain structure allows RanGAPs to spatially regulate RanGTPases in the cytoplasm (Bischoff et al., 1994) and promote a RanGTPase GDP/GTP gradient that is necessary for proper nuclear import and export (Gorlich and Kutay, 1999).
25 ArfGAP family
The ArfGAPs act primarily on the ArfGTPases and are intricately involved in
ER and golgi trafficking as well as actin dynamics (Randazzo and Hirsch, 2004). This subfamily is characterized by the necessity of a zinc finger in conjunction with two arginines to act as a GAP (Cukierman et al., 1995). Membrane association brings them in close proximity to the Arf GTPases, which associate with these membranes in a GTP- dependent manner to regulate coat assembly, trafficking, as well as maintaining the integrity of the golgi stack (Godi et al., 1999). Interestingly, the ArfGAP ARAP2 has both ArfGAP and RhoGAP domains, and has been shown to act as a GAP for both
ArfGTPases and RhoGTPases (Yoon et al., 2006), and can localize to and disrupt stress fibers and focal adhesions.
RabGAP family
The RabGAPs are a large family involved in vesicle trafficking and endo/exocytosis. Like their respective GTPases, there is still little known about the
RabGAPs. Interestingly, some reports have suggested that RabGAPs do not alter the function of certain RabGTPases (Rybin et al., 1996; Richardson et al., 1998), indicating
that either GDP/GTP cycling may not be important for some RabGTPases to function or
that the GAPs are ineffective at hydrolyzing GTP in the cell. These GAPs may also just
act as Rab-binding proteins in the cell.
26 RhoGAP family
The RhoGAP family is the largest group with over 80 potential members. This
family is involved in several cell processes (Moon and Zheng, 2003; Tcherkezian and
Lamarche-Vane, 2007), and has been extensively characterized in its involvement in
actin and MT signaling, mostly through their regulation of the GTPases RhoA, Rac1 and
Cdc42 (Peck et al., 2002; Watanabe et al., 2005). Similar to most other members of the
Ras superfamily of GAPs, the RhoGAP family has a conserved finger loop with an arginine finger that confers enzymatic function (Scheffzek et al., 1998). Their GTPase targets in vivo are influenced by their numerous domain structures and intense regulation by upstream regulators, and these functional domains confer membrane association, protein-protein interactions and enzymatic regions (Peck et al., 2002), (figure 1.4, borrowed with permission from Peck et al., 2002). Reflecting an importance to localize to certain signaling microdomains in cells, several independent domains have evolved to confer similar but not identical functions on different RhoGAP proteins. The C1
(Canagarajah et al., 2004), C2 (Ponting and Parker, 1996), pleckstrin homology (PH)
(Sakakibara et al., 2004), sec14 (Aravind et al., 1999), and the StAR-related lipid transfer
(START) (Strauss et al., 2003) domains are functional entities that each confer
association of membrane lipids or be targeted by lipids under the appropriate conditions.
In addition, RhoGAP proteins have many domains that bind to other proteins to
dictate which signaling complexes they are recruited to and their subcellular localization.
The src-homology 3 (SH3) domain is found in several RhoGAPs, including ARHGAP4
and the srGAPs (Wong et al., 2001), while the proline-rich and WW domains on RICH-1
(Richnau and Aspenstrom, 2001) and ARHGAP9 (Furukawa et al., 2001), respectively,
27 mediate protein-protein interactions in different ways. In addition, there are some
domains that confer different binding capabilities in different contexts and their target
binding motif is still unknown. The fes/fer/fps/cip4 homology (FCH) domain that is
found on ARHGAP4 and the srGAPs is one such example, and has been attributed to
actin association (Yeung et al., 1998), MT binding (Tian et al., 2000), as well as lipid
binding (Tsujita et al., 2006).
Well known for their ability to influence actin dynamics, the RhoGAP family regulates many processes that depend on actin, including growth cone and cell motility
(Wong et al., 2001; Barker et al., 2004). Tissue-specific cancer phenotypes can often be attributed to mutations in one or more RhoGAP proteins (Johnstone et al., 2004;
Goodison et al., 2005), indicating how important RhoGAP signaling is for proper
regulation of growth and motility. Moreover, RhoGAPs can localize to discrete areas of
cells to influence the actin cytoskeleton in different ways, including the golgi (Dubois
and Chavrier, 2005), focal adhesions and stress fibers (Burgstaller and Gimona, 2004;
Kawai et al., 2004), and at the leading edge of migrating cells (Lua and Low, 2004).
Several RhoGAP proteins can be regulated by other GTPase families, and the RhoGAPs
can modulate other families in turn. These crosstalking events have made it difficult to
streamline the understanding of signaling pathways. To understand RhoGAP regulation
of cell motility, axon outgrowth and other processes, actin dynamics must be considered
alongside of other events and not as the only factor that controls all these processes.
28 GAP protein mutations and developmental consequences
Due to the massive cellular signaling events that GAPs mediate, their
misregulation and mutations are linked to many serious disorders and developmental
abnormalities. The abundant p190RhoGAP exhibits GAP activity towards RhoA (Ridley et al., 1993) and is extremely important for normal development of the CNS. Knockout of p190RhoGAP results in mice that lack a corpus collosum, as well as incomplete closing of the optic fissure and hyperplasia of the retinal pigmented epithelium (Brouns et al., 2000). Defects in the oligophrenin-1 gene were linked to the cognitive impairment in patients with non specific X-linked mental retardation (Billuart et al., 1998). The oligophrenin-1 protein can act as a GAP towards RhoA, Rac1 and Cdc42 in vitro.
Patients with deletions in MEGAP/srGAP3 result in severe mental retardation (Endris et al., 2002), possibly due to defects in proper axon and dendrite outgrowth. Mutations and
deletions of the RhoA GAP, ARHGAP6, is attributed to microphthalmia with linear skin
defects (MLS), possibly through its effects on the actin cytoskeleton (Prakash et al.,
2000). Along with these well documented examples, several other examples of GAP
mutations and misregulation have been linked to many types of cancers (Crawford et al.,
2006; Kandpal, 2006; Ullmannova and Popescu, 2006).
Regulation of GAP proteins
While the GTPases can be referred to as the molecular switches that control
multiple signaling cascades, GAPs and GEFs are analogous to the power sources that
dictate the function of the GTPases. Much research is now being focused in
understanding how guidance cues and growth factors utilize GAPs and GEFs in a
29 temporal and spatial manner to harness the signaling potential of the GTPases. Ridley and Hall surmised that extracellular signals utilized the GTPases to alter the actin cytoskeleton (Ridley and Hall, 1992; Ridley et al., 1992), but the players involved upstream of the GTPases and the GAPs and GEFs are still not well known. While the
localization of GAPs and GEFs is critical for their function, there are additional
regulatory mechanisms. These include phosphorylation and dephosphorylation, protein-
protein interactions, lipid regulation, and potentially many more (Bernards and
Settleman, 2004). Overall, each of these contributes to the spatial as well as temporal regulation of GAP proteins.
Phosphorylation
Phosphorylation of GAP proteins is recognized as a common mode of regulation, and it is extensively used by guidance and growth factor receptors as well as by GTPase effectors to regulate other GTPase pathways through GAPs. Moreover, regulation via phosphorylation is a mechanism that is shared among most GAP families.
P190RhoGAP is phosphorylated by c-Src in response to epidermal growth factor receptor
(EGFR) activation, and this event activates p190RhoGAP to alter actin stress fibers
(Haskell et al., 2001). Activated plexin receptors can associate with and activate p190RhoGAP downstream of guidance cue signals (Barberis et al., 2005), demonstrating that guidance cue receptors utilize GAPs to locally alter cytoskeletal dynamics in growth cones. SYNGAP is a postsynaptic density-associated protein that targets H-Ras (Kim et
al., 1998). CAMKII can phosphorylate and inhibit SYNGAP when activated (Chen et al.,
1998), and this pathway has been linked to long term potentiation. Insulin signaling
30 causes the phosphorylation and subsequent inhibition of the Rab-family GAP AS160, and
this inhibition allows for GLUT-4 containing vesicles to cycle to the plasma membrane
(Sano et al., 2003). The Cdc42 and Rac1 GAP CDGAP can be phosphorylated in response to serum activation of the MAPK/ERK pathway, and this phosphorylation is
proposed to inhibit the activity of CDGAP (Tcherkezian et al., 2005). This is one
demonstration of how downstream signals from one GTPase cascade (Ras/MAPK), can
lead to changes in another GTPase signaling cascade (Rho family).
Protein-protein interactions
Aside from phosphorylation, protein-protein interactions are common ways to
regulate both position and activity of GAP proteins. The ability of guidance and growth
factor receptors to facilitate localized signaling to the GTPases requires the ability to
sequester and utilize GEFs and GAPs when needed. The intracellular domains of
receptors often directly recruit and activate GEFs and GAPs, or form multiprotein complexes that can recruit GEFs and GAPs and regulate their action. As mentioned
above there are several domains that can regulate protein-protein interactions. One of the
most studied is the SH3 domain, which binds to conserved proline rich regions. The SH3
domain of the srGAPs can bind to the proline rich region of the Robo receptor’s CC3
intracellular domain when Slit binds to the Robo receptor (Wong et al., 2001), and this
interaction activates the srGAPs’ GAP activity and leads to a local inhibition of Cdc42.
The srGAPs and ARHGAP4 also contain FCH domains that have been implicated in
binding to MTs (Tian et al., 2000), suggesting that this domain could recruit GAPs to
areas of MT cytoskeletal dynamics. Coiled-coil domains, also common among GAP
31 proteins, have the ability to oligomerize to form either homo or heterodimers. One example is the Rac1 and Cdc42 GAP RICH-1, which is capable of forming homodimers via its coiled-coil domain, an event that is necessary for RICH-1 to tubulate liposomes in conjuction with other functional domains (Richnau et al., 2004). All of these domains are important for regulation of both protein function and localization to signaling complexes.
In addition, while each domain has a common function, the domain combinations on
GAP proteins often confer synergistic effects that provide new ways to localize and recruit signaling proteins.
Other types of regulation
Second messengers, including calcium, can also regulate the activity of GAPs. In the presence of calcium, the RasGAPs CAPRI and RASAL are activated and associate with the membrane (Lockyer et al., 2001; Liu et al., 2005), and this is mediated through the calcium activation of the lipid binding C2 domain. Another way that GAPs can be controlled is through association with specific lipid moieties. P190RhoGAP is well known for its RhoA GAP activity, but when associated with certain phospholipids like phosphatidylserine (PS), its substrate preference and activity for RhoA can be inhibited, while activity towards Rac1 is activated (Ligeti et al., 2004). In addition, different types of lipids can have varying effects on different types of GAP proteins, potentially adding a new level of regulation in GTPase signaling (Tsai et al., 1989). Recruiting GAPs to specific membranes to inactivate GTPase pathways is important, because many GTP- bound GTPases associate with membranes through their lipid modifications (Heo et al.,
32 2006). Therefore, to inactivate a GTPase in a cell, a GAP protein must be recruited to the
same subcellular region, be it an organelle, vesicle, or lipid raft.
Although the regulation of GAPs and GEFs is still not well understood, it may be
one of the final links in connecting extracellular cues to the changes in the cytoskeleton.
In support of this, many guidance receptors are thought to signal directly to GAPs and
GEFs, which in turn locally regulate GTPase function (Huber et al., 2003). Although
there are still many signaling events that are still unknown downstream of GTPase signaling, there has been a great deal of work elucidating the signaling events downstream of RhoA, Rac1 and Cdc42. These GTPases regulate both the actin and MT
cytoskeletons, and many of these signaling events are conserved between migrating cells and axonal growth cones.
Actin and microtubule dynamics in growth cone and fibroblast motility
General actin dynamics
Actin dynamics have been of intense interest in understanding how growth cones and migrating cells rapidly alter their morphologies to traverse their environments (Dent
and Gertler, 2003; Ridley, 2006). The events that lead to actin polymerization and
depolymerization are tightly regulated by specific proteins that can sequester, stabilize,
sever, cap and crosslink actin filaments (Disanza et al., 2005). Actin polymers are
composed of different forms of actin monomer proteins: α-actin, β-actin and γ-actin
(Garrels and Gibson, 1976). These monomeric, globular (G)-actin proteins are
intrinsically asymmetric and form polarized filaments when polymerized (Reutzel et al.,
2004). ATP-bound G-actin is added to the plus ends of F-actin, and its addition in
33 combination with the loss of ADP-bound actin at the minus ends of F-actin determines the steady state of polymerization within cells (Korn et al., 1987). Evidence that F-actin was mostly assembled at the plus (barbed) end and disassembled at the minus (pointed) end came from experiments where labeled actin was incorported into F-actin in fibroblasts and then photobleached in small areas (Wang, 1985). These data showed that photobleached spots in F-actin moved from the plus end towards the minus end, reflecting the treadmilling of previously added actin subunits in response to each new subunit incorporated at the plus end. These experiments were done in fibroblasts where actin assembly was actively occurring. The plus end of F-actin is the most dynamic, undergoing the fastest rates of polymerization and depolymerization relative to the minus end (Pollard, 1986), and this dynamic nature makes it highly susceptible to proteins and drugs that modify the actin cytoskeleton.
Within cells, actin binding proteins (ABPs) regulate the rates of actin polymerization and depolymerization, and these events are heavily influenced by GTPase mediated signaling events (see above) (Ridley, 2006). Each ABP has a unique structure that confers function, and many different proteins can fall into specific categories based on how they affect actin. The F-actin severing protein ADF/cofilin severs actin filaments at the minus end and increases the amount of G-actin (Bamburg and Wiggan, 2002).
Profilin binds G-actin and can cooperate with several other ABPs to regulate actin filament assembly (Yarmola and Bubb, 2006). Certain ABPs are required to maintain specific actin morphologies in cells as well. Fascin is an actin filament crosslinking protein that is found along actin filaments in filopodia, and is required for highly crosslinked, straight actin bundles in these structures (Vignjevic et al., 2006). For a
34 growth cone or cell to move it must extend processes into its environment, and this often
requires not only polymerization of existing actin, but also nucleation of new actin
polymers. The actin related protein (ARP)2/3 can bind to F-actin in response to active
WAVE or SCAR/WASP proteins and nucleate new actin filaments at 70° angles to
existing filaments (Takenawa and Miki, 2001; Goley and Welch, 2006). Formin proteins,
like diaphanous (Dia), can also nucleate actin in the presence of profilin, and do so at the
plus ends of actin filaments (Kovar, 2006). Some ABPs are known to have multiple
functions, including the gelsolin family of proteins, which have been shown to sever, cap
and nucleate F-actin (Silacci et al., 2004) The net result of these proteins is to disrupt the
actin equilibrium in a spatial and temporal fashion downstream of regulated signals,
which leads to extension or retraction of actin-based structures in cells.
Several drugs and toxins have been shown to modify the actin cytoskeleton as
well. The cytochalasin drug family (cytochalasins A, B, C, D, E, H, and 21,22
dihyrocytochalasin-B) bind to F-actin barbed ends with high affinity and prevent F-actin
polymerization and depolymerization at the barbed ends (Cooper, 1987). The
latrunculins (A and B) bind and sequester monomeric actin and prevent polymerization
by decreasing the pool of available G-actin (Spector et al., 1989). Other actin modifiers
like jasplakinolide can alter the dynamics of G-actin and F-actin and effectively alter
actin dynamics (Bubb et al., 2000). One of the most popular toxins, the phalloidins, bind to actin filaments with high affinity and stabilize F-actin (Cooper, 1987). Due to its ability to bind F-actin, fluorescently-conjugated phalloidin has been used to visualize F- actin in several systems (Faulstich et al., 1988).
35 General microtubule dynamics
While dynamic actin controls processes that result in the guidance and migration of motile structures, the role of MTs was believed to be more passive. MTs serve important roles in cargo trafficking (Allan and Schroer, 1999; Palmer et al., 2005), polarity (Musch, 2004) and cytokinesis (Sato and Toda, 2004), and are now thought to be involved in more dynamics processes, including growth cone turning (Buck and Zheng,
2002), and cell migration (Watanabe et al., 2005). MTs are assembled from α-tubulin/β- tubulin heterodimers that assemble to form a polarized filament (Nogales and Wang,
2006). MT polymers are often enriched with GTP-bound tubulin at their plus ends, and this GTP-bound form is important for structural integrity, as GDP-bound tubulin at the plus ends of MTs is associated with the more MT catastrophes (Tran et al., 1997). MTs are often assembled from the centrosome in a minus to plus end direction, and often require the nucleating tubulin, γ-tubulin, and associated proteins (Heald and Nogales,
2002). As MTs grow, they undergo cycles of catastrophe (depolymerization) and rescue
(polymerization/stabilization) referred to as dynamic instability, and these events can be prevented or enhanced by drugs that act on tubulin and MTs or by proteins that modify the MT network (Nogales, 2001). Several drugs can stabilize or destabilize MTs, and have been used in the treatment of various cancers (Pellegrini and Budman, 2005).
Several of these drugs are widely used to study MT dynamics in migrating cells and growth cones, and two of the most common drugs are paclitaxel and nocodazole. The drug paclitaxel (also known as taxol), can bind to tubulin polymers and stabilize them, preventing depolymerization and catastrophes (Schiff and Horwitz, 1980). Another
36 widely used drug is nocodazole, which can severely destabilize MTs, leading to a
disolution of the MT polymers (Samson et al., 1979).
MTs that undergo high rates of rescue and catastrophe are considered dynamic,
and these dynamic MTs are often short lived in cells compared to stable MTs (Kreis,
1987). These dynamic MTs are often newly formed and are present in regions of a cell
that are highly motile, and can identified by the presence of a terminal tyrosine residue on
the carboxy-terminus of alpha-tubulin (Gundersen et al., 1984; Arregui et al., 1991;
Westermann and Weber, 2003). Over time, this region is detyrosinated and left with a
glutamate residue (Glu-tubulin), an event that is a consequence of a long-lived tubulin moiety (Gundersen et al., 1987), and these detyrosinated tubulin isoforms are highly
resistant to depolymerization and destabilizing drugs (Gundersen et al., 1987; Kreis,
1987). Other posttranslational modifications including phosphorylation, acetylation,
polyglycylation, polyglutamylation and palmitoylation can occur, reflecting populations
of MTs with different dynamics and binding characteristics in a cell (Westermann and
Weber, 2003). All but one of these modifications (acetylation), occur on the carboxy-
terminal tails of alpha and beta-tubulin, a region which is highly acidic due to several
glutamate residues (Westermann and Weber, 2003).
Dynamic MTs have been implicated in controlling growth cone turning in
response to an inhibitory cue (Challacombe et al., 1997; Kalil and Dent, 2005). They
have also been implicated in several aspects of cell migration (Watanabe et al., 2005),
and it is only recently that the signaling mechanisms controlling these events have begun
to be elucidated. Several proteins have been identified that alter MT dynamics, including
the microtubule associated proteins (MAPs) (Amos and Schlieper, 2005). These proteins
37 can be localized to discrete areas of polarized cells. Some well known markers for
neurons are the dendrite localized MAP2 (Caceres et al., 1986), axonal Tau (Brion et al.,
1988), and MAP1b, which is enriched in growth cones (Black et al., 1994). Although
these classical MAPs could be regulated by kinases and modulate MT dynamics,
GTPases did not seem to be influencing these processes. However, a fast growing family
of MAPs called the plus (+)-tip MT binding proteins (Akhmanova and Hoogenraad,
2005), can bind MTs in a dynamic fashion and potentially regulate MTs downstream of
GTPase signaling events (Wittmann et al., 2003). These proteins often associate with protein complexes that contain GTPase effector proteins and function to modulate MT dynamics (Wen et al., 2004; Wittmann et al., 2004). The end binding proteins (EBs), were found to track along MTs in a minus to plus direction, and cause stabilization and bundling of MTs when overexpressed (Bu and Su, 2001). They can also associate with the formin mDia and the +-tip protein adenomatous polyposis coli (APC) to stabilize
MTs downstream of RhoA activation (Wen et al., 2004). APC was initially described as a scaffolding protein in the Wnt/beta catenin pathway whose actions were dependent on its phosphorylation state (Ha et al., 2004). APC is capable of binding to and stabilizing
MTs in growth cones when its kinase, GSK-3β, is inhibited by nerve growth factor
(NGF) signaling (Zhou et al., 2004). In addition, APC can bind to IQGAP1, which links it to the peripheral actin cytoskeleton and directly activate the Rac1 GEF, APC- stimulated guanine nucleotide exchange factor (Asef) to control actin assembly (Akiyama and Kawasaki, 2006). While EB1 and APC are part of complexes that positively regulate cell motility and axon outgrowth, many other +-tip proteins have been shown to negatively regulate these processes. For example, the actions of the CLIP-170 protein
38 (CLASP) in conjunction with the motor protein dynein, were associated with increased
MT catastrophes in yeast (Grallert et al., 2006). In addition, the OP18/stathmin protein
can bind to and increase the rate of catastrophes when it is unphosphorylated (Wittmann
et al., 2004).
The role of these +-tip proteins is still unknown, but three general models have
been proposed to explain their functions: the Delivery model, Regulation model, and the
Search-Capture model (Vaughan, 2004). The delivery model argues that +-tip proteins
that track along MTs do not alter MT dynamics directly, but are only passengers that
target to the plus ends. This model could encompass plus-end directed motor proteins, yet many +-tip proteins have been shown to alter MT dynamics. The regulation model
argues that +-tip proteins directly alter the rate of catastrophes and rescues when bound to
the plus ends. Many +-tip proteins actions support this model, including EB1 and CLIP-
170, which have been shown to increase catastrophes when not able to bind to MTs
(Komarova et al., 2002; Tirnauer et al., 2002). Lastly, the search-capture model states
that the localization of +-tip proteins on MTs either mediate interactions with targets, or
are the result of subcellular tagets that have captured MT plus ends with preassembled
complexes. This model argues that the role of +-tip proteins is to bridge dynamic MTs to
subcellular targets like the actin cytoskeleton or membranes. This model has gained the most support over the last few years as more studies are examining the proteins that
assemble with the +-tip proteins. One example demonstrated that activated integrins at
the leading edge of migrating cells could capture and stabilize MTs through RhoA and its
effector mDia (Palazzo et al., 2004). Another study demonstrated that NGF signaling in
growth cones led to the association of APC with MTs and this interaction was important
39 for growth cone turning (Zhou et al., 2004), supporting the idea that activation of some
guidance receptors could capture and stabilize MTs by utilizing the APC protein.
