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Rho GTPases control cell dynamics during growth and development. They are activated by guanine nucleotide exchange factors and inactivated by GTPase-activating proteins (GAPs). Many GAPs exist with various protein modules, the functions of which largely remain unknown. We recently cloned and identified BPGAP1 as a novel RhoGAP that coordinately regulates pseudopodia and cell migration via the interplay of its BNIP-2 and Cdc42GAP homology, RhoGAP, and the proline-rich domains. To further elucidate the molecular mechanism underlying cell dynamics control by BPGAP1, we used protein precipitations and matrix-assisted laser desorption/ionization mass spectrometry and identified cortactin, a cortical actin binding protein as a novel partner of BPGAP1 both in vitro and in vivo. Progressive deletion studies confirmed that cortactin interacted directly and constitutively with the proline-rich motif 182-PPPRPPLP-189 of BPGAP1 via its Src homology 3 domain. Together, they colocalized to periphery and enhanced cell migration. Furthermore, substitution of prolines at 184 and 186 with alanines abolished their interaction. Consequently, this BPGAP1 mutant failed to facilitate translocation of cortactin to the periphery, and no enhanced cell migration was observed. These results provide the first evidence that a RhoGAP functionally interacts with cortactin and represents a novel determinant in the regulation of cell dynamics.
Small guanine nucleotide triphosphatases (GTPases) constitute a large superfamily of molecular switches that regulate important signaling networks in cell growth, cell dynamics, and tissue/organ development. For examples, the Ras family members are involved in cell proliferation, Rho family as regulators of cell dynamics, Arf family on intracellular trafficking, whereas Ran family controls nucleus export/import (Bishop and Hall, 2000 ; Etienne-Manneville and Hall, 2002 ). These pathways are activated by certain classes of guanine nucleotide exchange factors and inactivated by GTPase-activating proteins (GAPs).
GAP domain catalyzes the conversion of the active GTP-bound form of small GTPases to their inactive GDP-bound state through a canonical “arginine finger” motif (Sprang, 1997 ). Thus far, at least 53 distinct proteins harboring the GAP domain have been identified from the human genome database (Moon and Zheng, 2003 ). To date, there is no specific GAP for a single GTPase; instead, there exists a GAP that recognizes more than one GTPases, and a single GTPase can be a target of multiple GAPs. Furthermore, in vitro substrate profile can vary compared with the in vivo results (Ridley et al., 1993 ). Although it has been well established that GAP-containing proteins usually function to negatively regulate their cognate GTPase substrates, some are believed to function as an effector; for example, the RasGAP Neurofibromatosis 1 (Yunoue et al., 2003 ) and TcGAP that is involved in insulin-stimulated glucose transport (Chiang et al., 2003 ), whereas others potentiate the action of the protein they reside in, such as the RhoGAP domain in the regulatory subunits of phosphatidylinositol 3-kinase, p85, which interacts specifically with Rac1 and Cdc42 to stimulate its kinase activity in vitro, without detectable GAP activity (Zheng et al., 1994 ). Furthermore, GAPs can be subjects of signaling cross talk by providing multiple signaling modules linked to other signaling pathways such as tyrosine kinase, phosphoi-nositites, and serine/threonine kinases. The recent findings that p190-B RhoGAP unexpectedly regulates body size of mice by affecting insulin-mediated CREB transcriptional signaling and that Rho GTPases regulate a switch between adipogenesis and myogenesis immediately open up a wealth of new and exciting prospects for testing functions of other GAPs in vivo (Sordella et al., 2002 , 2003 ). All these point to the complexity in the nature of GAP and small GTPases regulation and highlighting the need to address the function of GAPs (and other regulators) in totality, including the roles of other domains they carry. Being the key signal transducers with multiple motifs and differential substrate specificities, GAPs are poised to coordinately interact with numerous molecules to converge or diverge signals inside the cells.
We have recently identified a novel RhoGAP, termed BPGAP1, that harbors three distinctive protein domains, the BNIP-2 and Cdc42GAP homology (BCH) domain that we first described (Low et al., 1999 , 2000a ,b ), a proline-rich region (PRR) and a functional GAP domain (Shang et al., 2003 ). BPGAP1 induces the formation of pseudopodia at the cell periphery in a process that required its BCH domain and the GAP domain. Furthermore, we previously showed that these two domains collaborate with the PRR to bring about enhanced cell motility. These results suggest that changes in cell morphology are coupled to determinant(s) in cell migration.
To further elucidate the molecular mechanism underlying cell dynamics control by BPGAP1, we used protein precipitations and matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF) mass spectrometry and identified cortactin, a cortical actin binding protein as a bona fide partner of BPGAP1. Cortactin has been shown to be involved in various intracellular signaling leading to membrane ruffling, endocytosis, and motility via reorganization of actin cytoskeleton (Weed and Parsons, 2001 ). In vitro and in vivo protein interaction studies confirmed that cortactin interacted directly and specifically with BPGAP1 in a constitutive manner that required its Src homology (SH)3 domain binding to the proline-rich motif of BPGAP1 and such interaction facilitated translocation of cortactin to the membrane periphery for enhanced cell migration. These results provide the first evidence that a RhoGAP functionally interacts with cortactin and represents a novel determinant in the regulation of cell dynamics.
Cells were lysed in lysis buffer [50 mM HEPES, pH 7.4, 150 mM sodium chloride, 1.5 mM magnesium chloride, 5 mM EDTA, 10% (vol/vol) glycerol, 1% (vol/vol) Triton X-100, a mixture of protease inhibitors (Roche Diagnostics, Indianapolis, IN)], 5 mM sodium orthovanadate, and 25 mM glycerol phosphate (Sigma-Aldrich, St. Louis, MO). The glutathione S-transferase (GST)-BPGAP1 proteins, coupled to glutathione beads, were incubated with precleared cell lysates. The bound proteins were resolved by SDS-PAGE and were visualized by silver-staining (Bio-Rad, Hercules, CA). The unique bands were excised and digested with trypsin (Shevchenko et al., 1996 ). Mass spectra were acquired with a matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometer (Voyager STR BioSpectrometry work station; Applied System) operating in the delayed-extraction reflectron mode. Peptide mass fingerprints of the tryptic peptides from MALDI-TOF mass spectrometric data were used to search the National Center for Biotechnology Information protein database with the programs MS-Fit (http://prospector.ucsf.edu/ucsfhtml4.0/msfit.htm) and Mascotsearch engine (http://www.matrixscience.com).
To obtain cortactin cDNA (GenBank NP_005222; Schuuring et al., 1992 ), 5 μg of total RNA isolated from HeLa cells (RNeasy; QIAGEN, Valencia, CA) was subjected to first-strand cDNA synthesis with avian myeloblastosis virus reverse transcriptase (Promega, Madison, WI) primed with oligo(dT) (QIAGEN Operon, Alameda, CA) and amplified by high fidelity, long-template DyNAzyme (Finnzymes, Espoo, Finland) by using specific primers. Various domains were generated from the full-length template by using specific polymerase chain reaction primers with BamHI (forward) and PstI (reverse) sites for cloning into FLAG epitope-tagged pXJ40 vector (Dr. E. Manser, Institute of Molecular and Cell Biology, Singapore) and maltose binding protein (MBP)-tagged pMAL-c2 × (Dr .T.S. Sim, National University of Singapore, Singapore) or with EcoRI site (reverse) into pGEX-4T1 vector (Amersham Biosciences, Piscataway, NJ). The BPGAP1 deletion or point mutants were generated by site-directed mutagenesis as described previously (Low et al., 2000a ). The hemagglutinin (HA)-tagged BPGAP1 constructs were generated after subcloning with appropriate restriction enzymes. Clones were verified after sequencing entirely in both directions and propagated in Escherichia coli strain XL1-blue and DH5α. All plasmids were purified using miniprep or midiprep kit (QIAGEN) for subsequent use in transfection experiments. Reagents used were of analytical grade, and standard protocols for molecular manipulations and media preparation were as described previously (Sambrook and Russell, 2001 ).