Coordinated signaling regulated by Rho-family GTPases
Support for the idea that the actin and MT cytoskeletons are controlled by the
same upstream factors comes from the work examining MT +-tip protein complexes that
often have several ABPs as well. These multidomain complexes support the idea of the
search-capture model for the +-tip proteins, and have led to the ideas that complexes on
MT plus ends can act on actin as they grow and retract, or that components complexed to cortical actin can capture and stabilize dynamic MTs in response to some signal. Both ideas require the interaction of actin and MTs, but the order of events is still not well understood. One well studied event that bridges the actin cytoskeleton to MTs involves the IQGAP1/CLIP170/Cdc42-GTP complex. IQGAP1 is a Rho-family GAP that lacks a conserved arginine finger, making it unable to stimulate GTP hydrolysis but can still interact with with GTP-bound Rac1 and Cdc42. When Cdc42 or Rac1 are activated at the plasma membrane, they can bind to IQGAP1, and form a complex that attaches to cortical actin, via IQGAP1, and can now capture CLIP170 on dynamic MTs (Fukata et al., 2002a), as well as recruit APC to bind MTs (Watanabe et al., 2004). This complex is highly involved in cell migration and polarity, and seems to function similarly in other cell types. In neurons, the Lis1 protein can associate with the IQGAP1/CLIP170/Cdc42-
GTP complex downstream of calcium influx, and this event is necessary for proper cell migration due to a similar mechanism (Kholmanskikh et al., 2006). These studies on
GTPase signaling events demonstrate that many diverse pathways can act together when
40 locally stimulated downstream of a common activator or inhibitor. The GTPases are
potent regulators of many of these multitude pathways and their regulation by GAPs and
GEFs determine when, where, and how these events play out.
Cell migration and functional parallels to growth cone guidance
The pathways that control both actin and MT dynamics are utilized not only by
growth cones but also by migrating cells (figure 1.5, borrowed with permission from
Rodriguez et al., 2003). When the body is wounded, fibroblasts must migrate into the
wounded area so healing can occur. These events require both highly organized and
dynamic cytoskeletons, but also the ability to polarize in the right orientation. In
addition, polarized neurons must send growth cones and dendrites to different locations
in order to wire the brain correctly.
Neurons are highly polarized structures with axons and dendrites, which have
differential distribution of proteins, cytoskeletal elements, and functions. The events
which polarize and guide the axon to its final destination are still widely unknown, but
are thought to be regulated by GTPase signaling (Watabe-Uchida et al., 2006) and
disproportionate regulation of cytoskeletal elements (Bradke and Dotti, 1999). Both
axons and dendrites respond to cues that guide them to their proper locations, and as the
axon navigates its environment its growth cone functions as a micro-polarized
appendage. Cytoskeletal components and proteins are unequally distributed in growth
cones (Dent and Gertler, 2003) and lead to a highly responsive structure whose goal is to
sense the environment and either locally protrude or retract in specific directions. The growth cone of an axon can be divided into 3 zones: central (C), transition (T), and
41 peripheral (P) (Dent and Gertler, 2003). The C zone is characterized as being MT rich
and is thought to be more passive than the rest of the growth cone in mediating guidance
and outgrowth. The area around the C domain, where the majority of MTs stop
extending and where the majority of actin starts growing into the P zone, is referred to as
the T zone. The P zone is a highly motile area of the growth cone that contains the
majority of actin and a few dynamic MTs that have grown out from the C zone. Due to
the fact that it is the first region of the growth cone to interact with the environment, it is
thought that the majority of signaling to the cytoskeleton from extracellular cues occurs in the P zone. In support of this micro-polarized zoning of the growth cone, several actin associated proteins and MT associated proteins are sequestered within different zones of the growth cone (Dent and Gertler, 2003). These proteins regulate the dynamic actin and
MT cytoskeletons which are known to mediate axon outgrowth and guidance
(Challacombe et al., 1997; Buck and Zheng, 2002; Gallo et al., 2002).
Fibroblasts are also highly motile and can polarize under the appropriate conditions. Migrating fibroblasts have polarized structures with disproportionate amounts of proteins that convey discrete functions, and often form a broad leading edge that is focused in the direction of migration and a trailing rear (Ridley et al., 2003). The fibroblast leading edge is an area composed of highly dynamic actin and MT elements
(Wehrle-Haller and Imhof, 2003), and is an area with highly activated GTPases. Many of the GTPase modulated pathways that control actin and MT dynamics in growth cones are also thought to function in fibroblasts. There are now several lines of evidence that show how important the GTPases, actin, and MT cytoskeletons are for migration and polarization (Fukata et al., 2003; Charest and Firtel, 2007).
42 Even though the GTPases RhoA, Rac1 and Cdc42 are capable of activating several pathways, their actions are often attributed to specific morphological events
during directed cell migration and growth cone motility. The small GTPase Rac1 is
important for lamellipodia formation and membrane ruffling (Ridley et al., 1992), as well
as the ability of growth cones and fibroblasts to protrude and spread, thus leading to directional movement (Wittmann and Waterman-Storer, 2001). RhoA activity is activated at the leading edges and also in the trailing edges of migrating cells and is thought to be involved in both the cycling of point contacts that are necessary for migration and for the detachment of the trailing edge (Wong et al., 2006). Cdc42 activity has been attributed to the formation of filopodia and is necessary for correct polarization to occur in fibroblasts (Yang et al., 2006). Polarization in fibroblasts is not just the formation of leading and trailing edges, but also the reorientation of the microtubule organizing center (MTOC) and the golgi complex in front of the nucleus. This reorientation of the MTOC and golgi is thought to provide long term support of cargo and membrane to the leading edge of the migrating cell and these events have been linked to specific pathways downstream of Cdc42 (Palazzo et al., 2001b; Cau and Hall, 2005).
The idea of the golgi and MTOC dictating polarity is also shared in neuronal polarity
(Zmuda and Rivas, 1998). Studies also suggest that the MTOC and golgi can be reoriented in response to actin and MT dynamics as well as signaling events downstream of other GTPases (Magdalena et al., 2003).
The leading edge of fibroblasts contains several protein complexes important for actin polymerization, and it is where many of the GTPase are activated in response to extracellular cues (Kurokawa et al., 2005). Some of the more common cues found in
43 serum are LPA, EGF, and PDGF, which are known to stimulate GTPase signaling events at the leading edge (Chan et al., 1998; Suetsugu et al., 2003; Sugimoto et al., 2006).
There are also several guidance cue receptors expressed in growth cones that utilize these pathways as well, but are not present in fibroblasts. Although many of the pathways downstream of the GTPases are thought to be conserved between growth cones and fibroblasts, it is still unknown how different receptors in different cell types use these pathways. With the same GTPases, GAPs and GEFs expressed between neurons and fibroblasts, there must be some way different receptors/upstream regulators can utilize the same GTPase signaling networks to influence cytoskeletal dynamics. With the enormous number of GAPs and GEFs that exist, and their wide array of domain arrangements, it seems likely that the answer may lie with what the GAPs and GEFs are capable of being complexed with. Since both the leading edge of a fibroblast and the growth cone are similar in terms of their function, it is probable that GAPs or GEFs localized there may have a similar function in both systems even if recruited by different upstream receptors or signaling proteins.
This study focuses on the GAP, ARHGAP4, which was found to be expressed in both growth cones of dentate granule cells and NIH/3T3 fibroblasts. In particular, this study aimed to determine if the phenotype and localization of ARHGAP4 was similar in both neurons and fibroblasts, and use this as a springboard to assess the structure-function of ARHGAP4 in two distinct yet conserved model systems.
Previous work on ARHGAP4
44 The ARHGAP4 gene is located on the X chromosome (Xq28) and encodes a 965 amino acid protein in rat (called ARHGAP4/C1) and a 946 amino acid protein in human
(p115 hematopoietic RhoGAP4/ARHGAP4/RG4/C1). Initial studies examining the human ARHGAP4 found that it was expressed in hematopoietic cell types, including T cell, B cell and myeloid cell subtypes. Overexpressed ARHGAP4 disrupted actin stress fiber formation, indicating that it may regulate actin dynamics (Tribioli et al., 1996).
More recent work found that human ARHGAP4’s GAP domain has GAP activity towards RhoA in vitro (Christerson et al., 2002) while the GAP domain of rat ARHGAP4 showed activity towards RhoA, Rac1, and Cdc42 (Foletta et al., 2002). The experiemts demonstrating that ARHGAP4 could alter the actin cytoskeleton and had a functional
GAP domain support the hypothesis that it is regulating cytoskeletal dynamics via Rho- family GTPase signaling events.
Most of the reports on ARHGAP4 have utilized the human protein and its role in hematopoietic cell lines. Recently, ARHGAP4 was found to co-immunoprecipitate with the membrane associated protein HEM1 by way of its SH3 domain (Weiner et al., 2006) from HL60 cells. The HEM1/ARHGAP4 complex also included proteins involved in leading edge cytoskeletal dynamics and cell motility. Proteins in this complex included cyfip1/2, Mypt1, VPS34, diaphanous, AKT2 and the catalytic subunit of myosin phosphatase, but not WAVE2, which associates with HEM1 in a separate complex.
These proteins are known to regulate actin dynamics and may form a functional signaling complex that locally regulates GTPase signaling in polarized HL60 cells. It was also shown that knockdown of HEM1 led to a decrease of ARHGAP4 protein in HL60 cells, as well as drastic reorganization of the leading edges of polarized HL60 cells. When
45 HEM1 was knocked down, leading edges reorganized from broad lammelipodia to
filopodial-like extensions, indicating reorganization of the actin cytoskeleton.
Interestingly, when HEM1 protein was knocked down, HL60 cells could no longer activate Rac1 in response to fMLP, and the Rac1 effectors PAK1 and AKT failed to be activated. This work suggested ARHGAP4’s protein-protein interaction with HEM1 was an important regulator of leading edge morphology and GTPase signaling in HL60 cells, but did not discern what role ARHGAP4 may play in this process. Human ARHGAP4’s
SH3 domain has also been found to associate with MEKK1 via yeast two-hybrid screen and by coimmunoprecipitation (Christerson et al., 2002). MEKK1 is a large multidomain scaffold that is involved in cell motility and JNK signaling (Xu et al., 1996; Xu and
Cobb, 1997), and associates with the actin cytoskeleton at focal adhesions through its binding to alpha-actinin (Christerson et al., 1999). Although there did not seem to be any
phosphorylation of ARHGAP4 by MEKK1, this interaction was shown to disrupt
MEKK1 activation of AP-1 mediated transcription (Christerson et al., 2002).
The rat ARHGAP4 is highly homologous to the human (85%) and mouse (90%)
ARHGAP4 proteins and was found to be expressed not only in hematopoietic tissues
(spleen and thymus), but also in the developing and postnatal central nervous system
(Foletta et al., 2002). ARHGAP4 mRNA was expressed in the spinal cord, telencephalon, midbrain and rhombencephalon at embryonic day 18 (E18). In the adult rat, mRNA expression is prominent in the olfactory bulb, cerebellum, hippocampus, olfactory tubercle and pontine nuclei, (many of these structures are associated with adult neurogenesis and axon outgrowth). A northern analysis revealed that there may be multiple isoforms of ARHGAP4 in both fetal and adult rat brain. In agreement with this
46 a western blot of ARHGAP4 demonstrates that there are multiple immunoreactive bands.
Specific bands (50 kDa) are present at E18 and disappear in the adult, while others (36, and 120 kDa) are not present at early stages but are found in the adult (Foletta et al.,
2002). This differential expression pattern of ARHGAP4 isoforms is one level of regulation that is still not well understood, and may confer unique functions in the embryo and adult. To simplify the scope of this study the role of the full length
ARHGAP4 (120 kDa) was assessed.
The study by Foletta et al. also recognized that ARHGAP4 was not simply a cytosolic protein, but rather had a unique distribution pattern in rat NRK cells (Foletta et al., 2002). It was highly expressed in perinuclear regions and colocalized with the golgi complex. In a few cells, ARHGAP4 colocalized with perinuclear MTs and with MTs of cells undergoing cytokinesis. In PC12 cells, ARHGAP4 still had strong perinuclear distribution, but was recruited to the tips of forming neurites in the presence of NGF.
These data support the idea that extracellular cues may signal through ARHGAP4 to regulate the protrusion of neurites and potentially regulate axon outgrowth. Based on these previous reports we decided to determine if ARHGAP4 played a role in axon outgrowth. It also seemed important to determine what role ARHGAP4’s conserved domains contributed to its function. Based on work by Tribioli et al., Christerson et al., and Foletta et al., ARHGAP4 had a functional GAP domain and could disrupt actin stress fibers. However, considering that GAPs are themselves spatially and temporally regulated a structure-function analysis of ARHGAP4 needed to be performed.
Conserved domains of ARHGAP4
47 FCH domain, extended FCH domain and ARNEY domain
The FCH domain (figure 1.6, green bar) is characterized as an amphipathic helical
stretch with a conserved RAEYL motif. This motif is always found in the FCH domain, but some of the amino acids are substituted in a subset of proteins (dashed box, figure
1.6). The alanine is the least conserved, while the arginine, glutamate, tyrosine and
leucine are almost always conserved. Some proteins lack the arginine (but it is often
substituted by lysine), and a scant few proteins have the tyrosine substituted with
phenyalanine or histidine (S. pombe RGA7 and RGA8, as well as yeast RGD1, and
RGD2). This motif may represent either a critical residue in the amphipathic helix
required for lipid binding (Tsujita et al., 2006) or a conserved tyrosine phosphorylation
motif (predicted by netphos 2.0) that may be targeted in response to extracellular cues
(Yeung et al., 1998; Chang et al., 2002). The function of the FCH domain is not
understood, but has been implicated in binding to actin (Yeung et al., 1998), MTs (Tian
et al., 2000), and specific lipids (Tsujita et al., 2006). Although binding partners
identified for the FCH domain have not yet revealed a consensus binding motif, they do
share some common features. Tubulin subunits often bind basic regions of proteins
through their carboxy-terminal tails, which are enriched in acidic (negatively charged)
residues (Karabay and Walker, 2003). Second, lipid moieties implicated in FCH domain
binding are often hydrophobic entities with negatively charged side chains, and include
phosphatidylserine (PS) and phosphatidylinositol (PI[4/5]P2) (Tsujita et al., 2006). The
FCH domain is highly alpha-helical with several interspersed, positively-charged
residues, and these residues often extend beyond the conserved FCH domain into a region
now recognized as the extended FCH domain (eFCH) (Tsujita et al., 2006), (figure 1.6,
48 orange bar). It is plausible that either an acidic residue binding pocket or specific
negatively charged lipids could bind to the positively charged residues of the FCH domain and its extended regions.
FCH domain containing proteins are categorized in three classes (Greer, 2002).
One class includes the protein kinase FCH domain proteins, which include Fps/Fes and
Fer, and contain a functional kinase domain in conjunction with an amino-terminal FCH
domain. The second and largest group is the adaptor family of FCH domain proteins,
which often have amino-terminal FCH domains in conjunction with carboxy-terminal
SH3 domains. This family includes the syndapins, amphiphysins, endophilins, CIP4 and
Cdc15. Cdc15 was first described in S. pombe as an FCH domain protein that associated
with acto-myosin contractile ring during cytokinesis, (Fankhauser et al., 1995). Cdc42
interacting protein 4 (CIP4) is an FCH domain containing protein which also contains a
COOH-terminal SH3 domain. It has been shown to bind directly to MTs and to specific
lipid moieties through its FCH domain (Tian et al., 2000; Tsujita et al., 2006). Formin
binding protein 17 (FBP17) and CIP4 bind directly to specific lipids through their FCH
and juxtaposed domains and can bend lipids in vitro (Tsujita et al., 2006), similar to the
function of BAR domains, which can sense curved membranes (Blood and Voth, 2006).
The last family is the RhoGAP family of FCH domain proteins, of which the srGAPs and
ARHGAP4 compose the main members. The srGAPs are also known by other names:
srGAP1/ARHGAP13, srGAP2/Formin-binding protein 2 (FBP2)/Formin-binding protein
27 (FBP27), and srGAP3/ARHGAP14/WAVE-associated Rac GTPase-activating protein
(WRP)/MEGAP.
49 While the GAP and SH3 domains of the RhoGAP family are better
charactrerized, the FCH domain has not been well characterized in this family. Two
simple possibilities may arise from having an FCH domain juxtaposed to a GAP domain.
The function of the FCH domain may be independent from that of the GAP and SH3 domains, or the FCH domain may function in a manner which confers novel functions for the GAP and SH3 domains. Experiments with the FCH-SH3 containing proteins have demonstrated that these domains can function together to bind membranes and dynamin to regulate endocytosis (Kamioka et al., 2004). It may also be possible that the FCH domain confers new abilities to the GAP domain as well, possibly by bringing new proteins in close proximity or by localizing the protein to regions not before accessible.
Interestingly, the FCH domain is usually found in tandem with a coiled-coil domain, and
these domains are thought to be functionally similar to the BAR domain, which has the
ability to dimerize and either bind to or bend specific lipid moieties.
In a small family of proteins, including the srGAPs and ARHGAP4, a conserved
motif referred to as the ARNEY domain has been reported (Coyle et al., 2004). This
domain is found carboxy-terminal to the FCH domain (figure 1.6, blue bar) and has not
yet been attributed to any function. Although there has been no reported function for this
domain, due to its proximity to the FCH and eFCH domains the ARNEY domain is
thought to function in tandem with these other domains to regulate events. ARHGAP4’s
FCH domain lies between amino acids 24-125, its extended FCH domain lies between
amino acids 126-288, its ARNEY domain is located between amino acids 244-283
(Coyle et al., 2004) (figure 1.6). ARHGAP4 may also contain coiled-coil domains
50 between amino acids 42-62, 144-164, and 472-495 (predicted by Ensemble, not shown on
alignment).
RhoGAP domain
The GAP domain enhances the hydrolysis of GTP to GDP on Rho family
GTPases. The GAP domains of Rho-family GAPs differ from other families of GAPs
(like the ARFGAPs, RanGAPs, etc), but nearly all GAP domains contain a conserved
finger loop with a protruding arginine residue (Scheffzek et al., 1998). From the crystal
structure of Ras in complex with the GAP domain of p120GAP, it has been proposed that
this arginine finger neutralizes charges during a GTPase’s GTP to GDP transition state
(Scheffzek et al., 1997), and supports the Arginine finger hypothesis. The arginine finger hypothesis argues that the rate of GTP hydrolysis is enhanced when an arginine inserts itself into the active site of a GTPase and stablilizes the transition state, and 3 criteria have arisen that dictate what constitutes an arginine finger (Scheffzek et al., 1998). First, the arginine is conserved among subfamily members. Second, it can not be replaced, even with similar amino acids like a lysine. Third, substitution of this arginine should result in significant decrease in GAP mediated GTP hydrolysis while still maintaining protein/protein interactions. ARHGAP4’s GAP domain (amino acids 527-714) is most homologous to the srGAPs, and its arginine finger is located at residue 562 (figure 1.7).
SH3 domain
The SH3 domain is a well known protein-protein binding domain found in several kinases as well as many actin binding proteins (Mayer, 2001). It can occur anywhere in a
51 protein sequence, unlike the FCH domain, and contains conserved glycine, tryptophan and proline residues (figure 1.8). The crystal structure of avian alpha-spectrin revealed that the SH3 domain was composed of 5 antiparallel beta-strands which comprise 2 beta- sheets that are perpendicular to each other (Musacchio et al., 1992). The conserved proline residue is important for overall structure due to its role in inducing one of the characteristic turns. The SH3 domain is well characterized and found in several proteins.
It can bind to the PxxP motif found in several proteins, and the selectivity of an SH3 domain for its PxxP ligand has been extensively studied using PxxP motifs from several different proteins (Tong et al., 2002). However, many proteins with SH3 domains can bind to several different proteins with PxxP motifs, supporting the idea that while the
SH3-PxxP interaction is promiscuous, each interaction may have a specific function. In support of this idea, ARHGAP4’s SH3 domain was found to associate with MEKK1
(Christerson et al., 2002), HEM1 (Weiner et al., 2006), and Applp1 (Scott Young, personal communication). Two of these interactions have been shown to be functionally important, each in different cell types. ARHGAP4’s SH3 domain lies between amino acids 770-824 and is most homologous to the SH3 domains of the srGAPs (figure 1.8).
Proline rich (PxxP) domains
Proline rich regions which follow the motif PxxP, act as protein-protein interaction motifs that bind to SH3 domains. Although the PxxP sequence is considered the minimal binding requirement, proline rich motifs often resemble class I SH3 ligands
(+xΦPxΦP), or class II ligands (ΦPxΦPx+), each which bind to SH3 domains with similar affinities, but in opposite orientations (Φ aromatic, x any, + basic, P proline)
52 (Tong et al., 2002). The PxxP motif interacts with the SH3 domains of proteins using the
carbon backbone on the proline residues and the conserved basic residue (Yu et al.,
1992). Since almost all PxxP motifs have the same basic residue and carbon backbones,
specificity of the SH3-PxxP interaction has been a complicated issue. This lack of
specificity can give rise to novel functions in the same protein, depending on where it is
localized in a cell and what neighbors are available for binding. As discussed above for
the SH3 domain, this ability for PxxP motifs to bind to several SH3 domains may give rise to novel functions in different cell types. ARHGAP4 contains 3 proline rich regions in its extreme carboxy-terminus, at positions 877-80, 944-50, and 955-8. Although these regions contain the minimal PxxP motif, they lack the aromatic and basic residues that would classify them into specific classes.
Research goals
The main focus of this project was to determine if ARHGAP4 had any function in the central nervous system. If so, did it have any role in axon outgrowth and by what mechanism? Since ARHGAP4 could have induced a phenotype by any or all of its domains, I made it a goal to assess ARHGAP4’s pheneotype using a structure-function approach. I also used a second model system (NIH/3T3 cells) that has many similarities to growth cones in order to see if ARHGAP4 had parallel functions in other cell types.
From these studies, I have generated data to support the hypothesis that ARHGAP4 functions to inhibit axon outgrowth and cell motility in dentate granule cell axons and
NIH/3T3 fibroblasts, respectively. These data show that the GAP activity of ARHGAP4 is necessary for this inhibition in both model systems and may function in a localized
53 fashion with growth cones and the leading edges of migrating cells. Targeting of
ARHGAP4 to growth cones and the leading edges of NIH/3T3 cells has been mapped to its first 71 amino acids, which contains the highly conserved RAEYL motif which is found in FCH domains. These data support a model in which ARHGAP4 is targeted to growth cones and leading edges via its FCH domain, and its GAP activity is dependent on some protein-protein interaction at these peripheral regions, leading to localized signaling.
54
55 Figure 1.1: Hippocampal anatomy and pathways
The mature Hippocampus contains 3 morphologically recognizable regions,
which are the dentate gyrus, cornu ammon horn (CA) 1 and 3. Axons from the entorhinal
cortex project to dentate granule cells (forming the perforant pathway), dentate granule
cell axons project to CA3 pyramidal cells (forming the mossy fiber pathway) and CA3 pyramidal cells project to the CA1 pyramidal cells (forming the schaffer collateral
pathway). The axons of the various cell types in the hippocampus project reproducibly to the same regions. The dentate gyrus can be subdivided into specific regions. The
molecular layer contains granule cell dendrites. The dentate granule cell layer (GCL)
contains the granule cell somas and the hilus (H) is the region where granule cell axons
(mossy fibers) project through before compressing into a narrow band called the stratum
lucidum (SL) that lies above the CA3 pyramidal cell layer. The SL contains the proximal
dendrites of the CA3 neurons which form synapses with mossy fiber axons. The border
between the GCL and the H is called the subgranular zone and is where new neurons are
generated in the adult.