293T or MCF7 cells were grown in RPMI 1640 medium supplemented with 10% (vol/vol) fetal bovine serum, 2 mM l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin (all from Hyclone Laboratories, Logan, UT), and maintained at 37°C in a 5% CO2 atmosphere, whereas HeLa cells were grown in DMEM (high glucose). They were starved for 18–24 h in serum-free medium before treatment with 10 ng/ml epidermal growth factor or platelet-derived growth factor (PDGF) (Sigma-Aldrich) for 10 min. The 293T and MCF7 cells were transfected using FuGENE 6 (Roche Diagnostics), whereas HeLa cells were transfected with Superfect (QIAGEN) according to the manufacturer's instructions.
Control cells or cells transfected with expression plasmids were lysed in 1 ml of lysis buffer as described above. Lysates were directly analyzed, either as whole-cell lysates (25 μg) or aliquots (500 μg) used in affinity precipitation/pulldown experiments with various GST fusion proteins (10 μg), or M2 anti-FLAG agarose beads (Sigma-Aldrich) as described previously (Low et al., 2000a ). For in vitro direct binding studies, immobilized GST fusion proteins were incubated with 60 ng of purified MBP-fusion proteins in 250 μl of lysis buffer at 4°C for 1 h. Samples were run in SDS-PAGE gels followed by Western blotting, and signals detected using the enhanced chemiluminescence system (Amersham Biosciences). Antibodies used were anti-FLAG (monoclonal and polyclonal; both from Sigma-Aldrich), polyclonal anti-HA (Zymed Laboratories, South San Francisco, CA), monoclonal anti-MBP (New England Biolabs, Beverly, MA), and cortactin antibodies (Cell Signaling Technology, Beverly, MA).
HeLa cells on sterilized glass coverslips were transfected with FLAG-BPGAP1 or FLAG-cortactin and HA-BPGAP1 plasmids, and then made quiescent or treated with PDGF (10 ng/ml) for 10 min. They were washed with cold phosphate-buffered saline (PBS) supplemented with 10 mM calcium chloride and 10 mM magnesium chloride (PBSCM) and fixed with 3% paraformaldehyde for 30 min at 4°C. Fixed cells were washed twice with PBSCM, twice with PBSCM containing 50 mM NH4Cl, and twice again with PBSCM, followed by permeabilization with 0.1% saponin (Sigma-Aldrich) (room temperature, 15 min) and incubation at room temperature for 1 h with 50 μl (0.5 μg) of anti-FLAG, anti-cortactin (BD Transduction Laboratories, Lexington, KY) or anti-HA polyclonal antibodies [in 2% (vol/vol) fetal bovine serum, 2% (wt/vol) bovine serum albumin in PBSCM]. Samples were washed three times (2 min) in 0.1% saponin-containing PBSCM before incubation with rhodamine-conjugated goat anti-mouse IgG (Chemicon International, Temecula, CA) or fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) or Alexa Fluor 350-conjugated goat anti-rabbit IgG (Molecular Probes, Eugene, OR). Filamentous actin was identified by staining with rhodamine-phalloidin (Sigma-Aldrich). After the final wash (5 times in 0.1% saponin containing PBSCM), coverslips were mounted with FluorSave (Calbiochem, San Diego, CA) and examined by confocal fluorescence microscopy (LSM510; Carl Zeiss, Jena, Germany). All images were captured with a 60× objective lens and presented by Microsoft PowerPoint software (Microsoft 2002).