56
57 Figure 1.2: Common guidance cues and receptors found in the hippocampus utilize various GTPases to effect growth cone guidance
Guidance cues activate specific receptors that recruit proteins to specifically target
GTPases of multiple families, and locally regulate cytoskeletal dynamics in growth cones. Slit signaling through its receptor Robo can recruit the slit robo GAPs (srGAPs) to inhibit Cdc42 activity. Ephs and ephrins can increase RhoA activity via the GEF
Ephexin. Semaphorins bind to neuropilins and/or plexin receptors to control GTPase signaling events and mediate cytoskeletal changes. Class 3 semaphorins can form a tertiary complex with neuropilin receptors. Plexin-A and plexin-B receptor cytoplasmic domains have GAP activity towards R-Ras, which is mediated by an interection with
Rnd1. The p190RhoGAP can be recruited to plexin-B receptors to downregulate RhoA.
58
59 Figure 1.3: The GTPase cycle and role of GAPs, GEFs and GDIs
GTPase cycle through GTP and GDP bound states. GTPases bound with GTP are able to associate with effector proteins, but will slowly hydrolyze this GTP to GDP over time and this process is irreversible. GDP-bound GTPases can either be sequestered by guanine dissociation inhibitors (GDIs) which sequester and localize GTPases to the cytoplasm, or come interact with guanine nucleotide exchange factors (GEFs) which remove the GDP from GTPases and allow free GTP to bind to the GTPase. The slow rate of GTP hydrolysis can be sped up drastically by GTPase activating proteins (GAPs) which stabilze the GTP to GDP transition state. Reprinted from TRENDS in Cell
Biology, 14, Bernards and Settleman, “GAP control: Regulating the regulators of small
GTPases,” 377-85, 2004, with permission from Elsevier.
60
61 Figure 1.4: Rho-family GAP proteins and their domains
Rho-family GAP proteins contain a wide assortment of functional domains.
Some domains can bind to lipids and include the cysteine-rich phorbol ester binding (C1) domain, calcium dependent lipid binding domain (C2), sec-14 like domain (sec14), eps15 homology (EH), plecstrin homology (PH), and START domains. Others are reported to have multiple binding partners like the fes/fer/cip4 homology (FCH) domain, which can interact with actin, microtubules and lipids. Some domains can bind to other GTPases, like the Ras-binding domain (RBD) and Ral binding domains. Domains like the src homology 3 (SH3), proline rich domains (P), WW domains, coiled coil (CC), IQ domains and src homology 2 (SH2) domains are known to facilitate protein-protein interactions.
Domains can also confer novel functions and enzymatic activities, including RhoGEF domains, ARFGAP domians, myosin motor domains, kinase domains and inositol phosphatase domains. Reprinted by permission of Federation of the European
Biochemical Societies from “Human RhoGAP-domain containing proteins”, by (Peck et al.), FEBS Letters, (528) 27-34, 2002.
62
63 Figure 1.5: Actin and microtubule cytoskeletons in migrating cells and growth cones
A) Migrating cells have several types of actin organization, and include parallel
actin in filopodia, branched actin in lamellipodia, and actin that lines the cortex. Parallel
actin filaments are also organized into stress fibers in cells, and the actin cytoskeleton can interact with the MT cytoskeleton through crosslinking proteins. MTs grow out from the centrosome/MT organizing center (MTOC) and can invade leading edges, trailing edges,
regions with stress fibers and regions of high cortical actin. B) Growth cone peripheral
(P) domains contain parallel actin bundles in filopodia, interspersed lamellipodia, and
dynamic MTs. The central (C) domain is enriched with more stable MTs, while the
transition (T) domain is the region where the majority of MTs and actin segregate.
Similar to migrating cells, dynamic MTs can interact with actin, and can grow out on
actin filaments as well as be forced back by actin retrogade flow. (Other abbreviations
and symbols: convergence zone [CZ], lammelipodia [LA], plus ends of actin or
microtubules [+]). Reprinted by permission from Macmillan Publishers Ltd: Nature Cell
Biology, (Rodriguez et al., 2003)
64
65 Figure 1.6: ARHGAP4’s amino terminus contains FCH, extended FCH and ARNEY domains
ARHGAP4 amino acids 1-289 were aligned with other FCH domain containing proteins. The FCH domain is predicted to be composed of amino acids 24-125 (green), while the extended FCH domain (orange) extends carboxy-terminal after the FCH domain from amino acids 126-288. The ARNEY domain (blue) is a small conserved motif that lies carboxy-terminal to the FCH domain and within the eFCH domain between amino acids 244-283. Alignments were made with Clustal W, and show invariant amino acids (*), strang conservation of groups (:), and weak conservation of groups (.). Dashed box shows the RAEYL motif of the FCH domain.
66
67 Figure 1.7: ARHGAP4 GAP domain alignment
ARHGAP4’s GAP domain (amino acids 527-714) is homologous to other Rho- family GAP domains. The GAP domain of ARHGAP4 is most homologous to the srGAPs and both the srGAPs and ARHGAP4 are highly homologous to the c. elegans
F12F6.5 protein and its GAP domain. ARHGAP4 contians an invariant arginine finger at position 562 (^), indicating it may possess enzymatic GAP activity.
68
69 Figure 1.8: ARHGAP4 SH3 domain alignment
ARHGAP4 contains an SH3 domain between amino acids 770-824 that is highly homologous to the SH3 domains of the srGAPs. PACSIN1 and CIP4 SH3 domains are more distantly related, but still contain the conserved residues that define the SH3 domain
(blue, *). (*) fully conserved amino acid, (:) strong conservation of groups, (.) weak conservation of groups.
70 Chapter 2
ARHGAP4 is a Spatially Regulated GTPase Activating Protein (GAP) Expressed in NIH/3T3 Fibroblasts and Dentate Granule Neurons
71 Summary
GAPs catalyze the hydrolysis of GTP to GDP on GTPase proteins, promoting
their inactive state. GTPases are expressed ubiquitously and are regulated by GAPs,
GEFs and GDIs. Many GAPs target common GTPases, but do so in different subcellular locales to induce unique signaling events. GAP-regulated GTPase control is often a consequence of GAP protein levels, subcellular targeting, and activation by upstream signaling events. Here we describe the spatial regulation of the Rho-family GAP
ARHGAP4 using a structure-function approach. ARHGAP4 is highly localized to the leading edges of NIH/3T3 fibroblasts and to growth cones of dentate granule neurons. Its targeting to these regions is dependent on its amino-terminal FCH domain and requires an intact actin cytoskeleton. In addition, ARHGAP4 associates with the leading edge actin cytoskeleton in migrating NIH/3T3 cells and this association may contribute to its
function in both cell migration and axon outgrowth.
Introduction
The signaling events by which Rho family GTPases regulate actin and
microtubules (MTs) in migrating cells and neuronal growth cones have been extensively
studied since Ridley and Hall first demonstrated that RhoA and Rac1 mediated the
formation of stress fibers and membrane ruffles respectively in response to extracellular
cues (Ridley and Hall, 1992; Ridley et al., 1992). Since that time many families of
GTPases have been characterized and attributed to multiple cell processes, including nuclear import/export, vesicle and organelle trafficking, transcriptional regulation, cell growth and cell division (Mitin et al., 2005). However, the most attention has focused on
72 the regulation of cytoskeletal dynamics mediated by the Rho family GTPases RhoA,
Rac1 and Cdc42 (Burridge and Wennerberg, 2004; Raftopoulou and Hall, 2004;
Wennerberg and Der, 2004). RhoA, Rac1 and Cdc42 are often ubiquitously expressed
and can associate with the plasma membrane when activated via a motif in their carboxy
tails (Heo et al., 2006). The majority of the cytoplasmic GTPase pool is sequestered by
Rho family Guanine nucleotide Dissociation Inhibitors (RhoGDIs) (Dransart et al.,
2005). Due to their ubiquitous nature, the GTPases are under tight control in cells by
Guanine nucleotide Exchange Factors (GEFs) and GTPase Activating Proteins (GAPs), which enhance the transition to their active or inactive states respectively (Geyer and
Wittinghofer, 1997; Bernards, 2003). The various GEFs and GAPs are far greater in number than the GTPases they act on, reflecting unique roles for each GEF and GAP in the regulation of GTPases. In addition, Rho family GAP proteins are often large multidomain proteins, with the differential combination of domains conferring novel functions to each GAP protein.
Here the Rho family GAP protein ARHGAP4 is assessed with respect to its functional GAP domain (Foletta et al., 2002), amino-terminal fes/fer/fps/cip4 homology
(FCH) domain and carboxy-terminal src homology 3 (SH3) domain. Previous work has demonstrated that ARHGAP4 is expressed in hematapoietic cell lines (Tribioli et al.,
1996) and the developing and postnatal central nervous system (Foletta et al., 2002). Its overexpression has been linked to a loss of stress fibers in cultured SAA fibroblast cells
(Tribioli et al., 1996), indicating a role in regulating actin dynamics. ARHGAP4 localizes to several subcellular locales in rat NRK cells, including the golgi apparatus,
perinuclear MTs and the extending tips of NGF treated PC12 cells (Foletta et al., 2002).
73 More recent evidence has shown that ARHGAP4 can associate with both HEM-1 in
HL60 cells (Weiner et al., 2006) and MEKK1 (Christerson et al., 2002), by way of its
SH3 domain.
To understand how ARHGAP4’s localization may affect its role in neurons and migrating cells, a structure-function approach was used to assess the role of the FCH,
GAP, and SH3 domains. While there have been several reports on the role of the FCH domain, there is not yet a consensus. It has been implicated in binding to actin (Yeung et al., 1998), MTs (Tian et al., 2000), and to specific lipids moieties (Tsujita et al., 2006).
Here it is demonstrated that ARHGAP4 localizes to the leading edges of NIH/3T3 cells and to growth cones via its FCH domain, and this localization at the leading edges of fibroblasts and growth cones may depend on tethering to the peripheral actin cytoskeleton.
Materials and Methods
Generation of constructs
ARHGAP4 cDNA was generated by polymerase chain reaction (PCR) using the pεMTH-ARHGAP4 plasmid as a template (Foletta et al. 2002). Specific primers (5’-
CTAGGCGGCCGCATGGCGGCGCACGGGAAGTTGCGG-3’; 5’-
GCTAGAATTCGACTGGCTTGCGAGTTGAATCTGG-3’) were used to insert NotI and EcoRI restriction sites (bold) into the 5’ and 3’ ends, respectively, of the ARHGAP4 cDNA. The NotI/EcoRI fragment was then subcloned into the multiple cloning site of the pCMV-Tag 4A vector plasmid (Stratagene, La Jolla, CA) to drive expression of a full-
74 length ARHGAP4 (encoding amino acids 1-965) fusion protein with a carboxy-terminal
FLAG epitope [(1-965)-FLAG].
To generate the vector encoding a truncated ARHGAP4 fusion protein (72-965)-
FLAG, specific PCR primers (5’-CATAGCGGCCGCATGGAACGCTTTACTAG-3’;
5’-GCTAGAATTCGACTGGCTTGCGAGTTGAATCTGG-3’) were used to generate
NotI/EcoRI fragments (bold) that were then subcloned into pCMV-Tag 4A as above. To generate the vector encoding a truncated ARHGAP4 fusion protein (1-770)-FLAG, specific PCR primers (5’-
CTAGGCGGCCGCATGGCGGCGCACGGGAAGTTGCGG-3’; 5’-
GCTAGAATTCAGCCTCCACAACTCCCTCG-3’) were used to generate NotI/EcoRI fragments that were then subcloned into pCMV-Tag 4A as above.
To generate the vector encoding amino acids 1-71 of ARHGAP4 fused to a carboxy terminal EYFP protein (1-71)-EYFP, PCR primers (5’-
AGAAAGATCTATGGCGGCGCACGGG-3’; 5’-
GGACGAATTCAGTATGTTCCAGCAGCAT-3’) were used to introduce BglII and
EcoRI restriction sites into the ARHGAP4 cDNA for subcloning into the pEYFP-N1 vector (Clontech, Mountainview, CA). The ARHGAP4 vector encoding (1-965)-FLAG was used as a template for site-directed mutagenesis to mutate arginine 562 to an alanine
(R562A) in the full-length protein. The Stratagene QuikChange Multi Site-Directed
Mutagenesis Kit (Stratagene #200514) was used according to the manufacturer’s protocol, and mutant cDNA was generated using the 5’-phosphorylated primer (5’-
CTGCAACATGAAGGCATCTTCGCGGTATCAGGTGCCCAGG-3’, base pair
75 changes shown in bold). All plasmids were purified using the Qiagen Maxi Plasmid Kit
(Qiagen, Inc., Valencia, CA) and modifications were verified by sequencing.
The GAP domains (amino acids 474-743) of wild type ARHGAP4 (GST-WT-
GAP) and R562A (GST-R562A-GAP) were generated by pcr using (1-965)-FLAG and
(R562A)-FLAG as templates. Primers: 5’-
GCAGAATTCCTGCAGGCCAAGCATGAAAAGCTCCAG-3’, and 5’-
CATAAGCTTCTAGCTCTCCAACTGGCCATCCCCCAG-3’. Novel sites EcoRI and
HindIII in bold were introduced to clone into the GST-fusion expression vector pET-41a
(Novagen). GST-1-289 was generated by PCR using the 1-289-EYFP as a template.
Primers: 5’-GACGAATTCATGGCGGCGCACGGGAAG-3’, and 5’-
GAGAAGCTTGGCAATCCATGAGGTCCAAGAT-3’, with novel sites EcoRI and
HindIII in bold introduced novel restriction sites to clone into the GST-fusion expression vector pET-41a (Novagen). Recombinant GST and GST-1-71 proteins were a generous gift from the lab of Matthias Buck (Case Western Reserve University).
Protein purification
Protein purification was performed as previously described (Tong et al., 2005).
Vectors encoding GST-WT-GAP and GST-R562A-GAP were transformed into One
Shot® BL21(DE3) bacterial cells (Invitrogen). Positive colonies were inoculated in LB
+ 50 μg/ml Kanamycin and grown overnight at 225 rpms, 37°C. Cultures were diluted
1:100 in fresh LB + Kanamycin, grown for 3 hours, then induced with 0.1 mM IPTG and grown overnight at 225 rpms, 30°C. Purification of GST-1-289 was performed similarly, except that cultures were grown at 37°C for 4 hrs after IPTG induction instead of
76 overnight. The cells were pelleted and resuspended in GST-binding buffer (50 mM
phosphate buffer pH 6.8, 100 mM NaCl, 4 mM DTT, 4 mM MgCl2) + sigma protease inhibitor cocktail, then sonicated on ice. Insoluble debris was cleared by centrifugation and the supernatant fractions were filtered through a 0.22 micron filter. GST-Bind™
Resin (Novagen) was equilibrated in GST-binding buffer then combined with the resulting lysates for 20 min. at RT. Lysates were removed by gravity filtration on a
column and the beads were washed 3x in GST-binding buffer. GST-fusion proteins were
eluted off the beads with (100 mM Tris pH 8.5, 20 mM glutathione), and verified by
SDS-PAGE and coommassie staining.
In vitro GAP assay
The RhoGAP Assay Biochem Kit #BK105 (Cytoskeleton) was used to determine the relative GAP activity of both GST-WT-GAP and GST-R562A-GAP recombinant proteins. Briefly, GST-WT-GAP, GST-R562A-GAP, RhoA-His, Rac1-His, and Cdc42-
His proteins were diluted to 50 μM in nanopure water at 4°C. Reactions were set up in triplicate with GST-WT-GAP or GST-R562A-GAP + RhoA-His, Rac1-His or Cdc42-
His, as well as reactions with either GTPase alone or the GST-WT-GAP or GST-R562A-
GAP alone. Reactions were combined with 1x Reaction Buffer in a 96 well plate on ice, and GTP was added to each well at a final concentration of 200 μM. The plate was shaken at 200 rpm for five seconds and then incubated at 37°C for 20 min. At the end of the reaction, 120 μl of Cytophos reagent (Cytoskeleton) was added to each well. The reactions were incubated for ten minutes at RT and then the absorbance was read at 650
77 nm to assay the level of GTP hydrolysis. Reactions containing 1x Reaction buffer +
Cytophos reagent only were used as background controls.
Targeting assay
NIH/3T3 cells or dissociated granule neurons were transfected with ARHGAP 1-
965-FLAG, R562A-FLAG, 1-770-FLAG, 72-965-FLAG or 1-71-EYFP and cultured for
an additional 48 hours before fixation with 4% paraformaldehyde. Cells and neurons
were stained and visualized for FLAG expression or native EYFP expression. NIH/3T3
cells were measured for fluorescence intensity at the protrusive tip of an extending
process and 20 μm from the tip using ImageJ software. Localized fluorescence was expressed as a ratio of (tip intensity/intensity 20 μm from the tip). In neurons,
fluorescence intensity was measured at the leading edge of growth cones and 40 μm from
the growth cone in the axon and expressed as a ratio as above.
NIH/3T3 culture
NIH/3T3 cells were obtained from the American Type Culture Collection (ATCC,
cell line CRL-1658) and cultured according to ATCC protocol using Dulbecco’s
Modified Eagle’s Medium (DMEM) supplemented with 10% bovine serum (BS).
Dissociated Neuron culture
The dentate gyrus was microdissected from 200 μm hippocampal slices prepared
from postnatal day 5-7 rats as previously described (Butler et al., 2004). After
microdissection, dentates were rinsed once in Neurobasal (supplemented with B27,
78 glucose and L-glutamine) and then dissociated in Neurobasal/B27 with papain (2 ug/ml
at 15-23 U/μg) for 30 minutes at RT. After incubation, dentates were allowed to settle,
media was removed, and one ml of new Neurobasal was added. Dentates were triturated
gently and then allowed to settle. The top portion was removed and the dense remaining material was triturated and recovered two more times. Recovered material was
centrifuged at 1000 x g for ten minutes. The pellet was resuspended in neurobasal + and
seeded at a concentration of about 50-100 cells/mm² onto polylysine/laminin coated coverslips. Neurons were incubated for a few hours to allow attachment to the coverslip,
then fed one ml of supplemented Neurobasal. The following morning, the old media
replaced with two ml of new media. Neurons were transfected at 3 days in vitro (DIV)
and analyzed at 5 DIV.
Transfections
NIH/3T3 cells were transfected using Lipofectamine (Invitrogen, Carlsbad, CA)
according to the manufacturer’s protocol when cells were 50% confluent (24 hrs after
seeding), and assessed 24-48 hrs later. Dissociated neurons were transfected with the
following modifications to reduce toxicity: In a 2 ml volume of media, 1.6 μg DNA was
complexed with 4 μl Lipofectamine 2000.
Antibodies and flourescently labelled reagents
The following antibodies were used at the stated concentrations for
immunofluorescence (IF) or western blot (WB) as indicated: βIII-tubulin (IF 1:500)
(Sigma); α-tubulin (IF 1:250, Molecular Probes); M2 anti-FLAG (IF 1:150, WB 1:1000,
79 Sigma); JL8 anti-GFP (WB 1:1000, Clontech), anti-C1/ARHGAP4 (IF 1:250 for
dissociated neurons and NIH/3T3 cells, 1:1000 for slices; WB 1:1000) (Foletta et al.,
2002), L1 (IF 1:2000) (gift from Vance Lemmon, University of Miami). Secondary
antibodies included: biotinylated goat anti rabbit (1:500, Molecular Probes, Eugene, OR), biotinylated horse anti-mouse (1:1000, Vector, Burlingame, CA), Texas red conjugated goat anti-mouse (1:500, Molecular Probes) and oregon green conjugated goat anti-mouse
(1:500, Molecular Probes). Biotin labelled secondary antibodies were visualized by using strepavidin conjugated flourophores (594 and 488, 1:1000, Molecular Probes).
Western blots were subjected to secondary antibodies: horseresdish peroxidase (HRP) conjugated rabbit anti-mouse (1:10000, Sigma), and HRP conjugated goat anti-rabbit
(1:10000), and detected by ECL. F-actin was visualized by incubating cells with
flourescently conjugated phalloidin (Molecular Probes) according to manufacturer’s
instructions.
Endogenous ARHGAP4 detection in hippocampal slices
150 μm slices were made from p21 Sprague-Dawley rats and the hippocampi were removed. The hippocampi were fixed with 4% paraformaldehyde and stained for endogenous ARHGAP4 using the rabbit ARHGAP4 antibody (Foletta et al., 2002).
Wound assays, drug treatments
NIH/3T3 cells were seeded on glass coverslips and transfected 24 hours later
(when cells were about 50% confluent). Cells were then grown to confluency in the presence of serum and wounded with an 18 guage needle. For drug studies, cells were
80 allowed to migrate for 3 hours in the presence of serum before applying either DMSO,
300 nM Nocodazole in DMSO or 1 μM Cytochalasin-D in DMSO for 5 minutes. Cells were then fixed in 4% paraformaldehyde for 15 minutes at room temperature for analysis.
MT cosedimentation assay
Co-sedimentation of ARHGAP4 proteins and MTs was performed using the
Microtubule Binding Protein Spin Down Assay Kit #BK029 (Cytoskeleton, Denver, CO).
Briefly, cells expressing FLAG or EYFP -tagged ARHGAP4 fusion proteins were lysed with general tubulin buffer (Cytoskeleton). Lysates were centrifuged at 100,000 x g for one hour at 4oC in a Beckman XL-100 Ultracentrifuge. Supernatant fractions were incubated for 30 min at RT with purified, taxol-stabilized microtubules. Microtubules were pelleted by ultracentrifugation (100,000 x g for 60 min at room temperature) and the supernatant and pellet fractions were subjected to SDS-PAGE. Resolved proteins were transferred to a membrane and analyzed by Western analysis using the M2 anti-FLAG antibody (Sigma) or the JL-8 anti-GFP antibody (BD Biosciences/Clontech).
Microtubule associated protein-2 (MAP-2) and bovine serum albumin (BSA) (provided with kit) were used as controls to test the specificity of the assay.
Statisical analysis
All statistical analyses were done with SigmaStat and graphs were generated in
SigmaPlot. Statistical significance was determined using one way analysis of variance
(ANOVA).
81 Results
ARHGAP4 localizes to NIH/3T3 cell leading edges via the FCH domain
In previous work, ARHGAP4 was found to localize to the perinuclear MT
network and golgi in rat NRK cells as well as to the tips of neurites in PC12 cells that
were stimulated with NGF (Foletta et al., 2002). One obvious question concerned the
role of ARHGAP4 in cells, and how its distribution to different subcellular regions
influences its function. To understand how ARHGAP4 localizes to different subcellular
areas, a structure-function approach was taken to assess what domains of ARHGAP4
were necessary for both localization and function. Deletion mutants designed to truncate
specific regions and a point mutant to abolish GAP activity were generated (figure 2.1).