Migration assays were carried out with six-well cluster plates (Transwell; Costar, Cambridge, MA) by using polycarbonate filters (8.0-μm pore size) coated with 0.5 μg of fibronectin (Sigma-Aldrich). Transfected cells were trypsinized and resuspended as 2.5 × 105 cell/ml in RPMI 1640 medium containing 0.2% bovine serum albumin. Cell suspension (1.5 ml) was added to the upper well and the lower wells of the chamber were filled with 2.6 ml of RPMI 1640 medium alone or 10 ng/ml PDGF in RPMI 1640 medium. After 12 h, migrated and nonmigrated cells from Transwell insert membranes were trypsinized, permeabilized, and labeled as described above, except that R-phycoerythrin–conjugated goat anti-mouse IgG (DakoCytomation California, (Carpinteria, CA) were used instead of rhodamine-conjugated goat antimouse IgG. Stained cells were suspended in PBS containing 1% paraformaldehyde and subjected to analyses by using fluorescence-activated cell sorting (FACSVantage SE). Each assay was done in at least duplicates, and the experiment was repeated at least twice. Statistical significance was analyzed with analysis of variance test. Data are means ± standard deviation (p < 0.005) and expressed as fold-stimulation over the control cells.
To better define the role of BPGAP1 in the control of cell dynamics, we used GST pulldown assays and MALDI-TOF to identify its cellular interacting partners. In addition to the full-length (FL) BPGAP1, one fragment carrying the BCH domain at the N terminus and the middle proline-rich region (NP fragment), whereas the other that harbored the same PRR together with the GAP domain at the distal C terminus (PC fragment) were used to isolate their putative targets (Figure 1A). Fragments containing PRR were used to maximize the chance of isolating any proteins that target to this region. These GST fusion proteins were coupled to glutathione-Sepharose beads and incubated with precleared lysate from the epithelial human cervical cancer, the HeLa cells. When GST-NP, GST-PC, or GST-FL were used for pulldown assays, several unique bands were consistently observed, including one with an apparent molecular weight of 80 kDa (Figure 1B, arrow), albeit with low amount. This band was excised from the gel and digested in-gel with trypsin. After this, the generated peptide mixture was analyzed by MALDI-TOF as described in MATERIALS AND METHODS. Based on their mass spectra, seven fragments could be clearly identified to be part of cortactin (Figure 1, C and D), a cortical actin binding protein known to be an important regulator of cytoskeleton. Cortactin was also identified when the same experiments were performed using lysates from the human embryonic kidney epithelial 293T cells and human breast epithelial MCF7 cells (our unpublished data).
To further confirm that cortactin was indeed a bona fide partner of BPGAP1, we used in vitro and in vivo protein interaction studies to examine the interaction of endogenous cortactin with various BPGAP1 domains that were prepared as bacterially expressed GST recombinants (Figure 2, A and B) or FLAG-tagged proteins expressed in mammalian cells (Figure 2C). Because the earlier proteomics-based pulldown assays detected cortactin in both NP and PC fragments that contained the proline-rich region (176-PPPTKTPPPRPPLP-189) as the common entity, it implied that this region could be involved in the interaction. To test this hypothesis, the N terminus without the PRR (NNP) was constructed for use in further binding studies. Figure 2A shows that when lysates prepared from HeLa cells maintained in culture condition were subjected to pulldown assays, the full-length BPGAP1 and the PC fragment interacted very strongly with endogenous cortactin, as indicated by marked enrichment compared with the cell lysates. The NNP fragment, however, did not reveal any binding to cortactin at all. Similar results were observed when MCF7 cells were used or when cortactin was overexpressed (our unpublished data). To further examine whether the interaction between BPGAP1 and cortactin is subjected to stimulation of cells by growth factors, HeLa cells were made quiescent by serum-deprivation for 24 h followed by treatment with or without PDGF for 10 min. Cell lysates were prepared and used for similar pulldown experiments. The result in Figure 2B again shows that BP-GAP1 interacted strongly and specifically via its fragment that contained the proline-rich region and that their binding did not depend on the activation of cells by PDGF.
To verify that BPGAP1 could indeed associate with endogenous cortactin in vivo, immunoprecipitation assays were carried out using lysates prepared from MCF7 cell transfected with FLAG-tagged BPGAP1 FL, NNP, NP, and PC fragments. The result shows that BPGAP1 FL, NP, and PC fragments, but not the NNP fragment, could be precipitated with endogenous cortactin (Figure 2C). Together, all these results confirm that BPGAP1 associates specifically and constitutively with cortactin in vitro and in vivo through its PRR.