The 72-965 mutant lacks the most conserved motif (RAEYL) in the FCH domain, while
the 1-71 peptide contains it. The 1-770 mutant is a carboxy-terminal deletion that results
in a loss of the SH3 domain and peripheral regions. The arginine finger in ARHGAP4 was mutated to an alanine to abolish GAP enzymatic activity for the R562A mutant.
While the wildtype protein enhances GTP hydrolysis over basal rates, the GAP domain of the R562A mutant does not, indicating this mutant lacks GAP activity (figure 2.2).
Unlike the GAP domain of human ARHGAP4, which is selective for RhoA (Christerson et al., 2002), the GAP domain of rat ARHGAP4 is not selective between RhoA, Rac1 and
Cdc42 in vitro. In addition, neither the FL or R562A GAP domains significantly enhanced the GTP hydrolysis on Ras.
To understand if ARHGAP4 localized in a similar manner in NIH/3T3 cells as it did in NRK and PC12 cells, we stained NIH/3T3 cells using an antibody that was previously used to characterize ARHGAP4 in this cell line (Foletta et al., 2002). Staining
82 of endogenous ARHGAP4 revealed that the protein had a strong perinuclear distribution
(data not shown) and also localized to the leading edges of NIH/3T3 cells (figure 2.3),
similar to the distribution in PC12 cells when stimulated with NGF. Cells were costained
for alpha-tubulin to determine if ARHGAP4 expression overlapped with the MT network
as described previously (Foletta et al., 2002). The distribution of perinuclear ARHGAP4 overlapped with the MT network, but not in a similar pattern, and at the leading edge of cells ARHGAP4 was found to line up with the protruding membrane in a pattern that resembles actin distribution but not the linear array found for MTs (figure 2.3).
The localization at the tips of protrusive cytoplasmic extensions placed
ARHGAP4 in a region of the cell that undergoes high cytoskeletal turnover and where
GTPase regulation occurs to control cell motility. To determine how ARHGAP4 was targeted to this location, full length ARHGAP4 and constructs with point or deletion mutants were transfected into NIH/3T3 cells and the proteins were analyzed for their ability to localize to the tips of protrusions. Full length ARHGAP4 (1-965), R562A, 1-
770, and 72-965 were used to determine if the FCH, GAP or SH3 domains could target
ARHGAP4 to leading edges. To assess targeting to the tips of cytoplasmic extensions, fluorescence intensity (using both FLAG staining and native EYFP) was first measured at the tip of each process and then measured 20 μm from the tip, with the two readings expressed as a ratio of fluorescence intensity (see methods) (figure 2.4). These data demonstrated that the 72-965 mutant did not target to the tips of extending processes, indicating that amino acids 1-71 are necessary for this localization. The GAP mutant
(R562A) and the carboxy-terminal truncation mutant (1-770) targeted in a similar manner as the full length protein, indicating that the GAP and SH3 domains were not necessary
83 for localization. To determine if amino acids 1-71 were sufficient for targeting, a 1-71
construct was transfected into NIH/3T3 cells and assessed in a similar manner. The 1-71-
EYFP protein targeted efficiently to the tips of processes. This targeting to the tips of
extending processes in NIH/3T3 cells suggests that the function of ARHGAP4 may be spatially restricted at the leading edge.
The FCH domain targets ARHGAP4 to growth cones
ARHGAP4 transcripts were previously found to be expressed in specific regions of the postnatal central nervous system, including the hippocampus (Foletta et al., 2002).
Expression of ARHGAP4 protein increases from postnatal day 7 to 21 and parallels the postnatal maturation of the dentate gyrus and the axogenesis that follows (figure 2.5 B).
By p21 there was an enrichment of ARHGAP4 protein in the stratum lucidum area of the hippocampus (figure 2.5 A, A”), but only faint staining in the molecular and hilus regions
(figure 2.5 A’). This area is where the mossy fiber axons grow through as they synapse onto the CA3 pyramidal cells, and suggests that ARHGAP4 protein is localized in mossy fiber axons and potentially growth cones as well. To get a better understanding of
ARHGAP4 localization in dentate granule neurons, dissociated neurons were cultured for
5 days on poly-l-lysine/laminin and then stained for ARHGAP4. ARHGAP4 protein could be seen perinuclear and weakly throughout some dendrites and the axons (data not shown), and was highly enriched in growth cones (figure 2.6). The expression in the growth cone extended from the central domain into the peripheral domain, where it was found to be present at the tips of filopodia (arrows, figure 2.6) and along the leading
84 edges of the interspersed lamellipodia. This enrichment of ARHGAP4 in growth cones is
similar to the targeting of ARHGAP4 to the tips of extending processes in NIH/3T3 cells.
To determine if ARHGAP4 targeted to growth cones in a similar manner as in
NIH/3T3 cells, ARHGAP4 expression vectors were transfected in dissociated neurons at
3 DIV and assessed for localization at 5 DIV. By this stage, many neurons are polarized with committed axons and dendrites, yet still maintain a highly motile growth cone. Full length ARHGAP4 (1-965), R562A, 1-770, 72-965 and 1-71 were assessed and compared to the distribution of the cytoplasmic EYFP protein (figure 2.7). The 1-965, R562A, 1-
770 and 1-71 proteins localized to growth cones and their fluorescence intensity ratios were not significantly different from each other. However, their ratios were significantly different from that of EYFP, which had a diffuse pattern within the neuron (figure 2.8).
The 72-965 protein did not localize well to growth cones but could be found in axons
(figure 2.7 J-L). There was no significant difference between EYFP and 72-965 fluorescence intensity ratios, indicating that the 72-965 mutant is mostly diffuse in neurons. Although the level of EYFP protein was much higher than 72-965, the growth cone to axon pixel intensity was similar for both, arguing for a diffuse distribution of
each protein. These data indicate that amino acids 1-71 are both necessary and sufficient
for targeting ARHGAP4 to growth cones and also demonstrate that ARHGAP4 is
targeted to peripheral regions in a similar manner in both neurons and NIH/3T3 cells.
These parallels suggested that ARHGAP4 may be under the same type of spatial control
in both model systems, at the leading edges of NIH/3T3 cells and in growth cones.
85 The FCH domain indirectly associates with microtubules
To determine what role the FCH domain plays in ARHGAP4’s function and localization, we next investigated whether ARHGAP4 is associated with the MT network, either indirectly or directly. The study from Foletta et al. (2002) demonstrated that ARHGAP4 could colocalize with the perinuclear MT network. Also, the FCH domains of CIP4 and Rapostlin have been shown to interact with MTs (Tian et al., 2000;
Fujita et al., 2002). This suggested that the FCH domain of ARHGAP4 may function to bind MTs. To determine if ARHGAP4 could bind MTs, ARHGAP4 mutants were expressed in NIH/3T3 cells and the cytosolic fraction of the lysates were combined with polymerized tubulin using the Microtubule Binding Protein Spin Down Assay Kit
(Cytoskeleton). Proteins that can associate with polymerized tubulin will pellet at
100,000 x g, while those that do not will remain in the supernatant. MAP2 binds MTs and was found in the pellet fraction, while cytoplasmic BSA is enriched in the supernatant (figure 2.9). Both full length ARHGAP4 and R562A cosedimented with
MTs, but the 72-965 protein was only present in the supernatant fraction. This indicated that amino acids 1-71 were necessary for an interaction with tubulin. To assess this further, 1-71 was expressed in NIH/3T3 cells and analyzed in a similar manner. Amino acids 1-71 cosedimented with MTs, indicating that this protein was sufficient to associate with MTs. These data demonstrated that ARHGAP4’s FCH domain was capable of associating with MTs, but since these data were performed with overexpressed proteins from cell lysates it was still unknown if this was a direct or indirect interaction.
To determine if the association with MTs was direct or indirect, recombinant proteins were made to use in the MT cosedimentation assay. Making a full length GST-
86 tagged ARHGAP4 recombinant protein was not a likely possibility due to the size
(predicted 146 kDa), so amino acids 1-71 and 1-289 were used instead. While the 1-71 protein contains the minimal fragment required to associate with MTs, the 1-289 protein would include the extended FCH domain (Tsujita et al., 2006) and conserved ARNEY motif (Coyle et al., 2004) (figure 1.6), which may function in tandem with the FCH domain. When these recombinant proteins were tested for their ability to bind directly to
MTs, neither the GST-1-71 or GST-1-289 proteins cosedimented with MTs. This indicated that although the FCH domain of ARHGAP4 could associate with tubulin indirectly, this did not appear to be due to direct binding.
ARHGAP4 is tethered to the leading edge actin cytoskeleton
Recombinant proteins purified from bacteria are not posttranslationally modified, yet these modifications may be important for ARHGAP4 function. For instance,
ARHGAP4 may associate with MTs in a state dependent manner at its site of action, which cannot be assayed in a MT cosedimentation assay with recombinant proteins.
Another hypothesis is that the ARHGAP4/MT association is only important for transport, and that this association is not necessary once ARHGAP4 reaches its destination. To test
If ARHGAP4’s interaction with MTs was occurring at the leading edge, cells expressing
ARHGAP4 mutants were treated with either MT or actin destabilizing drugs. If
ARHGAP4 was associated with either cytoskeletal element, than mislocalization should occur in the presence of nocodazole or cytochalasin-D.
Full length ARHGAP4 and the 1-770 and 72-965 mutants were assessed to see if they mislocalized when MTs were disrupted with 300 nM nocodazole, or when actin was
87 disrupted with 1 μM cytochalasin-D. Cells were transfected with the ARHGAP4 expression vectors and wounded to induce polarization and migration. In migrating cells,
actin and MTs are extremely dynamic at the leading edges, making this region very
sensitive to drugs. After three hours of migration, cells were treated with nocodazole or cytochalasin-D for five minutes and then stained to determine the localization of the
ARHGAP4 proteins. The peripheral MT and actin cytoskeletons were greatly disrupted
after 5 minutes in drugs (figure 2.10). The full length ARHGAP4 is localized at the
leading edges of migrating cells and disruption of the MT network did not alter its
distribution (figure 2.11 D-F). Interestingly, disruption of the actin cytoskeleton causes
ARHGAP4 to assume a more diffuse distribution, and it is no longer found at the leading
edge (figure 2.11 G-I). The 1-770 mutant, which lacks the SH3 domain and peripheral
regions is also found at leading edges (figure 2.12 A-C), and addition of nocodazole does
not alter its localization at the leading edge (figure 2.12 D-F). The 1-770 mutant was a
little more diffuse in both the DMSO and nocodazole than full length, but was still
localized at the leading edge. Similar to full length protein, the 1-770 protein also
mislocalized when actin filaments were disrupted (figure 2.12 G-I). The 72-965 mutant
did not target to leading edges and disrupting the MT or actin networks had no effect on
localization (figure 2.13), supporting the idea that this protein is diffuse. These data
suggest that once ARHGAP4 is localized to leading edges, neither MT binding nor
association are required for ARHGAP4 to remain, lending support to the targeting
hypothesis. However, it also demonstrates that ARHGAP4’s localization at the leading
edge may require some sort of tethering to the leading edge actin network. Both full
length and 1-770 proteins mislocalized when actin was disrupted but not when MTs were
88 disrupted, suggesting the domain that tethers ARHGAP4 to this region may lie with amino acids 1-770, which does not include the SH3 domain.
Discussion
ARHGAP4 is a multidomain Rho-family GAP protein with FCH, GAP and SH3 domains. This study examined the structure-function of ARHGAP4 with respect to subcellular localization and cytoskeletal interactions. Previous work had demonstrated gross expression patterns of ARHGAP4 in the developing central nervous system and noted multiple subcellular localization patterns in rat NRK cells and differentiated PC12 cells (Foletta et al., 2002). This work extended the focus to understand how the particular domains of ARHGAP4 are related to its subcellular targeting and function. By expressing deletion and point mutants in both fibroblasts and primary neurons, a targeting role was discovered for the amino terminal 71 amino acids which contain the majority of the FCH domain and its conserved RAEYL motif. This region was found to be both necessary and sufficient for targeting ARHGAP4 to NIH/3T3 cell leading edges and to dentate granule cell growth cones (figures 2.4, 2.7, 2.8). Interestingly, neither a functional GAP domain nor the SH3 or peripheral domains on the carboxy-terminus are necessary for this targeting (figures 2.4, 2.7, 2.8).
Since ARHGAP4 was found colocalized with perinuclear MTs in rat NRK cells
(Foletta et al., 2002), and other FCH domains have been implicated in binding to MTs
(Tian et al., 2000; Fujita et al., 2002), it seemed likely that ARHGAP4’s targeting may be the result of trafficking along MTs. The same amino acids that were shown to target
ARHGAP4 to leading edges and growth cones were found to be both necessary and
89 sufficient for MT association but not direct MT binding (figure 2.9). These results indicate that unlike CIP4 (Tian et al., 2000), ARHGAP4 may not bind directly to MTs.
The initial MT binding experiments (figure 2.9 A) utilized the cell lysates of overexpressed proteins and combined those lysates with purified tubulin. These cell lysates contain several cytosolic proteins that could link ARHGAP4 to MTs, including motor proteins, small trafficking vesicles or scaffolding proteins.
When recombinant proteins were tested for their ability to bind MTs, neither the
GST-1-71 or the larger GST-1-289 proteins cosedimented with MTs (figure 2.9 B). This indicated that the domain responsible for linking ARHGAP4 to the MT cytoskeleton was not doing so directly. When taken together with the targeting data, it supports a role for
MT based transport of ARHGAP4 to leading edges and growth cones. This idea raises the new question of what is ARHGAP4’s FCH domain binding to in order to target to these locales? Although these data can not answer that question, there have been a plethora of papers demonstrating that the FCH domain may also bind to actin or lipids that can target them to sites of action (Yeung et al., 1998; Lippincott and Li, 2000; Icking et al., 2006; Tsujita et al., 2006; Wachtler et al., 2006). Similar to the targeting of
GTPases to membranes upon activation and binding to the phosphoinositides (Heo et al.,
2006), FCH domain containing proteins may also be targeted to areas where GTPases are activated by phosphoinositides at the membrane. Many of these FCH domain containing proteins are able to bind phosphatidylserine (PS) and phosphatidylinositol 4,5- bisphosphate (PI[4,5]P2), and be recruited to membranes through conserved basic residues in the FCH and extended FCH domains (Tsujita et al., 2006). In addition, brightfield images of NIH/3T3 cells have many membrane ruffles and membrane
90 foldings at the leading edge, and ARHGAP4 proteins that contain the FCH and eFCH
domains are highly localized in these subregions (figures 2.11 C, 2.12 C). These
membrane foldings and ruffles are areas of high actin and membrane dynamics, and
many GTPase signaling events downstream of EGF and PDGF activation are thought to
take place here (Maddala et al., 2003). EGF and PDGF are well known inducers of
leading edge membrane ruffling and activators of Rac1. In addition, treatment of
NIH/3T3 cells with cytochalasin-D to disrupt the peripheral actin cytoskeleton
mislocalized FL ARHGAP4, as well as the 1-770 mutant from the leading edge. This
indicates that ARHGAP4 is tethered to the leading edge actin cytoskeleton in some
manner and that the domain responsible is between amino acids 1-770. While the 1-770
mutant lacks the SH3 domain, ARHGAP4 contains other domains that could mediate
protein-protein interactions that link it to the actin cytoskeleton, including the FCH
domain and predicted coiled-coil regions within the eFCH domain.
It it also possible that ARHGAP4’s localization at leading edges could be regulated by extracellular signals. The FCH domain containing protein CIP4/2 is known to translocate to the plasma membrane downstream of insulin signaling, and this event requires a functional FCH domain (Chang et al., 2002). How this signal regulates FCH domain containing proteins is still unknown, but some work has demonstrated that FCH domain proteins are tyrosine phosphorylated in response to serum stimulation, suggesting that phosphorylation may regulate FCH domain function and affinity for its targets
(Yeung et al., 1998). In support of this hypothesis, the tyrosine residue in the conserved
RAEYL motif (figure 1.6) is highly conserved among FCH domains and is predicted to be a tyrosine phosphorylation motif (netphos 2.0). The FCH domain may also be part of
91 a larger functional domain. Several FCH domains are juxtaposed to an extended FCH
(eFCH) domain that is loosely homologous to the lipid-sensing BAR domain (Tsujita et al., 2006). The eFCH domain is defined as a helical region with interspersed basic amino acids that may be exposed to sense acidic residues or positively charged moieties.
Mutation of these basic amino acids disrupted the ability of these proteins’ FCH + eFCH domains to bind and curve specific lipids in vitro, suggesting that these residues are functioning in a similar manner to the BAR domain. Considering that ARHGAP4 did not bind to MTs directly but it was still able to associate in cell lysates to MTs, there is the possibility that ARHGAP4 may bind to lipid moieties and be recruited to sites of action.
This interaction may sequester ARHGAP4 at the leading edges of cells where high levels of specific lipids are required for membrane protrusion and polarity, and where active
GTPases are recruited to mediate signaling (Heo et al., 2006).
The other domains of ARHGAP4 did not influence targeting to the leading edges of NIH/3T3 cells or growth cones, suggesting some other function. ARHGAP4’s GAP domain has been examined before and was found to act as a GAP towards RhoA, Rac1 and Cdc42 in vitro (Foletta et al., 2002). In agreement with those findings, ARHGAP4 also enhanced GTP hydrolysis on RhoA, Rac1 and Cdc42 in this study, but not the Ras
GTPase (figure 2.2). While these experiments demonstrate that the GAP domain has enzymatic activity, it does not address the in vivo selectivity for a particular GTPase, or lack of selectivity. These experiments show that ARHGAP4 is localized at the leading edges of NIH/3T3 cells and growth cones (figures 2.4, 2.7, 2.8), and this localization may dictate what substrates are available for ARHGAP4 to target as cells migrate and growth cones navigate their environments. Another group has reported that human ARHGAP4
92 selectively targets RhoA in vitro (Christerson et al., 2002), suggesting that ARHGAP4 orthologues may function differently or that in vitro assays may not be a good standard to determine substrate selectivity. Future experimnts should address what GTPases are candidate targets in cells and can be substrates for full length ARHGAP4, and not just its
GAP domain.
The SH3 domain of ARHGAP4 did not target ARHGAP4 to leading edges or growth cones either. The SH3 domain of ARHGAP4 has been shown to have different functions in separate cell lines, but was able to associate with important scaffolding or signaling proteins in each type. One report demonstrated that ARHGAP4’s SH3 domain directly interacts with MEKK1 (Christerson et al., 2002), a protein which is found at focal contacts and stress fibers through its association with alpha actinin (Christerson et al., 1999). This interaction was found to be important for AP-1 transcriptional regulation.
In hematopoietic HL60 cells, ARHGAP4’s SH3 domain associated with HEM-1, which is a protein known to be part of the inhibitory WAVE complex in hematopoietic cells, and multiple cytoskeleton and polarity proteins, including another member of the complex that inhibits WAVE (Cyfip1/2), a class III PI3 kinase (Vps34), and Diaphanous
(Weiner et al., 2006). While no function was examined for ARHGAP4 in this system, it was suggested that ARHGAP4 may be part of a leading edge polarity complex with
HEM-1 that could regulate leading edge actin in migrating neutrophils. These studies suggest that ARHGAP4’s SH3 domain may mediate indirect binding to actin through any number of signaling or scaffolding intermediates. With this in mind, it was expected that the ARHGAP4 mutant 1-770 (which lacks the SH3 domain) would mislocalize from leading edges when the tubulin cytoskeleton was disrupted. However, the 1-770 protein
93 was still found at leading edges in the presence of nocodazole, but was severely mislocalized when actin filaments were disrupted with cytochalasin-D (figure 2.12), suggesting that ARHGAP4 is tethered in some way to the actin cytoskeleton via a region in amino acids 1-770, which lack the SH3 domain. However, there was a significant disruption in localization of the 1-770 mutant in perinuclear regions when nocodazole was applied (figure 2.12 D). These results suggest that there is some interaction with
ARHGAP4 and the perinuclear MT network via the SH3 domain and/or other regions in the carboxy-terminus.
With the various protein-protein interactions reported for ARHGAP4’s SH3 domain, and another report that it interacts with dendrin (Foletta et al., 2002) and Appbp1
(Scott Young, personal communication), it seems likely that ARHGAP4 can integrate several signaling components in different cell types via its SH3 domain. Some of these interactions have functional consequences, but the pathways involved are not known to be related or working together. In addition, some of the proteins that bind to
ARHGAP4’s SH3 domain are not expressed in every cell type that ARHGAP4 is expressed in. For example, HEM-1 is a hematopoietic specific protein, but not expressed in the brain or NIH/3T3 cells. However, the related protein HEM-2 (also known as Nck associated protein-1, NAP-1) is expressed in neurons, and is necessary for localized Rac1 induction of lamellipodia (Steffen et al., 2004) and axon guidnace (Hummel et al., 2000).
Even though there is a great deal of literature supporting several binding partners for
ARHGAP4’s SH3 domain, deletion of the carboxy-terminus has no effect on targeting to leading edges or growth cones. This suggests that protein-protein binding mediated by the SH3 domain is secondary to localization of ARHGAP4.
94 One hypothesis to explain localization is that ARHGAP4 associates with protein
complexes bound for leading edges and growth cones via the SH3 domain and/or
carboxy-terminal domains, but this hypothesis is complicated by two factors. First, the
72-965 protein has a diffuse distribution in cells and neurons, suggesting that even if the
SH3 domain and carboxy tail are trafficked on MTs, they are not arriving at the same destinations as the full length protein or its 1-71 fragment. Second, amino acids 1-71 are sufficient to target to leading edges and growth cones, indicating that these amino acids may be the sole way for ARHGAP4 to target. It is posible that amino acids 1-71 are simply sequestered by leading edges and growth cone targets and over time they build up in these regions. However, with the large distance from neuronal somas to growth cones, it seems unlikely that diffusion first, followed by sequestration, is the case. In some rare cases, the 1-71 protein seems to be tracking along MTs (figure 2.7 M-O), suggesting that
this domain may be targeting ARHGAP4 to sites of action in cells. Another interesting
possibility is that ARHGAP4 is differentially activated and capable of binding to certain
substrates at leading edges and growth cones as opposed to more cytosolic domains in
perinuclear locales. In this scenario, ARHGAP4 may traffic along MTs via amino acids
1-71 until it reaches leading edges and growth cones, and be recruited to signaling
networks along the way via its SH3 domain. In this model, several ARHGAP4 proteins
would traffic along MTs and occasionally hop off to locally regulate signaling events (for
example, at focal adhesions) by binding to activated signaling proteins. Those proteins
that make it to the leading edge may incorporate into the leading edge or be recruited in a similar manner when needed. If the MT cytoskeleton was disrupted, than proteins that were only able to travel to the leading edges and growth cones and not hop off on the way
95 would be mislocalized, while those that made it to the leading edge and were recruited by different complexes would be properly localized.
96
97 Figure 2.1: ARHGAP4 constructs used in transfections and bacterial protein expression.
A) Full length ARHGAP4, FCH and SH3 deletion mutants, and a point mutation in the arginine finger of the GAP domain were generated to express as FLAG or EYFP fusion proteins in mammalian cell lines. B) The GAP domain and peripheral regions (amino acids 474-743) of wildtype (WT) and R562A ARHGAP4 were generated in frame to
GST to express and purify from bacteria.
98
99 Figure 2.2: In vitro GAP assay of WT and R562A ARHGAP4 GAP domains.
The enzymatic GAP activity of ARHGAP4 and the R562A mutant were assessed
using purified GST-fusion proteins encoding ARHGAP4’s GAP domain and peripheral
regions (amino acids 474-743). GTP-loaded Ras-His, RhoA-His, Rac1-His, or Cdc42-
His proteins were combined with either GST-WT-GAP or GST-R562A-GAP proteins
and incubated together for 20 minutes. To determine levels of hydrolyzed GTP, cytophos
buffer was added to the reactions and the levels of cleaved, inorganic phosphate were
read by a spectrophotometer at 650 nm. Levels of GTP hydrolysis in the presence of either GST-WT-GAP or GST-R562A-GAP + GTPase were normalized to levels with
GTPase alone and are presented as the fold induction of GTPase hydrolysis. GST-WT-
GAP enhanced the hydrolysis of GTP on RhoA, Rac1 and Cdc42 but not Ras, while the
GST-R562A-GAP protein did not show significant GAP activity with any of the
GTPases. The dashed line represents relative Ras, RhoA, Rac1 and Cdc42 GTP hydrolysis levels. Each condition was performed in triplicate 2-3 times, and data are represented as the mean ± SEM, (* p<0.05, **p=0.006).
100
101 Figure 2.3: Endogenous ARHGAP4 localizes to the leading edge of NIH/3T3 cells.
NIH/3T3 cells were seeded on glass coverslips and grown for 2 days in the presence of serum before fixing and staining for alpha tubulin and ARHGAP4. Alpha
tubulin was detected with a mouse monoclonal antibody and subsequent staining with a
Texas red conjugated goat anti-mouse secondary. ARHGAP4 was detected using a rabbit
polyclonal antibody, followed by a biotinylated goat anti-mouse secondary and
subsequent staining with streptavidin conjugated to Alexa 488. Immunofluorescence
revealed a leading edge distribution of ARHGAP4 that extends beyond the tubulin
cytoskeleton. Confocal images were taken with a 63x oil objective with a 4x digital
zoom, scale bar 5 μm. Arrow points to leading edge of cell.
102
103 Figure 2.4: Amino acids 1-71 are necessary and sufficient to target ARHGAP4 to the tips
of NIH/3T3 cell cytoplasmic extensions.
NIH/3T3 cells were transfected with an EYFP vector, FL (1-965) (A), R562A
(B), 1-770 (C), 72-965 (D), or 1-71 (E) constructs and fixed after 48 hours. Proteins were visualized by FLAG staining, followed by staining with biotinylated horse anti-mouse
secondary and subsequent visualization with Streptavidin conjugated Avidin alexa 488,
or by directly observing native EYFP. Proteins with amino acids 1-71 were found at
leading edges (arrows). F) Western analysis of NIH/3T3 cells tranfected with
ARHGAP4 constructs demonstrating that proteins are detectable at the right size, (1-71-
EYFP western analysis is shown in figure 4.1). Blot was stained for FLAG using the M2 antibody, followed by secondary staining with an HRP-conjugated rabbit anti-mouse antibody and detection by ECL. G) Pixel intensity of either FLAG or EYFP proteins were measured using ImageJ software at the tips of extending cytoplasmic extensions and
20 μm away from the tip of cytoplasmic extension. 1-965, R562A, 1-770 and 1-71 proteins were enriched at the tips of cytoplasmic extensions and were significantly different than EYFP alone, indicating a non-diffuse distribuion in cells. The 72-965 protein was not significantly different from EYFP, suggesting a more diffuse distribution.
Data are from 3 independent experiments and are presented as ± SEM, *p<0.05.
104
105 Figure 2.5: ARHGAP4 is enriched in the mossy fiber axons of the hippocampus.
A) 150 um slices from p21 rats were fixed and stained for endogenous
ARHGAP4 protein using a rabbit polyclonal ARHGAP4 antibody, and subsequently detected with a biotinylated goat anti-rabbit secondary and incubation with streptavidin- conjugated Alexa 594. Staining revealed a restricted expression within the stratum lucidum layer of the hippocampus A”, with a little fluorescence seen in the hilus and molecular layers of the dentate gyrus A’. (abbreviations: ML molecular layer, DG dentate granule cell layer, H hilus, SL stratum lucidum, CA3 CA3 pyramidal cell layer).
B) Hippocampal tissue was harvested from p7, p14, and p21 rats and 20 μg total protein added to each lane. SDS-PAGE gels were run and blots were probed for ARHGAP4, followed by an HRP-conjugated goat anti-rabbit antibody and ECL detection.
ARHGAP4 immunoreactive bands migrated at 120 (predicted full length), 80, 50, 42 and
36 kDa. Blot is representative of at least three independent experiments.
106
107 Figure 2.6: Endogenous ARHGAP4 is enriched in mossy fiber growth cones.
Dissociated dentate granule cells were seeded on polylysine/laminin coated coverslips and grown for 5 days before fixing and staining for beta III tubulin and
ARHGAP4. Microtubules were visualized with a βIII tubulin antibody, followed by an
Oregon Green conjugated goat anti-mouse secondary, and ARHGAP4 was visualized with the rabbit polyclonal antibody, followed by detection with a biotinylated goat anti- rabbit secondary and incubation with streptavidin conjugated Alexa 594. The red and green images were changed to green and red, respectively, in photoshop.
Immunofluorescence revealed distribution of ARHGAP4 in both the central (C) and peripheral (P) domains of growth cones. Confocal images were taken with a 63x oil objective with a 4x digital zoom. Arrows point to filopodial extensions. Scale bar = 5
μm.
108
109 Figure 2.7: ARHGAP4 expressed proteins localize to growth cones via amino acids 1-
71.
Dissociated dentate granule cells from p5-7 rats were seeded on coverslips coated
with poly-L-lysine /laminin and grown for three days before transfecting with ARHGAP4
full length and mutant expression vectors. Transfected cultures were grown for an additional two days before fixing and then incubated with L1 or βIII-tubulin antibodies to
identify neurons, and transfected neurons were visualized by staining for FLAG or native
EYFP. M2 staining was visualized using an Oregon Green conjugated goat anti-mouse
secondary, L1 staining was detected with a biotinylated goat anti-rabbit secondary
followed by incubation with Streptavidin conjugated Alexa 594, and beta III tubulin was
visualized using a biotinylated horse anti-mouse secondary followed by incubation with
Streptavidin conjugated Alexa 594. The FL (1-965)-FLAG (A-C), R562A-FLAG (D-F),
1-770-FLAG (G-I) and 1-71-EYFP (M-O) proteins localized to growth cones while the
72-965-FLAG (J-L) protein was seen in the axon shaft but not enriched in the growth
cone. Arrows point to localization of proteins in filopodia, while asterick (M-O) shows a
filopodial extension that contains 1-71-EYFP protein but lacks microtubules. All images
were taken with a 100x objective with a 4x digital zoom. Scale bar = 5 μm.
110
111 Figure 2.8: Amino acids 1-71 are necessary and sufficient to target ARHGAP4 to growth
cones.
Dissociated dentate granule cells from p5-7 rats were cultured on
polylysine/laminin for 3 DIV, transfected with EYFP or ARHGAP4 mutants, and then cultured for an additional 2 days. A) Pixel intensities of FLAG stained or native EYFP neurons were recorded in the growth cone and 40 μm down the axon to determine the
ratio of growth cone to axon protein enrichment. B) Neurons EYFP alone were used as a measurement of a diffuse protein and were compared to other proteins. The FL (1-965),
R562A, 1-770 and 1-71 ratios were significantly growth cone enriched compared to
EYFP alone. The 72-965 protein had a similar ratio to EYFP and was not significantly different, suggesting a more diffuse distribution in axons and growth cones. Data are representative at least 3 independent experiments and are presented as ± SEM,
(* p<0.05).
112
113 Figure 2.9: Amino acids 1-71 can associate indirectly but do not bind microtubules in an in vitro microtubule cosedimentation assay.
A) NIH/3T3 cells were transfected with either FL (1-965), R562A, 72-965 or 1-
71 and grown for 48 hours. Cells were lysed and centrifuged at 100,000 x g for one hour to clear macromolecular complexes, polymerized actin and polymerized tubulin. Lysates containing overexpressed proteins were then combined with polymerized microtubules and allowed to complex for 30 minutes at RT before centrifuging the mixture through a
50% glycerol buffer at 100,000 x g for one hour. Supernatant (containing proteins that did not bind to polymerized microtubules) and pellets (polymerized microtubules and associated proteins) were collected and subjected to SDS-PAGE and blotting for FLAG or EYFP, followed by staining with a HRP-conjugated rabbit anti-mouse secondary antibody and subsequent detection with ECL. 1-965, R562A and 1-71 proteins were able to associate with the polymerized microtubules, but the 72-965 protein was only found in the supernatant. B) Recombinant proteins (1-71 and 1-289) as well as BSA and MAP2 were used to determine direct binding to microtubules. Purified proteins were combined with purified microtubules and complexed at RT for 30 minutes, then centrifuged for
100,000xg for 1 hour. Supernatant and pellet fractions were run on SDS-PAGE gels and proteins were visualized by coommassie. The cytosolic protein BSA was enriched in the supernatant fraction, while the microtubule associated protein MAP2 was enriched in the pellet. Neither the 1-71 nor the 1-289 proteins pelleted with microtubules. All proteins were verified to not pellet in the absence of microtubules (- MT lanes). Data represent at least three independent experiments per condition.
114
115 Figure 2.10: Leading edge distribution of microtubules and F-actin in the presence of nocodazole or cytochalasin-D.
Wound assay experiments were performed on NIH/3T3 cells, and three hours after wounding, the cells were treated for 5 minutes with vehicle alone (DMSO), 300 nM nocodazole, or 1 μM cytochalasin-D to disrupt MTs and actin filaments, respectively.
Wounded cells have leading edge microtubules (A-C) and F-actin (G-I) in the presence of
DMSO. Microtubules were visualized by staining for alpha tubulin and subsequent detection with a biotinylated horse anti-mouse antibody and subsequent incubation with
Streptavidin conjugated Alexa 594. F-actin was detected by incubating in Alexa 594- conjugated phalloidin. 300 nM nocodazole for five minutes is sufficient to disrupt the leading edge microtubules (D-F), while 1 μM cytochalasin-D for 5 minutes is sufficient to disrupt leading edge actin filaments (J-L). Images in A, D, G and J were taken with a
63x objective, while images in B, C, E, F, H, I, K and L were taken with a 63x objective and 4x digital zoom. Fluorescent images were merged with brightfield images (C, F, I,
L) to show leading edge boundaries. Scale bar = 5 μm.
116
117 Figure 2.11: Full length ARHGAP4 (1-965) leading edge distribution in the presence of nocodazole or cytochalasin-D.
Wound assay experiments were performed on NIH/3T3 cells that expressed full length FLAG-tagged ARHGAP4 (1-965-FLAG). Three hours after wounding, the cells were treated for five minutes with vehicle alone (DMSO), 300 nM nocodazole, or 1 μM cytochalasin-D to disrupt MTs and actin filaments, respectively. Transfected cells were detected by staining for flag, followed by incubation with an Oregon Green conjugated goat anti-mouse secondary antibody. Images in A, D, and G were taken with a 63x objective, while images in B, C, E, F, H, and I were taken with a 63x objective and 4x digital zoom. Fluorescent images were merged with brightfield images (C,F,I) to show leading edge boundaries. Scale bar = 5 μm.
118
119 Figure 2.12: 1-770 leading edge distribution in the presence of nocodazole or
cytochalasin-D.
Wound assay experiments were performed on NIH/3T3 cells that expressed the
carboxy-terminal truncation mutant (1-770-FLAG). Three hours after wounding, the
cells were treated for five minutes with vehicle alone (DMSO), 300 nM nocodazole, or 1
μM cytochalasin-D to disrupt MTs and actin filaments, respectively. Transfected cells
were detected by staining for FLAG, followed by incubation with an Oregon Green conjugated goat anti-mouse secondary antibody. Images in A, D, and G were taken with a 63x objective, while images in B, C, E, F, H, and I were taken with a 63x objective and
4x digital zoom. Fluorescent images were merged with brightfield images (C,F,I) to show leading edge boundaries. Scale bar = 5 μm.
120
121 Figure 2.13: 72-965 leading edge distribution in the presence of nocodazole or cytochalasin-D.
Wound assay experiments were performed on NIH/3T3 cells that expressed the amino-terminal truncation mutant (72-965-FLAG). Three hours after wounding, the cells were treated for five minutes with vehicle alone (DMSO), 300 nM nocodazole, or 1 μM cytochalasin-D to disrupt MTs and actin filaments, respectively. Transfected cells were detected by staining for flag, followed by incubation with an Oregon Green conjugated goat anti-mouse secondary antibody. Images in A, D, and G were taken with a 63x objective, while images in B, C, E, F, H, and I were taken with a 63x objective and 4x digital zoom. Fluorescent images were merged with brightfield images (C,F,I) to show leading edge boundaries. Scale bar = 5 μm.
122 Chapter 3
ARHGAP4 is an Inhibitor of NIH/3T3 Cell Motility and Dentate Granule Cell Axon Outgrowth
123 Summary
Wound healing and axon outgrowth are distinct events that share several signaling
parallels with respect to GTPase signaling. Both events require polarized cells that
translate guidance and growth cues into action. These events require strict control of the
Rho-family GTPases within discrete locales, notably the growth cones of axons and the
leading edges of migrating fibroblasts. Here it is demonstrated that the Rho-family GAP
ARHGAP4 inhibits NIH/3T3 cell migration and dentate granule cell axon outgrowth.
This inhibition requires specific domains on ARHGAP4 to mediate its effects. Using a targeting deficient ARHGAP4 mutant, a GAP mutant and a mutant that lacks a protein- protein interaction domain, it is shown that ARHGAP4 is a functional GAP that must be properly localized to inhibit cell migration and axon outgrowth.
Introduction
Axon outgrowth and cell migration are highly organized events that share many
similar mechanisms (Rodriguez et al., 2003). Both neurons and migrating fibroblasts
polarize, and have morphologically distinct structures that act in different ways in their immediate environment. Neurons send out neurites that commit to be dendrites or axons, and the growth cones of axons have distinct sets of receptors and signaling proteins that translate extracellular cues to mediate cytoskeletal elements (Huber et al., 2003). These signaling events primarilily utilize the Rho family of small GTPases to dynamically regulate actin and tubulin, and this regulation is under the tight control of spatially localized Rho family GEFs and GAPs (Bernards and Settleman, 2004). Although migrating fibroblasts do not respond to all the same extracellular cues as neuronal growth
124 cones, they do utilize Rho family GTPases, GEFs and GAPs to alter the actin and microtubule (MT) cytoskeletons in the same way (Rodriguez et al., 2003; Raftopoulou and Hall, 2004).
RhoGAP family members that are expressed in growth cones and fibroblasts are potent regulators of cytoskeletal dynamics, and this regulation is dependent on their localization and interaction with other signaling complexes. Often the same RhoGAP protein can be utilized in different ways within differing cell types, but can lead to the same result. For example, p190RhoGAP inhibits RhoA signaling in neurons and migrating cells, leading to similar effects on axon outgrowth and cell motility (Brouns et al., 2000; Haskell et al., 2001; Barberis et al., 2005). These experiments demonstrated that p190RhoGAP could be activated by semaphorin receptors in neurons and downstream of EGF and focal adhesions in fibroblasts to regulate RhoA in the same way.
Similarly, ARHGAP4 is another protein that may function similarly in different cell types. In the previous chapter, ARHGAP4 was shown to localize to the leading edges of fibroblasts and growth cones via the FCH domain (figures 2.4, 2.7, 2.8), and both of these cellular regions are similar in that they are polarized structures that have dynamic actin and MT cytoskeletons.
ARHGAP4 is expressed in dentate granule cells (figures 2.5, 2.6) and in NIH/3T3 fibroblasts (figure 2.3), and contains a functional GAP domain (figure 2.2; (Foletta et al.,
2002). Here, ARHGAP4 is shown to be an inhibitor of axon outgrowth and cell migration and mediates this function in similar ways. Through the combination of siRNA mediated knockdown and overexpression of ARHGAP4 and its mutants in
NIH/3T3 cells, an inhibitory role for ARHGAP4 has emerged. In addition,
125 overexpression of the R562A (GAP mutant) seems to act in a dominant-negative fashion to promote cell migration and axon outgrowth while deletions of either the FCH or SH3
domains abrogate ARHGAP4’s function. These experiments suggest that ARHGAP4
may mediate similar signaling events in neurons and migrating cells, and describe the
first functional role for ARHGAP4 as an inhibitory protein in cell migration and axon
outgrowth.
Materials and methods
Dentate explant cultures
Dentate gyrus explant cultures were prepared from postnatal day 5-7 Sprague-
Dawley rat pups as previously described (Butler et al., 2004). Rats were decapitated,
and 200μm horizontal hippocampal slices were prepared using a WPI Vibroslice. The
dentate gyrus was isolated from the hippocampal slices using a #15 scalpel. Each dentate
gyrus isolated from the hippocampal slices yielded 2 explants. After microdissection, the
dentates were trimmed to remove any remaining pyramidal cell and hilar regions.
Dentate explants were almost entirely composed of the granule cell layer and molecular
layer, with minimal hilus included. Dentate explants were placed on 25mm Nunc
Anopore membranes coated with polylysine (Sigma, St. Louis, MO) and laminin
(Invitrogen, Carlsbad, CA) and cultured in 6 well tissue culture plates containing 1.5ml
Neurobasal/B27 medium (Invitrogen).
126 Dissociated dentate granule cell cultures
The dentate gyrus was microdissected from 200 μm hippocampal slices prepared from postnatal day 5-7 rats as previously described (Butler et al., 2004). After microdissection, dentates were rinsed once in Neurobasal (supplemented with B27, glucose and L-glutamine) and then dissociated in Neurobasal/B27 with papain (2 ug/ml at 15-23 U/μg) for 30 minutes at RT. After incubation, dentates were allowed to settle, media was removed, and one ml of new Neurobasal was added. Dentates were triturated gently and then allowed to settle. The top portion was removed and the dense remaining material was triturated and recovered two more times. Recovered material was centrifuged at 1000 x g for ten minutes. The pellet was resuspended in neurobasal + and seeded at a concentration of about 50-100 cells/mm² onto polylysine/laminin coated coverslips. Neurons were incubated for a few hours to allow attachment to the coverslip, then fed one ml of supplemented Neurobasal. The following morning, the old media replaced with two ml of new media. Neurons were transfected at 3 DIV and analyzed at
5 DIV. Cultures were stained with M2 anti-FLAG antibody or visualized for native
EYFP to identify transfected neurons, and stained for either L1 or βIII tubulin to verify neuronal identify.
ARHGAP4 siRNA generation
Small interfering RNAs (siRNAs) were generated using the Silencer siRNA
Construction Kit (Ambion, Inc.). Nucleotides 203-223 of mouse ARHGAP4 mRNA were chosen as targets for silencing using the Ambion siRNA Target Finder program.
Briefly, complementary DNA oligonucleotides containing this region of ARHGAP4
127 cDNA and a complementary region of the 3’ end of the T7 promoter were annealed to a
T7 promoter primer and filled in with Klenow DNA polymerase to generate double- stranded templates. ARHGAP4 oligonucleotides: 5’-
ACAAGTTGGCTGAACGCTTTACCTGTCTC-3’, and 5’-
TAAAGCGTTCAGCCAACTTGTCCTGTCTC-3’. Control oligonucleotides: 5’-
ACAAATTAGCGGAGCGATTCACCTGTCTC-3’, and 5’-
TGAATCGCTCCGCTAATTTGTCCTGTCTC-3’ (6 underlined nucleotides encode silent mutations). The resulting double-stranded DNA templates were used to generate siRNA strands in in vitro transcription reactions using T7 RNA polymerase. ARHGAP4 or Control reaction products were combined to permit annealing of the siRNA strands, and DNA template removal, single-stranded RNA removal, and purification, were done according to the manufacturer’s protocol. siRNAs were transfected into NIH/3T3 cells using Lipofectamine 2000 at a final concentration of 2 nM. After transfection, cells were cultured an additional 2 days prior to wounding.
ExGen 500 transfection of dentate explants
20 μg DNA was added to 50μl of sterile 5% glucose and 3.6μl ExGen 500 in vivo transfection reagent (#R0521, Fermentas, Hanover, MD). The DNA was condensed for
10 minutes at room temperature with the ExGen 500. Then, 1µl of the transfection solution was applied to each explant using a micropipette. Explants were transfected 1 hour after culturing and then returned to the incubator for 48 hours before fixing.
128 Antibodies and reagents
βIII-tubulin (immunoflourescence 1:500, WB 1:1000, Sigma), M2 anti-FLAG (IF
1:150, Sigma), C1/ARHGAP4 (WB 1:1000) (Foletta et al., 2002), GFAP (IF 1:500, MP
Biomedicals), L1 (IF 1:2000) (gift from Vance Lemmon, University of Miami), and
Actin (C-2) (1:1000, SantaCruz Biotechnology). Secondary antibodies included: biotinylated goat anti rabbit (1:500, Molecular Probes), and oregon green conjugated goat anti-mouse (1:500, Molecular Probes). Biotin-conjugated secondary antibodies were visualized by using strepavidin conjugated Alexa 594 (at 1:1000, Molecular Probes).
Western blots were subjected to secondary antibodies: horseresdish peroxidase (HRP) conjugated rabbit anti-mouse (1:10000, Sigma), and HRP conjugated goat anti-rabbit
(1:10000), and detected by ECL or ECL+. DAPI (Molecular Probes) was used to stain
cell nuclei, (final dilution of 1:333 in PBS), and streptavidin conjugated HRP was used to
detect biotinylated antibodies for western analysis.
Dissociated axon outgrowth measurement criteria
Dissociated granule cells were seeded at 50-100 cells/mm² and cultured for 3 DIV
before transfecting with Lipofectamine 2000. Cultures were fixed at 5 DIV in 4%
paraformaldehyde for 30 minutes and stained. Axons were clearly distinguishable from
dendrites/immature neurites by 4-5 DIV, and were measured from the region where they
emanated from the soma to the tip of the growth cone. To be counted, the neuron could
not be touching any other cell or process. Only neurons with axons longer than 30 μm
were measured, otherwise they were considered immature or unhealthy.
129 Statistical analysis
Statistical analyses were performed with SigmaStat and graphs were generated
with SigmaPlot. Statistical significance was determined by one way analysis of variance
(ANOVA) and band densities were analyzed using Kodak 1D gel software.
Results
ARHGAP4 knockdown increases cell motility
The first description of ARHGAP4 linked its overexpression to the disassembly of actin stress fibers in SAA fibroblasts cells (Tribioli et al., 1996). Aside from a few reports suggesting that ARHGAP4 can associate with signaling proteins involved in mediating cytoskeletal function, the Tribioli et al., (1996) paper was the only description of ARHGAP4’s function in relation to the cytoskeleton. To further examine
ARHGAP4’s functional role, ARHGAP4 was knocked down using siRNA in the wound assay model system (figure 3.1). Within the wound assay, cells polarize and migrate into the wounded area and this process requires intense cytoskeletal rearrangements downstream of the GTPases (Rodriguez et al., 2003; Desai et al., 2004). If ARHGAP4
regulates cell motility or polarity, than depleting ARHGAP4 protein should lead to
different rates of migration in cells in a wound assay. The siRNA probes were tested for
their ability to knock down ARHGAP4 protein in NIH/3T3 cells. Western analysis
showed that the siARHGAP4 probe targeted against ARHGAP4 resulted in a 40% reduction in protein levels after 2 days in culture compared to a control siRNA probe
(siControl) (figure 3.2 A). To visualize cells that were transfected with siRNA, siRNA was cotransfected with an EYFP expression vector. Cells expressing EYFP had a
130 reduction in ARHGAP4 IF compared to their neighbors (data not shown). To verify that cells transfected with siControl were not unhealthy, their rate of migration was normalized and compared to EYFP transfected cells alone. The siControl+EYFP cells migrated the same as EYFP cells, but when the siARHGAP4+EYFP cells were normalized to the EYFP values, there was a significant increase in migration (figure 3.2
B). This indicated that a reduction in ARHGAP4 enhanced cell motility, suggesting that the endogenous role for ARHGAP4 is inhibitory towards migration.
ARHGAP4 inhibits cell migration via its GAP domain
Since ARHGAP4 was thought to be involved in mediating actin cytoskeletal dynamics (Tribioli et al., 1996) and a knockdown of endogenous protein resulted in increased migration (figure 3.2 B), it seemed likely that ARHGAP4 was regulating cytoskeletal dynamics to inhibit migration. ARHGAP4’s GAP domain was previously shown to have enzymatic activity towards RhoA, Rac1, and Cdc42 in vitro (Foletta et al.,
2002) (figure 2.2), and was the most likely domain to lead to changes in leading edge cytoskeletal components that regulate migration. To test if the GAP activity of
ARHGAP4 was capable of mediating ARHGAP4’s inhibition of cell motility, both full length (1-965) and R562A expression vectors were transfected into NIH/3T3 cells and assessed for their ability to alter cell motility in the wound assay. In agreement with the siRNA data, overexpressing full length ARHGAP4 inhibits cell motility as early as 2 hours after wounding, and continues to be inhibitory up to 8 hours after wounding (figure
3.3). Contrary to the effects of the full length protein, cells transfected with the GAP mutant (R562A) had an increase in cell migration over control cells transfected with
131 EYFP that was pronounced by 4 hours after wounding, and grew larger by 8 hours (figure
3.3). This dramatic difference in cell motility indicates that ARHGAP4 inhibits NIH/3T3 motility in a wound assay and that GAP activity is necessary for this inhibition.
ARHGAP4 requires FCH, GAP and SH3 domains to inhibit axon outgrowth
Considering that ARHGAP4 targets to NIH/3T3 cell leading edges (figure 2.4) and growth cones (figures 2.7, 2.8) in a similar manner, it seemed plausible that
ARHGAP4 would function in the same inhibitory manner in dentate granule neurons. To test this hypothesis, dentate gyrus explants were cultured as previously described (Butler et al., 2004) and transfected with either an empty vector, full length ARHGAP4 (1-965),
R562A, 1-770, or 72-965 and assessed for protein localization . Explants were assessed for axon outgrowth 2 days after transfection and each condition was normalized to axon outgrowth for empty vector alone (figure 3.5 A). To verify transfections, ARHGAP4 fused to EYFP was transfected into explants and detected by western from explant lysates
(figure 3.4 F). Full length ARHGAP4 overexpression greatly reduced axon outgrowth from dentate explants, indicating that it was also inhibitory in this model system (figures
3.4 B, 3.5 B). Explants transfected with R562A showed an increase in the amount of axon outgrowth (figures 3.4 C, 3.5 B). These data indicate that the GAP activity of
ARHGAP4 is also necessary for inhibition of axon outgrowth. Interestingly, explants transfected with either 1-770 or 72-965 failed to alter axon outgrowth (figures 3.4 D, 3.4
E, 3.5 B). This suggested that the FCH domain and carboxy-terminal region which includes the SH3 domain are necessary for ARHGAP4 inhibition of axon outgrowth.
132 Although western analysis demonstrates that full length ARHGAP4 is detectable
in transfected dentate explant lysates (figure 3.4 F), the individual transfected cells in the
explants could not be distinguished over background by IF. Either neurons, astrocytes or both could be transfected and cause the changes in axon outgrowth from explants. To
assess if ARHGAP4’s inhibitory function was being caused indirectly by astrocytes,
astrocyte outgrowth was measured from dentate explants transfected with an empty
vector, 1-965 or R562A. There was no significant difference in astrocyte outgrowth
(figure 3.6), indicating that the changes in axon outgrowth were not due to changes in
astrocyte morphology or extensions from the explant. To further test the function of
ARHGAP4 in neurons, dissociated granule neurons were cultured at low concentrations
and transfected with an EYFP vector, full length 1-965, R562A, 1-770, or 72-965 (figure
3.7). Neurons in these cultures did not contact other neurons or non-neuronal cells and
were allowed to grow out on poly-l-lysine/laminin. Neurons were transfected at 3 DIV
and allowed to grow an additional 2 days before measuring axon outgrowth.
Interestingly, the trends in axon outgrowth from dissociated cells were very similar to
those seen for axon outgrowth from explants (figure 3.8). Overexpression of the full
length ARHGAP4 reduced axon outgrowth compared to EYFP transfected neurons,
indicating that ARHGAP4 is also inhibitory in dissociated neurons, while the R562A
expression had a significant increase in axon outgrowth. The 1-770 and 72-965 proteins
did not significantly alter axon outgrowth, supporting a role for the FCH and SH3
domains in this inhibitory phenotype. This inhibition requires proper localization to
neuronal growth cones and a functional carboxy terminus. Since there was no change in
astrocyte migration from explant cultures, it may suggest that the actions of ARHGAP4
133 could require interactions with specific proteins or other factors found in neurons and
NIH/3T3 cells but that are lacking in astrocytes.
Discussion
Before these studies, the only description of ARHGAP4’s function was in the
SAA cell line, where it was correlated to a disruption of actin stress fibers (Tribioli et al.,
1996). Although in vitro assays have demonstrated that ARHGAP4 has a functional
GAP domain (figure 2.2) (Christerson et al., 2002; Foletta et al., 2002), no one had demonstrated if the GAP activity had any functional relevance. Here, the first functional role for ARHGAP4 is described in the CNS and in migrating cells, as an inhibitor of both processes. First, siRNA mediated knockdown of ARHGAP4 resulted in an increase in migration in NIH/3T3 cells (figure 3.2), which supports the hypothesis that endogenous levels of ARHGAP4 function to inhibit cell motility. In addition, the overexpression of
ARHGAP4 inhibited cell motility (figure 3.3), further supporting this hypothesis. To understand this inhibition in more detail, a series of deletion and point mutants were used to assess the role of each functional domain in ARHGAP4. Overexpression of
ARHGAP’s GAP mutant R562A, led to opposite effects on cell motility (figure 3.3) and axon outgrowth (figures 3.5 B, 3.8), indicating that GAP activity is central to
ARHGAP4’s inhibitory role. By itself, GAP activity as a reason for inhibition is not surprising. However, ARHGAP4 also required functional FCH and carboxy-terminal domains (including the SH3 domain) to inhibit axon outgrowth (figures 3.5 B, 3.8).
Since previous data showed that the FCH domain was required for targeting ARHGAP4 to fibroblast leading edges and to growth cones (figures 2.4, 2.8), it supports the
134 hypothesis that ARHGAP4 signals to the cytoskeleton only in discrete subcellular
locations. In addition, even when properly targeted, the mutant lacking the carboxy-
terminus (1-770) had no effect on axon outgrowth (figures 3.5 B, 3.8). This finding was
very surprising, considering that many mutants can have a dominant-negative effect by
sequestering proteins needed for the endogenous protein to function. This lack of an
effect supports a model in which ARHGAP4 may be recruited when needed via its SH3
domain or proline rich repeats in its carboxy-terminus, independently of, or secondary to
the events that traffic it to sites of activity.
The inhibition of NIH/3T3 cell motility and axon outgrowth argues for similar
signaling mechanisms in NIH/3T3 cells and growth cones. The GAP activity is necessary for inhibition in both of these model systems and may be targeting the same
GTPase(s) when activated in each cell type. While rat ARHGAP4’s GAP domain is able
to stimulate GTP hydrolysis of RhoA, Rac1 and Cdc42 in vitro (figure 2.2) (Foletta et al.,
2002), there is no data showing which GTPase is the substrate in either model system in
vivo. Despite the clear inhibitory role of ARHGAP4, the GTPase which it targets to
cause inhibition is hard to predict, as no one GTPase is totally growth promoting or
growth inhibiting. ARHGAP4 may target RhoA, Rac1 and Cdc42, as well as some or
many of the other 19 Rho-family GTPases. Rac1 is often considered a growth promoting
GTPase necessary for cell migration and axon outgrowth (Wittmann and Waterman-
Storer, 2001; Wittmann et al., 2003), and is an excellent candidate as a target for
ARHGAP4 GAP activity. However, if ARHGAP4 targeted Cdc42, it could disrupt
Cdc42’s regulation of NIH/3T3 cell polarity and axon polarity (Watanabe et al., 2004),
which would inhibit the ability of both cells and axons to grow out effectively. Even
135 RhoA could be a substrate for ARHGAP4 and lead to the same inhibitory results. RhoA
can signal through Rho kinase pathways or mDia pathways to regulate actin dynamics
(Nakano et al., 1999; Faix and Grosse, 2006). While several Rock pathways have been
attributed to acto-myosin contraction and inhibition of migration and axon outgrowth
(Mills et al., 2005; Sahin et al., 2005; Wu et al., 2005; Gallo, 2006), they are also
necessary for axon outgrowth when working in conjunction with Rac1 to form and
abolish point contacts (Woo and Gomez, 2006). In addition, mDia is activated
downstream of RhoA to control polarity, actin polymerization and MT stabilization, all
events necessary for cell motility and axon outgrowth (Palazzo et al., 2001a; Wen et al.,
2004). The human ARHGAP4 protein co-immunoprecipitates with a signaling complex
that contains proteins involved in Rac1 and WAVE signaling (Cyfip1/2), a Rac1 positive
feedback loop at leading edges (the class 3 PI3K, vps34), and the RhoA effector
Diaphanous (Weiner et al., 2006), indicating that it could be regulating either one or both
of these GTPases and their respective pathways. Therefore, in vivo assays are necessary
to discern which GTPase(s) are inhibited downstream of ARHGAP4 at cell leading edges
and growth cones.
The upstream factors that could utilize ARHGAP4 are also numerous and differ
between systems. NIH/3T3 cells migrating in a wound assay are exposed to serum
factors that are known to sculpt leading edge morphology and enhance cell motility, and include but are not limited to, EGF, PDGF and LPA (Chan et al., 1998; Suetsugu et al.,
2003; Sugimoto et al., 2006). Each of these factors differentially regulates GTPase
signaling cascades and may recruit common signaling complexes, which include
ARHGAP4, to locally inhibit GTPase signaling. There are also several inhibitory
136 guidance cues that are expressed at the same time as ARHGAP4 and affect mossy fiber outgrowth in the hippocampus. Two of the best candidate pathways include the semaphorin/plexin/neuropilin pathways and the Slit/Robo signaling pathways (Itoh et al.,
1998; Chen et al., 2000; Sahay et al., 2005). Although semaphorin mediated signaling events are known to influence mossy fiber outgrowth and are expressed around the same time, there is little other evidence to suggest that ARHGAP4 is recruited by this pathway.
In contrast, ARHGAP4 is highly homologous to the slit/robo GAPs (srGAPs), and could be recruited to the Robo2 receptor in mossy fibers, in a similar manner as other srGAPs.
Robo2 is highly expressed in the postnatal dentate gyrus at the same time as ARHGAP4
(Marillat et al., 2002), however, there has been no examination of the Robo2 pathway in the hippocampus, nor the potential involvement of ARHGAP4.
The carboxy-terminus (which includes the SH3 domain), is also necessary for inhibition of axon outgrowth in both explants (3.5 B) and dissociated neurons (figure
3.8). ARHGAP4 localizes normally to both the leading edges of NIH/3T3 (figure 2.4) cells and to growth cones (figures 2.7, 2.8) when the carboxy-terminus is deleted, but neither acts as a dominant-negative mutant like the R562A nor does it recapitulate the inhibitory actions of full length (1-965) ARHGAP4. This lack of function may suggest that the 1-770 protein is misfolded and may be degraded to some extent in cells.
However, when expressed in NIH/3T3 cells the 1-770 protein migrates to the expected size on a protein gel (figure 2.4 F) and there were no obvious break down products. In addition, NIH/3T3 cells expressing 1-770 and treated with nocodazole to disrupt MTs, lost immunostaining in the perinuclear regions but not at the leading edges (figure 2.12).
This suggests that amino acids 771-965 may be able to recognize some protein substrate
137 and be sequestered along the way to the leading edge. This hypothesis would argue that
ARHGAP4 proteins lacking amino acids 771-965 could be trafficked normally to leading edges and growth cones, and that there are different signaling complexes that ARHGAP4 could be recruited by. When MTs are disrupted in cells expressing the full length
ARHGAP4, there is no mislocalization of ARHGAP4 protein perinuclear or at the leading edge (figure 2.11), indicating that there is some other binding partner for
ARHGAP4 in both the cytoplasm and at the leading edge. The SH3 domain of
ARHGAP4 can bind to several different proteins in different cell types and conditions
(Christerson et al., 2002; Foletta et al., 2002; Weiner et al., 2006), and is the most likely domain to recruit ARHGAP4 to signaling complexes. Future experiments should address what signaling proteins can bind to ARHGAP4’s SH3 domain, in both neurons and
NIH/3T3 cells. Although there may be several proteins that could bind to ARHGAP4’s
SH3 domain, it is highly likely that these proteins may recruit ARHGAP4 to similar signaling areas in both neurons and NIH/3T3 cells, including focal adhesions, leading edges with Rac1 and PI3K, or to growth and guidance cue receptors.
138
139 Figure 3.1: Wound assay model system and quantification of cell migration in
individually transfected cells
A) NIH/3T3 cells are seeded and grown until confluent. Transfections occur at 24
hours after seeding, when the cells are about 50% confluent, and then allowed to grow until confluent. When the cells are confluent, a wound is made with an 18 guage needle and cells migrate as a monolayer in the direction of the wound. Two distinct borders are easily identified. The wound edge is the border that the cells migrate over to access the wound, while the leading edge is the border where the fastest cells form the extent of cell migration into the wound. B) The rate of cell motility/migration is expressed as a ratio of those transfected cells found at the leading edge over the total number of migrating cells
between the wound edge and the leading edge (top value of formula). These values are
normalized to cells transfected with EYFP (bottom value of formula), which reflect a
random distribution of migrating cells.
140
141 Figure 3.2: siRNA mediated knockdown of ARHGAP4 increases cell migration in
NIH/3T3 cells
A) Western blot stained for ARHGAP4 protein from NIH/3T3 cell lysates that were treated with an siRNA probe against ARHGAP4 (si-ARHGAP4) or one containing silent mutations (si-Control). ARHGAP4 protein levels were decreased by 40% in the presence of the si-ARHGAP4 siRNA when normalized to an actin loading control.
ARHGAP4 was detected with an ARHGAP4 rabbit antibody followed by staining with an HRP-conjugated goat anti-rabbit secondary and ECL. Blot was stripped and reprobed for actin with the actin (C-2) antibody followed by detection with an HRP-conjugated rabbit anti mouse secondary and ECL. B) NIH/3T3 cells were seeded and grown for 24 hours before cotransfecting either the si-ARHGAP4 or si-Control probes with an EYFP vector, and then grown to confluency and wounded to induce polarization and cell migration. Cells that were treated with the si-Control probe did not differ in their cell migration compared to EYFP alone (dashed line), while the si-ARHGAP4 treated cells had a significant increase in cell migration compared to the EYFP alone cells. Data represent 3 independent experiments and are presented as ± SEM, (* p<0.05).
142
143 Figure 3.3: ARHGAP4 inhibits NIH/3T3 cell migration via its GAP domain
NIH/3T3 cells were transfected with EYFP, 1-965-EYFP or R562A-EYFP
expression vectors and grown until confluent. Once confluent, cells were wounded with
an 18 gauge needle and the cells migrated for either 2, 4 or 8 hours before fixation. Cells
were then stained for α-tubulin to visualize the wound edge and leading edge and
transfected cells were visualized by native EYFP. At each time point, transfected cells
were counted and expressed as a percentage that were present at the leading edge relative to the total number of transfected cells migrating, and this ratio was normalized to the same values attained for EYFP transfected cells. Data are representative of at least 3 independent experiments and are presented as ± SEM. To determine significance, 1-965-
EYFP and R562A-EYFP data sets were compared to EYFP (dashed lined) at each time point. (*p<0.001).
144
145 Figure 3.4: ARHGAP4 inhibits axon outgrowth from dentate explant cultures through its
FCH, GAP and SH3 domains
Dentate explant cultures were microdissected from 200 μm rat hippocampal slices and cultured on filters coated with polylysine and laminin. Explants were transfected with ARHGAP4 expression vectors on the day of culture and grown an additional 2 days before fixing and visualizing axon outgrowth by staining for βIII-tubulin, followed by detection with an oregon green conjugated goat anti-mouse secondary. Explants transfected with FL (1-965)-FLAG (B) showed a marked decrease in axon outgrowth compared to explants transfected with an empty vector (A). Explants transfected with either 1-770-FLAG (D) or 72-965-FLAG (E) did not inhibit axon outgrowth, while those transfected with R562A-FLAG (C) had an increase in axon outgrowth. Scale bar in (A) =
100 μm. F) Western blot demonstrating expression of FL (1-965)-EYFP protein after transfection with PEI from dentate explant cultures. Expressed ARHGAP-EYFP protein was detected by blotting with the JL8 anti-GFP antibody, followed by incubation with a biotinylated horse anti-mouse secondary and incubation with streptavidin-HRP, then visualized with ECL+. The blot was stripped and reprobed with a βIII tubulin antibody, followed by incubation with an HRP conjugated rabbit anti-mouse secondary and detection by ECL.
146
147 Figure 3.5: Dentate explant axon outgrowth is inhibited by ARHGAP4 in an FCH, GAP
and SH3 domain manner
A) Explants were stained for either βIII-tubulin (neurons) or GFAP (astrocytes)
and DAPI (cell bodies). Outgrowth from the explant was determined by measuring the
entire area of outgrowth and subtracting the area of the cell bodies, then normalizing this value to the size of the explant by dividing by the DAPI area. To minimize variability,
outgrowth values from transfected conditions were normalized to explants transfected
with a parent vector alone for each independent experiment. B) Dentate explants were
transfected on the day of culture and fixed 2 days later. The FL (1-965)-FLAG
transfected explants exhibited an inhibition of axon outgrowth compared to vector alone,
while R562A-FLAG increased axon outgrowth. 1-770-FLAG and 72-965-FLAG had no
effect on axon outgrowth. Data are representative of at least 3 independent experiments
and are presented as ± SEM, (p<0.05).
148
149 Figure 3.6: Dentate explant astrocyte outgrowth is not altered by ARHGAP4
Dentate explant cultures were transfected with FL (1-965)-FLAG, R562A-FLAG or vector alone, and fixed 2 days later. Explants transfected with either 1-965-FLAG or
R562A-FLAG did not have significant differences in astrocyte outgrowth compared to vector alone. Data are representative of at least 3 independent experiments and are presented as ± SEM.
150
151 Figure 3.7: ARHGAP4 inhibits axon outgrowth in dissociated granule cells in a GAP dependent manner
Dissociated granule cells from P5-7 rats were plated on polylysine and laminin coated coverslips, transfected at 3DIV and fixed at 5DIV. Neurons were visualized by staining for FLAG (B, C, D, E) or native EYFP (A). Transfected neurons were stained for FLAG and visualized with an oregon green conjugated goat anti-mouse secondary, and were also stained for L1, followed by staining with biotinylated goat anti-rabbit secondary and incubation with streptavidin conjugated alexa 594. EYFP transfected cells were visualized for native EYFP. Neurons expressing FL (1-965) (B) had shorter axons compared to EYFP controls (A), while those expressing R562A (C) had longer axons.
Axon outgrowth did not seem to be inhibited by the 1-770 (D) or 72-965 (E) proteins. F)
Due to a lack of protein in growth cones, the 72-965 image from (E) is shown with an L1 stained image to show the axon. Images were acquired with a 20x objective, scale bar =
50 μm.
152
153 Figure 3.8: ARHGAP4 inhibits axon outgrowth in dissociated granule cells in a GAP
dependent manner
Dissociated granule cells from P5-7 rats were plated on polylysine and laminin
coated coverslips, transfected at 3DIV and fixed at 5DIV. Neurons expressing FL (1-
965) decreased axon outgrowth, while R562A expression increased axon outgrowth.
Although 72-965 expression did not significantly alter axon outgrowth, there was a trend towards axon outgrowth inhibition. 1-770 expression had no effect on axon outgrowth.
Data are representative of at least 50 neurons from 3 independent experiments and are presented as ± SEM, (p<0.05).
154 Chapter 4
Mechanistic insights into the function of ARHGAP4
155 Summary
ARHGAP4 is an inhibitory GAP protein involved in NIH/3T3 cell migration and
mossy fiber outgrowth. This inhibition requires a functional GAP domain as well as an intact FCH domain and carboxy-terminus. ARHGAP4 may be regulated locally in
NIH/3T3 cells and growth cones via its FCH domain, and its localization to leading edges is dependent on an intact actin cytoskeleton. Despite these observations, the signaling pathways that ARHGAP4 functions in are still unknown. In addition, ARHGAP4’s amino terminus contains eFCH and ARNEY domains that may also be involved in
ARHGAP4’s inhibitory function. To understand the mechanisms involved a more detailed look at GTPase regulation was pursued, as well as a more in depth study of the entire amino-terminus. ARHGAP4’s amino terminus is capable of inhibiting cell migration when the FCH, eFCH, ARNEY domains are intact, and the actin cytoskeleton may be the target of this inhibition. ARHGAP4 can downregulate levels of GTP-RhoA in NIH/3T3 cells and this GAP activity may also function in other systems, including adult DRGs.
Introduction
Experiments using the actin and microtubule (MT) destabilizing drugs, cytochalasin-D and nocodazole, demonstrated that ARHGAP4 was localized to NIH/3T3 cell leading edges by an intact actin cytoskeleton (figures 2.11, 2.12, 2.13). This localization did not depend on its carboxy-terminus, which includes the SH3 domain
(figure 2.12), but targeting to leading edges does require amino acids 1-71, which
comprises the most conserved region of the FCH domain. In addition, there are several
other regions in ARHGAP4’s amino-terminus which may contribute to its inhibitory
156 actions on cell migration and axon outgrowth. The eFCH domain is thought to be an
extension of the FCH domain which may mediate binding to lipid moieties via conserved
basic residues found interspersed along its length (Tsujita et al., 2006). ARHGAP4’s
eFCH domain also contains the conserved ARNEY domain (figure 1.6), which may be a
functional unit or an important component of the FCH and eFCH domains. Here we show that ARHGAP4’s amino terminus may be more than the sum of its parts. The
amino terminus is capable of inhibiting cell migration, but only when the ARNEY
domain is included. In addition, cells expressing the FCH/eFCH/ARNEY domain protein
have a disrupted actin cytoskeleton at their leading edges.
The regulation of axon outgrowth and cell migration is highly dependent on
GTPase signaling events that control the actin and MT cytoskeletons (Rodriguez et al.,
2003). Signaling proteins transduce extracellular cues to activate or inhibit GAPs and
GEFs, and these proteins in turn dictate when and where specific GTPase signaling cascades are activated (Bernards, 2003). The spatial and temporal control of GAPs and
GEFs is also under strict control in cells, and this control is mediated by their conserved domains that interact with specific protein signaling cascades, localize them to sites of action or are needed to be regulated by upstream factors (Bernards and Settleman, 2004).
Understanding the particular function of a GAP or GEF involves understanding how they
respond to extracellular cues and what changes are incurred in the cytoskeletal elements
downstream of their actions. In particular, determining which GTPases are targets of a
particular GEF or GAP in cells can narrow the scope of potential signaling pathways in
which any GEF or GAP may be participating. Here we show that ARHGAP4 can
downregulate levels of GTP-RhoA in NIH/3T3 cells, despite a lack of specificity in vitro.
157 The upstream signals that utilize ARHGAP4 are still unknown, but many
inhibitory pathways are potential candidates, including the proteoglycans. Similar to mossy fiber axons, dorsal root ganglion (DRG) neurons can grow out and regenerate into adulthood, but after injury glial scars develop and are enriched in inhibitory proteoglycan proteins (Carulli et al., 2005). These proteoglycans prevent axon regeneration and the signaling pathways that regulate this inhibitory function are still widely unknown, but are thought to require signaling events that RhoA and its effector ROCK (Monnier et al.,
2003). Here we demonstrate that overexpression of the R562A mutant may partially
block some of the inhibitory actions of the proteoglycan, aggrecan.
Methods
DNA construct generation
Vectors encoding amino acids 1-241, 1-289 and 1-536 of ARHGAP4 were
generated in frame to a carboxy terminal EYFP protein using the pEYFP-N1 vector
(Clontech, Mountainview, CA). The 1-71 forward primer (described in chapter two methods) was the forward primer for all the PCR reactions. Reverse primers included:
(1-241) 5’ CAGAATTCTCCGCCCTCCTTTCTTGAGAGAG 3’, (1-289) 5’
CAGAATTCGGCAATCCATGAGGTCCAAGAT 3’, and (1-536) 5’
CTGAATTCCGCCTGTGCTCTGGATAAACTTC 3’ (1-536), with novel EcoRI sites introduced in bold.
158 NIH/3T3 cell culture
NIH/3T3 cells were obtained from the American Type Culture Collection (ATCC,
cell line CRL-1658) and cultured according to ATCC protocol using Dulbecco’s
Modified Eagle’s Medium (DMEM) supplemented with 10% bovine serum (BS).
Transfections
NIH/3T3 cells were transfected using Lipofectamine (Invitrogen) according to the
manufacturer’s protocol when cells were 50% confluent (24 hrs after seeding), and
assessed 24-48 hrs later. DRGs were transfected according to the manufacturer’s
instructions without modifications.
Antibodies and flourescently labeled reagents
The following antibodies were used at the stated concentrations for immunofluorescence (IF) or western blot (WB) as indicated: JL8 anti-GFP (WB 1:1000,
Clontech); βIII tubulin (IF 1:1000, Sigma); RhoA (26C4) (WB 1:200, Santa Cruz
biotechnology); Rac1 (WB 1:1000, Upstate Biotechnology, Lake Placid, NY); Cdc42 (B-
8) (WB 1:200, Santa Cruz biotechnology). Western blots were subjected to the
horseresdish peroxidase (HRP) conjugated rabbit anti-mouse secondary antibody
(1:10000, Sigma) and detected by ECL. F-actin was visualized by incubating cells with
flourescently conjugated phalloidin 594 (Molecular Probes) according to manufacturer’s
instructions.
159 GTPase loading assays
NIH/3T3 cells were cultured in 10 cm tissue culture dishes and transfected with
either a Tag4A empty vector or a vector encoding full length ARHGAP4, and grown for two days. On the second day, the media was replaced with reduced serum Optimem media (Invitrogen) overnight and then stimulated for 10 minutes the next morning with
DMEM supplemented with 10% bovine serum. After 10 minutes, cells were rinsed with
cold PBS and then collected in Magnesium lysis buffer (50 mM Tris pH 7.5, 10 mM
MgCl2, 0.3 M NaCl2, 2% Igepal) + Protease inhibitor cocktail (Sigma). Lysed cells were
centrifuged at 10,000 xg for 15 minutes to pellet insoluble debris, and remaining lysates
were combined with PAK-PBD or Rhotekin-RBD protein GST beads (Cytoskeleton) for
1 hour at 4°C and rotated. Beads were then pelleted and washed three times and
resuspended in protein loading buffer. Bead and lysate fractions were subjected to SDS-
PAGE and probed for RhoA, Rac1 or Cdc42.
DRG culture
Dorsal root ganglion (DRG) cells were a generous gift from the lab of Dr. Jerry
Silver (Case Western Reserve University) and were prepared as previously described
(Davies et al., 1997). Briefly, dorsal root ganglia from lumbar levels 4-6 were dissected
from female Sprague-Dawley rats and dissociated with dispase/collagenase in calcium
and magnesium free (CMF) HBSS. DRGs were resuspended in Neurobasal A
supplemented with B27 and seeded at around 1000-3000 cells/coverslip. Coverslips were
coated uniformly with poly-L-lysine/ laminin or poly-L-lysine/laminin + aggrecan. Cells
160 were transfected on the day of culture and cultured an additional two days before fixation and staining.
Results
The FCH, eFCH and ARNEY domains are involved in NIH/3T3 cell migration
ARHGAP4 overexpression inhibited axon outgrowth (figures 3.5 B, 3.8) and
NIH/3T3 cell motility (figure 3.3), while a knockdown increased cell motility (figure
3.2), indicating that its normal function is as an inhibitory protein. Interestingly, every conserved domain contributed to this phenotype in neurons. ARHGAP4 must be properly localized to growth cones via its FCH domain to be inhibitory (figs 2.4, 2.8), but proper localization itself yielded no phenotype when the carboxy-terminus was deleted
(figures 3.5 B, 3.8), suggesting that protein-protein interactions are required for its activity. Most importantly, the inhibitory phenotype was GAP dependent, because the point mutant that lacks GAP activity (R562A) not only prevented inhibition of axon outgrowth and cell motility, but acted in a dominant negative fashion (figures 3.3, 3.5 B,
3.8). To get a better understanding of the domain contributions to ARHGAP4’s inhibitory function, fragments encoding increasingly larger pieces of the amino-terminus were used to determine what, if any, role these domains may have in cell migration.
Each fragment was generated in frame to EYFP and expressed as a fusion protein in
NIH/3T3 cells (figure 4.1 A). The 1-71-EYFP protein encodes the most conserved region of the FCH domain and was previously shown to be sufficient for an indirect interaction with tubulin (figure 2.9), as well as targeting ARHGAP4 to NIH/3T3 cell leading edges (figure 2.4) and growth cones (figure 2.8). The 1-241-EYFP protein
161 encompasses the entire FCH domain and most of the extended FCH (eFCH) domain,
while the 1-289-EYFP protein includes FCH, eFCH and conserved ARNEY motif. The
1-536-EYFP protein encodes all the domains in the amino terminus up to the start of the
GAP domain. Each protein migrated at the predicted molecular mass (figure 4.1 B) in
NIH/3T3 cells. It was hypothesized that some minimal fragment may associate with and sequester ARHGAP4 binding proteins, acting in a dominant-negative manner and increasing cell migration.
When assayed, neither the 1-71 nor 1-241 proteins had any effect on cell
migration compared to cells transfected with EYFP alone (figure 4.2). However, cells
expressing the FL (1-965), 1-289 and 1-536 had an inhibition of cell migration compared
to EYFP controls (figure 4.2). These results argue that amino acids 1-289 compose a
minimal sequence required to inhibit cell migration. Interestingly, these findings did not support the hypothesis that fragments of ARHGAP4 may sequester and block ARHGAP4 activation, but rather argue that overexpression of amino acids 1-289 are acting to inhibit cell migration in a similar manner as full length ARHGAP4. These findings could be explained by two competing hypotheses. The 1-289 fragment may inhibit migration through some gain of function, and due to its loss of several functional domains, may be associating with new proteins and activating or inhibiting other pathways. Alternatively, the 1-289 fragment may regulate the same pathway as ARHGAP4, and the combination of the FCH, eFCH and ARNEY regions can activate ARHGAP4 downstream effectors, or possibly activate endogenous ARHGAP4 through dimerization.
Previous experiments demonstrated that some domain or domains between amino
acids 1-770 are responsible for tethering ARHGAP4 to the actin cytoskeleton in NIH/3T3
162 cells (figure 2.11, 2.12, 2.13). The major domains within these amino acids are the GAP,
FCH, eFCH and ARNEY motif (figures 1.6, 1.7). It is unlikely that the GAP domain is
targeting ARHGAP4 to actin rich regions in cells, but the FCH, eFCH and ARNEY
motifs may have some ability to link ARHGAP4 to the actin cytoskeleton. In addition,
the difference in cell motility between cells expresing amino acids 1-289 and smaller
fragments may involve a disruption of actin dynamics at the leading edge. To assess if
these domains were targeting differently to actin rich areas of NIH/3T3 cell leading edges
or causing changes in the actin cytoskeleton, the 1-71 or 1-289 fragments were expressed in migrating NIH/3T3 cells and compared with F-actin. Both the 1-71 and 1-289 proteins were found to colocalize with F-actin at the leading edges of NIH/3T3 cells, and the 1-71
protein had a distribution that colocalized with leading edge cortical actin and the tips of actin fibers (figure 4.3 C). Interestingly, the actin cytoskeleton looked very different in cells transfected with 1-71 and 1-289. The 1-71 protein colocalized with but did not seem to alter the actin cytoskeleton in any way. However, when 1-289 was expressed, there were few F-actin fibers, and small circular actin filaments were visible behind the leading edge (figure 4.3 E), suggesting that that the 1-289 fragment was inducing some alteration in the leading edge actin cytoskeleton.
GAP function
Since the GAP phenotype was consistant in both NIH/3T3 cells (figure 3.3) and neurons (figures 3.5 B, 3.8), it seemed likely that one of the Rho family GTPases may be inhibited by ARHGAP4. Both our experiments (figure 2.2) and previous experiments
(Foletta et al., 2002) demonstrate that the GAP domain of ARHGAP4 can enhance GTP
163 hydrolysis on RhoA, Rac1 and Cdc42 but not Ras. However, in vitro assays do not
always translate to what happens in vivo. To determine if ARHGAP4 can act as a GAP
for RhoA, Rac1 or Cdc42 in whole cells, GTPase loading assays were performed on cells
overexpressing ARHGAP4. GTP loading assays utilize GTPase effector proteins to bind to the GTP-bound forms of GTPases from whole cells, and are used to determine how
GTPase activity changes in the presence of a particular GAP or GEF in whole cells.
NIH/3T3 cells expressing either an empty vector or full length ARHGAP4 were harvested and subjected to GTP loading assays. If ARHGAP4 acts as a GAP towards any of the GTPases, than lower levels of GTP-bound GTPases will be evident in the presence of full length ARHGAP4 compared to empty vector on western blots when normalized to total GTPase levels in the lysates. Western blots revealed that levels of GTP-bound Rac1 and Cdc42 were not visually different between ARHGAP4 overexpression and empty vector when normalized to lysates (figure 4.4). However, there was a dramatic decrease in the levels of GTP-bound RhoA with ARHGAP4 overexpression (figure 4.4), suggesting that RhoA may be an in vivo target of ARHGAP4 in NIH/3T3 cells.
The R562A mutant may help neurons overcome inhibitory cues
ARHGAP4’s inhibition of axon outgrowth suggests that it may be utilized by inhibitory guidance cues to modulate GTPase signaling pathways. In addition, expression of ARHGAP4’s R562A GAP mutant may abrogate signaling downstream of inhibitory guidance cue receptors. To determine if the R562A mutant could overcome inhibitory cues, EYFP and the R562A mutant were expressed in dorsal root ganglion
(DRG) neurons and assessed for their ability to grow out on laminin or laminin and the
164 proteoglycan aggrecan. DRGs were transfected with either EYFP or R562A-EYFP and grown on either laminin or laminin + aggrecan substrates (figure 4.5 A-D). Obvious trends in axon outgrowth and branching were evident on both laminin or laminin + aggrecan substrates (figure 4.5 E). DRGs expressing the R562A mutant displayed trends in increased longest axon length and total axon outgrowth, as well as more branching on laminin compared to DRGs expressing EYFP. While both EYFP and R562A expressing
DRGs on laminin + aggrecan showed decreases in axon outgrowth, those expressing the
R562A mutant had axon outgrowth levels that were equal to or greater than EYFP expressing DRGs on laminin. This suggests that the R562A protein may overcome or partially abrogate the inhibitory cues mediated by aggrecan.
Discussion
Here we describe the potential mechanisms by which ARHGAP4 may be
activated, and inhibit NIH/3T3 cell migration and axon outgrowth. It was established
that ARHGAP4 is targeted to the leading edges of migrating cells (figure 2.4) and to
growth cones (figure 2.8), and this targeting may dictate where ARHGAP4 is
functioning. This localization required an intact actin cytoskeleton (figures 2.11, 2.12),
and both the full length protein and a mutant lacking the entire carboxy terminus seemed
tethered to the leading edge actin cytoskeleton. This suggested that some domain in the
amino-terminus may be linked to the actin cytoskeleton. To further analyze the role of
the many domains in ARHGAP4’s amino-terminus, protein fragments fused to EYFP
were utilized in the wound assay model system to determine if some minimal fragment
could alter cell motility. Suprisingly, none of the fragments acted as dominant-negative
165 proteins, but those containing the ARNEY domain could inhibit cell migration (figure
4.2). Since the ARNEY domain lies within the eFCH domain, this could mean that either
the ARNEY domain itself is activating some inhibitory pathway or activating full length
ARHGAP4, or that the entire eFCH domain could mediate these events. Other proteins that contain FCH and eFCH domains can dimerize in a similar manner to the lipid sensing BAR domain (Tsujita et al., 2006). It is possible that the FCH and eFCH domains comprise a functional unit that could homodimerize and lead to activation of
ARHGAP4. One complication to this hypothesis is that the 1-770 mutant had no effect
on axon outgrowth, even though it contains intact FCH and eFCH domains (figures 3.5
B, 3.8). It may be possible that the carboxy-terminus (via the SH3 or PxxP domains),
must bind to some other protein to be activated, and this activation can expose the amino-
terminal regions. Since the 1-289 protein has no other domains, it may not be masked
and has the ability to bind and activate endogenous ARHGAP4.
The actin cytoskeleton is dramatically altered when the 1-289 protein is expressed
(figure 4.3), suggesting that actin dynamics are intricately involved in ARHGAP4
signaling. It was also interesting to note that the 1-71 protein localized to actin fibers and
along the cortical actin at the leading edge (figure 4.3). Decoration of the actin
cytoskeleton by this minimal fragment suggests that amino acids 1-71 may be the minimal fragment to target and sequester ARHGAP4 to leading edges and growth cones.
Earlier experiments demonstrated that amino acids 1-71 were necessary and sufficient for indirect MT binding (figure 2.9), but could not rule out that other linker proteins were tethering this protein to the MT cytoskeleton. These data support the hypothesis that amino acids 1-71 are the minimal unit sufficient to target ARHGAP4 to the leading edge
166 actin cytoskeleton, potentially by binding proteins or lipids that are targeted to leading
edges by trafficking anterogradely along MTs.
ARHGAP4 is able to inhibit both NIH/3T3 cell migration and axon outgrowth in
a GAP dependent manner, suggesting that some GTPase is inhibited by ARHGAP4 to
mediate these events. Rac1 signaling has been attributed to several events that promote
cell motility and axon outgrowth (Ridley et al., 2003; Causeret et al., 2004), and it was
hypothesized that ARHGAP4 may act as a GAP towards Rac1 to inhibit cell migration
and axon outgrowth. However, when GTP loading assays were performed, only GTP-
bound RhoA seemed to be altered by the overexpression of ARHGAP4 (figure 4.4).
While the trend is interesting, these experiments need to be repeated to determine if these
events holds true. The RhoA/ROCK pathway is linked to acto-myosin contractility and is utilized by several inhibitory guidance cues (Amano et al., 2000; Huber et al., 2003). So, downregulation of GTP-RhoA levels would be expected to increase cell motility and axon outgrowth. However, the RhoA effector diaphanous is associated with actin
polymerization and stabilization of MTs, events required for proper cell migration and axon outgrowth (Arakawa et al., 2003; Faix and Grosse, 2006). In support of the
hypothesis that ARHGAP4 may regulate a RhoA/diaphanous pathway, ARHGAP4 was
co-immunoprecipitated with diaphanous in a signaling complex thought to regulate
neutrophil chemotaxis from HL60 cells (Weiner et al., 2006).
The upstream regulators of ARHGAP4 are still unknown, and due to the many
binding partners found for ARHGAP4’s SH3 domain (Christerson et al., 2002; Foletta et
al., 2002; Weiner et al., 2006), it is possible that there are many potential pathways.
Whatever pathways are utilizing ARHGAP4, the GAP activity seems necessary for
167 inhibition of both cell migration (figure 3.3) and axon outgrowth (figures 3.5 B, 3.8).
Since R562A elicited a dominant-negative effect in both model systems, it is possible that this mutant could overcome inhibitory cues in other model systems as well. DRGs expressing this mutant had increased trends in axon outgrowth and branching on laminin compared to EYFP. Even though there was a trend in decreased axon outgrowth on aggrecan, DRGs expressing R562A still had greater levels of axon outgrowth on aggrecan than EYFP expressing DRGs on laminin. This suggests that the R562A mutant could either partially interfere with the inhibitory signaling of aggrecan, or that it is utilizing some other pathways to promote axon outgrowth even in the presence of this inhibitory cue. These experiements did not determine the mechanism by which R562A increased axon outgrowth, but it is unlikely that aggrecan is the only inhibitory molecule that could be overcome by overexpression of R562A. However, the trends in increased axon outgrowth on an inhibitory substrate suggest that the R562A protein, inhibition of
ARHGAP4’s GAP activity, or inhibition of ARHGAP’s effector proteins may be potential therapeutic tools to help overcome inhibition of axon outgrowth after injury.
168
169 Figure 4.1: ARHGAP4 amino-terminal protein design and expression
ARHGAP4 amino-terminal protein fragments were genrated in frame to EYFP for
expression in mammalian cells. A) The 1-71 protein encodes the majority of the FCH domain (maroon) and the conserved RAEYL motif, 1-241 encodes the entire FCH domain and the majority of the extended FCH (eFCH) domain (gray + yellow), 1-289 encodes the entire eFCH domain which includes the ARNEY motif (yellow), and 1-536
encodes the entire amino terminus but does not include the GAP domain (blue) or the
SH3 domain (orange). B) EYFP fusion proteins were expressed in NIH/3T3 cells and migrate at the expected size on protein gels. Proteins were identified by blotting for
EYFP with the JL8 anti-GFP antibody, and detected with an HRP conjugated rabbit anti- mouse secondary followed by ECL.
170
171 Figure 4.2: Amino acids 1-289 are the minimal unit required to inhibit NIH/3T3 cell
migration
Full length (1-965), 1-71, 1-241, 1-289, and 1-536 proteins were transfected into
NIH/3T3 cells and grown until confluent. Cells were then wounded and allowed to migrate for 4 hours and then fixed and visualized for transfected cells. Cell migration was scored in the same manner as figures 3.2 and 3.3 and normalized to EYFP transfected alone (dashed line). The 1-965 protein inhibited cell migration, but proteins with only FCH domains (1-71) or FCH and eFCH (1-241) did not inhibit migration.
Cells expressing proteins that included the ARNEY domain were significantly inhibited compared to EYFP controls, and the addition of the entire amino-terminus inhibited migration at the same level. Data are representative of 3 independent experiments and are presented as ±SEM, (*p<0.05)
172
173 Figure 4.3: Amino acids 1-71 and 1-289 of ARHGAP4 result in different actin
phenotypes in migrating NIH/3T3 cells
1-71-EYFP and 1-289-EYFP proteins were expressed in NIH/3T3 cells and
analyzed for localization with F-actin in migrating NIH/3T3 cells. The 1-71-EYFP
protein localized to actin rich regions (arrows) that resembled actin stress fibers and
leading edge cortical actin (A-C). The 1-289-EYFP protein localized to leading edges
(D) and colocalized with actin rich regions (arrow, F). The actin cytoskeleton appeared normal in cells expressing 1-71-EYFP, with characteristic actin bundles seen during migration. However, cells expressing 1-289-EYFP had a disrupted actin cytoskeleton with a loss of actin filaments and several circular actin structures (astericks in E, F).
Scale bar = 5 μm.
174
175 Figure 4.4: Full length ARHGAP4 overexpression decreases levels of GTP-RhoA in
NIH/3T3 cells
NIH/3T3 cells were transfected with either the Tag4A empty vector or a vector encoding full length ARHGAP4 (1-965) and grown for two days before harvesting cells for protein. Cleared lysates from transfected cells were combined with beads conjugated to PAK-PBD or Rhotekin-RBD to immunoprecipitate the GTP-bound forms of RhoA,
Rac1 or Cdc42. Recovered GTPases bound to the beads and lysates were subjected to
SDS-PAGE and blotted for RhoA, Rac1 or Cdc42, and band intensities were normalized to levels of total GTPases in the lysates. Cells expressing ARHGAP4 had a noticable decrease in GTP-RhoA compared to those transfected with empty vector, while levels of
GTP-Rac1 and GTP-Cdc42 were not as different. Western blots and graphs are representative of 1 experiment for RhoA, 2 experiments for Rac1 and 1 experiment for
Cdc42.
176
E Substrate Condition Longest ± Total ± Branch ± Axon SEM Axon SEM Points SEM (μm) Length (μm) Laminin EYFP 227.3 40.6 378.9 68.5 2 0.5 R562A 297.8 60.2 697.0 149.3 4 0.7 Laminin EYFP 182.3 62.6 282.8 114.7 2.3 1.3 + R562A 228.5 59.5 470.5 12.5 3 2 Aggrecan
177 Figure 4.5: Trends in axon outgrowth and branching are altered in DRGs expressing the
R562A mutant on laminin and aggrecan
DRGs were cultured on laminin or laminin + aggrecan and transfected with vectors expressing an EYFP or R562A-EYFP proteins and grown an additional two days before fixation. DRGs expressing EYFP and R562A had greater axon outgrowth on laminin alone versus the addition of aggrecan. E) DRGs expressing R562A had a greater tendendcy for longer axons, larger total axon length and more branching on laminin and on aggrecan. Arrows: growth cones.
178 Chapter 5
General Discussion
179 There have been few examinations of the ARHGAP4 gene or protein to date, and no prior studies have examined the role of the FCH domain or entire amino terminus.
The first description of ARHGAP4, (then known as C1), correlated its overexpression with the disruption of stress fibers in SAA fibroblast cells, as well as characterizing it as a hematopoietic cell specific protein (Tribioli et al., 1996). Later studies demonstrated that the rat orthologue of ARHGAP4 was not only expressed in hematopoietic tissues, but also in the developing central nervous system (Foletta et al., 2002), particularly in areas that develop postnatally or are associated with adult neurogenesis and regeneration.
Human ARHGAP4 has also been implicated as an important polarity protein via its association with the hematopoietic specific (HEM-1) protein (Weiner et al., 2006).
Weiner et al. (2006) demonstrated that ARHGAP4’s SH3 domain could coIP with HEM-
1 and a complex of proteins involved in leading edge actin dynamics, which include
Vps34 (human PI3K subunit), diaphanous, and components of the WAVE inhibitory protein complex. Human ARHGAP4’s SH3 domain was also shown to bind to MEKK1
(Christerson et al., 2002), a protein that is known to associate with alpha-actinin and localize to actin stress fibers and focal adhesions (Christerson et al., 1999). All of these studies implicated that ARHGAP4 was involved in pathways with cytoskeletal modifying proteins, yet no mechanism or function was determined for ARHGAP4 in any of these systems.
ARHGAP4 is a large gene with 22 coding exons that may be alternatively spliced.
While this study focused on manipulating the levels of full length ARHGAP4, the function of other isoforms is still unknown. Many lines of evidence suggest, but do not prove, that other ARHGAP4 isoforms exist. First, an antibody against ARHGAP4’s
180 amino terminus, (which was raised against residues in exon 1), reproducibly identifies several bands by western blot, which seem to be developmentally regulated (figure 2.5 B,
(Foletta et al., 2002). Even though the full length ARHGAP4 is detected in NIH/3T3 cells, the predominant band is 80 kDa, not the 120 kDa attributed to the full length protein. It is possible that some of these bands could be breakdown products. However, reverse transcriptase (RT)-PCR from rat hippocampus also identifies several different
ARHGAP4 isoforms which may be the templates for the different protein isoforms seen by western analysis (data not shown).
Interestingly, ARHGAP4 mRNA can be detected in the dentate granule cell layer,
CA3 and CA1 areas of the hippocampus using a probe that recognizes the central region of ARHGAP4 (Foletta et al., 2002). Yet when an antibody was used that only recognizes exon 1, protein was only visible in the mossy fiber axons of the dentate granule cells
(figure 2.5). This might indicate that protein is not made in the pyramidal cells of CA3 and CA1 even though mRNA is detectable, or that the protein made in these cells lacks exon 1 and cannot be detected with this antibody. If this is the case, then it raises the possibility that different isoforms could be expressed in different cell types and these
ARHGAP4 isoforms may have differing functions based on the combination of domains present in each. Furthermore, the combination of various ARHGAP4 isoforms could elicit unique functions in cells. In support of this, expressing amino acids 1-289 of
ARHGAP4 in NIH/3T3 cells inhibited cell migration in the same manner as overexpression as full length (figure 4.2), and disrupted the actin cytoskeleton (figure
4.3). Future experiemnts will need to consider what other ARHGAP4 isoforms are expressed in these model systems to fully understand their functions. RT-PCR products
181 from hippocampus could be cloned and sequenced to determine which domains are
present in these splice forms, and these splice variants could also be expressed in neurons and in migrating cells to assess if they also regulate cell migration and axon outgrowth like full length ARHGAP4.
This study sought to expand the knowledge of the full length ARHGAP4 protein and determine its role in neurons and migrating fibroblasts. To assess the role of
ARHGAP4 in these model systems, a structure-function approach was utilized. Through the deletion of critical regions, point mutagenesis, and expression of minimal fragments,
a model has been proposed to explain how ARHGAP4 localizes to sites of action and
functions to inhibit axon outgrowth (Figure 5.1). It was found that the carboxy-terminus
of ARHGAP4 was necessary for inhibition of axon outgrowth in both dentate explants
and dissociated granule cells (figures 3.5 B, 3.8). While the SH3 domain is the main
functional domain in this region that was deleted, there is the possibility that three PxxP
motifs could act as protein-protein interaction domains as well. This region had no effect on the localization of ARHGAP4 to growth cones (figure 2.8) or the leading edges of migrating fibroblasts (figure 2.4). While other studies have demonstrated that the SH3 domain of ARHGAP4 can associate with HEM-1 (Weiner et al., 2006) and MEKK1
(Christerson et al., 2002) in some cell lines, it may bind to other proteins in different model systems. In support of this, ARHGAP4’s SH3 domain was found to interact with dendrin (Foletta et al., 2002), a protein translated in dendrites of dentate granule cells
(Herb et al., 1997), and amyloid beta (A4) precursor protein-binding, family B, member 1 interacting protein (Apbb1ip) (accession # XM_225631) (Scott Young, personal communication). It is plausible that the role of ARHGAP4’s SH3 domain and carboxy
182 terminus varies from cell to cell, without altering the downstream events mediated by
ARHGAP4. This hypothesis would assume that different upstream regulators, like
guidance cue receptors, or signaling proteins that harbor PxxP rich regions, could recruit
ARHGAP4 via its SH3 domain to locally regulate GTPase levels. Interestingly, even
with proper localization, the 1-770 mutant does not inhibit axon outgrowth like the full-
length protein (figures 3.5 B, 3.8), indicating that the carboxy-terminus is critical to its
inhibitory function. Moreover, if the 1-770 mutant could still bind to effector proteins a
dominant-negative effect might have been expected, instead of inhibition, resulting in
increased outgrowth. However, neither the normal inhibitory function nor a dominant
negative role was found for this mutant.
These results suggest that the carboxy-terminus may need to be activated by some
signaling pathway or be bound by some upstream factor in order for the rest of the
protein to be active and recruit effector proteins. The most homologous proteins to
ARHGAP4, the srGAPs, function in such a manner. When the srGAPs (via their SH3
domain) are bound to the CC3 cytoplasmic domain of an activated Robo receptor, their
GAP activity is activated to target Cdc42 (Wong et al., 2001).
While many labs refer to ARHGAP4 and the srGAPs in the same vein, no one has
reported on the ability of ARHGAP4 to function as a GAP downstream of Slit/Robo
signaling. One of the reasons for this may be that screens for Robo receptor binding
proteins have been carried out on embryonic libraries, and the full length ARHGAP4 is
not highly expressed at this time. ARHGAP4 is enriched postnatally in regions where the
Robo2 receptor is known to be expressed (Marillat et al., 2002). An interaction between
ARHGAP4 and Robo2 has not been tested but is plausible, especially considering that
183 ARHGAP4 seems to be acting in the expected manner for a member of this family (i.e.
growth cone enriched, requires functional carboxy terminus to be inhibitory, GAP
activity decreases axon outgrowth, and is highly homologous to known srGAPs). This
hypothesis could be tested by expressing ARHGAP4 or its SH3 domain in neurons that
express the Robo receptors, performing immunoprecipitation experiments for ARHGAP4
and probing for an interaction with Robo receptors either without or in the presence of
Slit, which is predicted to mediate the recruitment of srGAPs to the Robo receptors
(Wong et al., 2001). If ARHGAP4 could transduce Slit/Robo signaling to postnatal
granule cell axons, than disruption of ARHGAP4 function may not only disrupt the
normal targeting of mossy fiber axons during development, but may also disrupt the
ability for newly born granule cells to integrate into the restrictive adult environment.
ARHGAP4 misregulation could lead to miswiring of excitatory mossy fibers in the dentate gyrus, and cause excitotoxicity of neurons and potential epileptic seizures.
Mossy fibers are thought to sprout during temporal epilepsy and form new synapses,
potentiating excitatory feedback (Buckmaster et al., 2002), and ARHGAP4
overexpression could also be used as a therapeutic tool to prevent this unwarranted
growth in people suffering from temporal lobe epilepsy. Improper targeting of mossy
fiber axons during development or in the adult could also put an animal or human at risk
for learning and memory deficits. To assess this, ARHGAP4 could be knocked out or
overexpressed locally in the hippocampus, and these mice could be assessed behaviorally
for basic learning and memory deficits (Gerlai, 2001). If ARHGAP4 is found to be a
determining factor in proper mossy fiber development and learning memory, it would be
of interest to determine if ARHGAP4 protein levels are altered or if mutations could be
184 found in human patients suffering from mental retardation, epilepsy or learning and
memory behavior deficits.
One of the main foci of this study dealt with the role of the fes/fer/fps/cip4
homology (FCH) domain. Several ideas describe the role of the FCH domain, including
a binding motif for actin (Yeung et al., 1998), MTs (Tian et al., 2000), and specific lipid moieties (Tsujita et al., 2006). FCH domains may be promiscuous domains that can do all of the above, or each of these findings could be examining the consequences of overexpression or circumstantial evidence. Studies with MAYP (Yeung et al., 1998) and cdc15p (Lippincott and Li, 2000) implicate actin binding, while binding to tubulin has come from studies examining rapostlin (Fujita et al., 2002) and CIP4 (Tian et al., 2000).
When rapostlin and CIP4 were overexpressed, they could be found bound colocalize with
MTs in cells, and in vitro assays demonstrated binding to tubulin. However, a CIP4 splice form (CIP4/2), which is similar to CIP4 except for an additional 56 amino acid insertion in its middle is not able to bind MTs (Chang et al., 2002), even though it retains other functions attributed to CIP4 and a nearly identical FCH domain. Instead, it associates with the plasma membrane in response to insulin activation.
FCH domains and their extended regions may compose a larger functional domain that is loosely homologous to the lipid-sensing BAR domain (Itoh and De
Camilli, 2006). The BAR domains of several proteins have been crystallized and found to form concave hydropathic structures with exposed basic amino acids that can act as binding sites for curved membranes upon dimerization (Gallop et al., 2006). These positive residues are often interspersed arginines, and mutations of these residues in FCH domains as well as extended FCH domains of several proteins prevents them from
185 binding to specific lipid moieties (Tsujita et al., 2006). ARHGAP4 is also somewhat homologous to these FCH + eFCH domain proteins (figure 1.6) and may possess similiar functions. The FCH + eFCH domain crystal structures have revealed critical residues that could aid in a better understanding of these domains (Shimada et al., 2007). The many basic residues within the FCH and eFCH domains are critical for binding to lipids, and mutation of these residues should generate proteins that are unable to bind their substrates. The conserved tyrosine residue within the RAEYL motif of the FCH domain is necessary for dimerization of these domains, and mutations in this residue could be used as a tool to determine if homodimerization is necessary for localization of function of these proteins. The carboxy-terminal region of the eFCH domain is predicted to be necessary for these domains to form higher order oligomers in order to control membrane dynamics. Since ARHGAP4 seems to be associated with regions of high membrane and actin dynamics at the leading edges of NIH/3T3 cells, it may be of considerable interest to follow up on the idea that ARHGAP4 may be recruited to leading edge membranes by
PS or PI[4/5]P2, each specific lipid moieties that can bind directly to FCH/eFCH domains. Specific lipids are enriched at the leading edges of migrating cells and these lipids recruit GTP-bound GTPases to signal at leading edge membranes (Heo et al.,
2006). To assess a role in membrane binding, we could test for an interaction of
ARHGAP4’s FCH + eFCH domains with PS or PI[4/5]2 as previously described (Tsujita et al., 2006). In addition, these lipids proposed to bind to FCH + eFCH domains are heavily enriched in lipid raft fractions. We could could test for a functional role for lipid raft association by determining if ARHGAP4 or mutants are enriched in isolated lipid raft fractions, or within cultured cells and neurons by disrupting lipid rafts using methyl-β-
186 cyclodextran to deplete cholesterol (Pike and Miller, 1998), and look for mislocalization
of ARHGAP4 from cell leading edges and neuronal growth cones.
While some role has been attributed to ARHGAP4 domains during targeting and
its function in inhibition, the upstream regulators of ARHGAP4 are still unknown.
Amino acids 1-71 target to the tips of actin bundles in migrating cells and to cortical actin
at the leading edge (figure 4.3). If this targeting is due to sequestration of ARHGAP4’s
FCH domain or actual targeting to sites of action is still unclear. To test for a role in
actively targeting ARHGAP4 to cell leading edges or to growth cones, cells or neurons
expressing EYFP tagged verisons of ARHGAP4 and its mutants could be photobleached
at cell leading egdes or growth cones and functional recovery after photobleaching
(FRAP) studies would test if constructs with a functional FCH domain were actively
moving to these photobleached areas in live cells.
ARHGAP4 could be part of signaling complexes at the leading edge or within focal contacts and adhesions. Within growth cones, several guidance cues may utilize
ARHGAP4. While the number of possible upstream regulators is great, determining what pathways could use ARHGAP4 would help immensely in understanding how to use
ARHGAP4 as a therapuetic tool for regenerating axons. To determine which pathways could be involved, ARHGAP4 could either be acutely knocked down in slice cultures or overexpressed in the mossy fibers of the hippocampus in slice culture. In this model system, the endogenous cues are all in place, and since many guidance cue defects are often localized to very distinct areas, any pathfinding errors could point out what potential pathways may be candidates to study in the future.
187 Guidance cues are not the only potential upstream regulators. One recent report
suggests that metabotropic GABA B receptor signaling could also influence levels of
ARHGAP4 (Ghorbel et al., 2005), indicating that neuronal activity may also utilize
ARHGAP4 to control axon outgrowth or potentially synapse development and plasticity.
ARHGAP4’s inhibition of NIH/3T3 cell migration (figure 3.3), mossy fiber
outgrowth (figures 3.5 B, 3.8), and similar trends in DRG axon outgrowth (figure 4.5) support the hypothesis that its GAP activity is necessary for this inhibition. However,
GTP-loading assays demonstrated that ARHGAP4 targets RhoA, and not Rac1 or Cdc42
in NIH/3T3 cells. Classic studies examining the roles of these GTPases have established
a dogma whereby Rac1 and Cdc42 are often considered the growth promotive GTPases
while RhoA is considered the growth inhibitory GTPase. While there is a great deal of
evidence to support these roles in both migrating cells and axons, what actually happens
is not as clear cut, especially for RhoA.
The RhoA/ROCK pathway can lead to actomyosin contraction and is associated
with cells in a static state (Amano et al., 2000), and increased ROCK activity is also
known to decrease levels of active Rac1 (Salhia et al., 2005). However, recent reports
have demonstrated a growth promotive role for active RhoA in cell migration and axon
outgrowth (Arakawa et al., 2003; Vicente-Manzanares et al., 2003; Wen et al., 2004).
The chemokine SDF1α can promote axon outgrowth from cerebellar granule cells, and
does so by activating RhoA and its effector mDia1 (Arakawa et al., 2003). Why RhoA
would select mDia1 in one case and ROCK in another has been an area of interest. This
selectivity may be regulated by phosphorylation of RhoA on serine 188 by PKA, which is
activated downstream of NGF activation of TrkA. When RhoA is phosphorylated on
188 ser188, it can bind to its effectors Rhotekin, PKN and mDia with much higher affinity,
while affinity for ROCK is decreased (Nusser et al., 2006). In addition, RhoA activity and effectors are spatially restricted in cell leading edges and trailing rears. Localized
RhoA and mDia activation is prominent in the leading edge membranes of migrating cells and are important for proper cell migration, while ROCK is more diffuse (Kurokawa and Matsuda, 2005). This level of spatial control opens up the possibility that locally inhibiting RhoA could lead to decreased axon outgrowth and NIH/3T3 cell migration if it is inhibited when it is normally complexed with mDia. ARHGAP4 is localized to the leading edges of migrating NIH/3T3 cells, and may be locally inhibiting RhoA when it is complexed with mDia or some other diaphanous-related formin. In support of this hypothesis, ARHGAP4 was co-immunoprecipitated with diaphanous from HL60 cells
(Weiner et al., 2006). To understand if ARHGAP4 is inhibiting NIH/3T3 cell motility and axon outgrowth by regulating a RhoA/diaphanous pathway, further experiments need to be performed to see if ARHGAP4 could selectively alter levels of active ROCK or mDia in cells.
Even though ARHGAP4 does not bind directly to MTs (figure 2.9), if it alters levels of active mDia by decreasing levels of GTP-bound RhoA, it may inhibit signaling events that locally stabilize MTs and increase actin polymerization at the same time.
MTs are also targets of Rho GTPase activity, and mDia can bind to the +-tip proteins
EB1 and APC to stabilize MTs and promote cell migration (Wen et al., 2004). Localized inhibition of mDia or other diaphanous-related formins may lead to inhibition of growth promotive events at the leading edge of migrating cells and in growth cones. To determine if ARHGAP4 is locally inhibiting specific GTPases at cell leading edges and
189 in growth cones, ARHGAP4 could be coexpressed with Flourescence Resonance Energy
Transfer (FRET) probes for Rac1, Cdc42 or RhoA (Nakamura et al., 2006) and assessed to see if local GTPase activity changes with ARHGAP4 or its mutants.
Overall, ARHGAP4 is a potent regulator of cell migration and mossy fiber outgrowth, and its signaling events could be major targets of therapeutic interventions to enhance axon regeneration and promote repair. This study has attempted to dissect out the function of each conserved domain, but more work needs to be done to understand this protein in more detail. Interestingly, expressing ARHGAP4 in a cell is not sufficient in itself to inhibit process outgrowth. Although there were inhibitory roles attributed to
ARHGAP4 and its GAP activity in neurons and migrating NIH/3T3 cells, astrocytes in dentate explant cultures did not change their migration/outgrowth properties when expressing ARHGAP4 or the R562A mutant (Figure 3.6). One hypothesis is that
ARHGAP4 mRNA could be expressed in astrocytes but is not utilized unless signaled to, and these mRNA levels could be assessed from astrocyte cultures and compared to neuronal enriched populations. If this is the case, it may reveal a level of transcriptional and translational control which astrocytes and neurons respond differently to. It would also be of interest to determine if ARHGAP4 mRNA and protein levels differ in astrocytes that are quiescent compared to those activated at sites of injury.
It is also probable that novel signaling modules in neurons and NIH/3T3 cells are necessary to recruit and activate ARHGAP4’s GAP activity but are lacking in astrocytes,
or that ARHGAP4 protein is turned over rapidly in astrocytes but could be stabilized in
neurons or NIH/3T3 cells. ARHGAP4 protein is stabilized by HEM-1 complexes via its
SH3 domain (Weiner et al., 2006). To determine if HEM-1 or its neuronal enriched
190 isoform HEM-2/Nck associated protein 1 (Nap1) is the minimum binding partner
required for ARHGAP4 activation, ARHGAP4 and HEM-1 or HEM-2/Nap1 could be
coexpressed in astrocyte cultures to determine if they alter astrocyte morphology or
migration compared to expression of either protein alone. Nap1 is a highly regulated
protein required for neuronal differentiation and axon outgrowth (Yokota et al., 2007) suggesting it would not be found in astrocytes, and recruits proteins like WAVE-1 to membrane leading edges to activate a Rac1 positive feedback loop (Weiner et al., 2006) to ensure sustained axon outgrowth. ARHGAP4 may be activated specifically in neurons and NIH/3T3 cells to act as a stop signal to this Rac1 positive feedback loop, but remain dormant in astrocytes without the HEM-1/HEM-2/Nap1 protein, similar to the lack of a phenotype in neurons when ARHGAP4 lacking the COOH-terminus was expressed. We could also express ARHGAP4 and its mutants in astrocyte or neuronal and NIH/3T3 cells, immunoprecipitate ARHGAP4, and run out immunoprecipitated complexes on protein gels to identify novel bands in the neuronal and NIH/3T3 samples. We predict to find novel interactions in neurons and in NIH/3T3 cells with proteins like Nap1 that associate with ARHGAP4 via its SH3 domain. In addition, the GTPase Rac3 has similar growth promoting properties as Rac1, but has restricted expression in NIH/3T3 cells
(Joyce and Cox, 2003) and neurons but not astrocytes (Bolis et al., 2003), suggesting that it may also be an excellent candidate to interact with ARHGAP4.
Understanding how the molecular mechanisms of GTPase signaling events in cell migration and axon outgrowth are regulated can also help elucidate the mechanisms that underlie many cancer phenotypes. Both overexpression (figures 3.3, 3.5 B, 3.8) and siRNA experiments (figure 3.2) demonstrate that ARHGAP4 protein levels can impact
191 both cell migration and axon outgrowth. It is possible that ARHGAP4 mutations or
alterations in gene expression could lead to uncontrolled migratory behavior or invasion
of cells into different tissues. In support of this, ARHGAP4 expression is downregulated
in certain ameloblastomas (Heikinheimo et al., 2002), and upregulated in chemotherapy
resistant ovarian tumors (L'Esperance et al., 2006) and in plasminogen activator inhibitor-
1 stimulated D54Mg glioma cells (Hjortland et al., 2004). While these expression studies do not prove that levels of ARHGAP4 are directly linked to these abnormalities, they suggest that ARHGAP4 may be involved. Understanding ARHGAP4 signaling mechanisms in both growth cones and in migrating cells will not only help develop therapeutic tools to aid axon regeneration and outgrowth, but could also contribute to developing better treatments for people with certain cancers.
192
193 Figure 5.1: Model of ARHGAP4 activity
ARHGAP4 contains functional FCH, eFCH, GAP and SH3 domains. These domains contribute to localization and inhibition of ARHGAP4. The full length ARHGAP4 (1) is enriched in mossy fiber growth cones and inhibits axon outgrowth, potentially by local activation and subsequent disruption of the cytoskeleton. Without the functional SH3 domain (2), ARHGAP4 can localize to growth cones but can not inhibit axon outgrowth, potentially by not being able to interact with signaling modules or guidance cue receptors. Without the functional FCH domain (3), ARHGAP4 is not enriched in growth cones and has a diffuse distribution, suggesting that the FCH domain may bind to a growth cone targeted signaling complex or vesicle, or that it is sequestered by some factor only found in growth cones.
194 Chapter 6
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195
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