Depending on the context of sequence motif, PRR could serve as cognate ligands for specific interaction with three independent protein modules, namely, SH3, WW, and Enabled/VASP homology domains (Zarrinpar et al., 2003 ). Indeed, cortactin contains an SH3 domain at its C terminus that could well be the functional interactive domain for BPGAP1's proline-rich region. To confirm our hypothesis, various deletion constructs of cortactin were made as described in Figure 3A. BPGAP1 FL, NNP, and PC fragments were expressed as FLAG-tagged proteins in 293T and subjected to pulldown assays with cortactin GST recombinants of either the full-length, or its NT, PSH3, or SH3(1) domains. As shown in Figure 3, B–D, FLAG-tagged BPGAP1 NNP fragment that lacked the proline-rich region did not bind to any of the cortactin constructs, whereas FLAG-tagged PC or BPGAP1 full-length interacted with full-length cortactin, SH3(1), or PSH3 but not the NT fragment that did not have the SH3 domain. To confirm that the flanking region (amino acid 453–494) of SH3(1) domain was not partly responsible for the binding, another set of GST recombinant proteins were made, namely, the NTP (amino acid 1–494) and SH3(2) (amino acid 495–550). These GST recombinants were used for pulldown with cells transfected with FLAG-tagged BP-GAP1 FL. The result in Figure 3E strongly suggested that the SH3 domain (amino acid 495–550) was indeed sufficient to mediate its interaction with BPGAP1. Hence, it can be concluded that cortactin associates specifically with BPGAP1 through its SH3 domain.
The PRR of BPGAP1 (176-PPPTKTPPPRPPLP-189) harbors various potential consensus target sequence motif (PXXP) for SH3 binding. To further delineate the specific motif within this region, various internal deletion (as opposed to truncation mutants used in earlier experiments) or point mutants were made as GST recombinant constructs and subjected to pulldown assays by using lysates isolated from HeLa cells that expressed endogenous cortactin (Figure 4). As shown in Figure 4B, deletion of the entire proline rich region (P1 mutant) or the KTPPPRPPLP sequence in P2 mutant both abolished the binding between BPGAP1 and cortactin, suggesting that this stretch of sequence is essential for interacting with cortactin SH3 domain. Consistent with this was the observation that P3 mutant, which still retained the sequence KTPPPRPPLP, could indeed pull down endogenous cortactin. This target sequence is unique because it differs from the consensus binding site (+PPΨPxKP) (where + and Ψ indicate basic and hydrophobic amino acid residues, respectively) (Sparks et al., 1996 ) for many partners of cortactin SH3 domain identified to date (Figure 4C). To further assess which proline residues within this region were important in mediating this interaction, proline residues at 184 and 186 were substituted with alanines (termed PP mutant) because they conform best to the consensus core motif PXXP (Figure 5A). The PP mutant was expressed as FLAG-tagged and GST-tagged recombinants for further analyses. Pulldown assays were performed, and the results show that unlike the wild-type full-length BPGAP1, the GST-PP mutant failed to interact with endogenous cortactin from HeLa cell lysates (Figure 5B) or with MBP-cortactin SH3(2) in the direct binding assays (Figure 5C). Consistent with these is the lack of coimmunoprecipitation of endogenous cortactin with FLAG-tagged PP mutant (Figure 5D).
Together, all these results confirm that PXXP core motif within the PRR of BPGAP1 is indispensable for directly targeting BPGAP1 to cortactin at its SH3 domain to form a stable constitutive complex. Furthermore, the target sequence represents a novel recognition motif for the binding of cortactin SH3 domain. Delineation of these interaction domains/motifs should allow us to probe for their possible roles in regulating cellular process as described in the following section.
We recently observed that BPGAP1 induced pseudopodia via its BCH and GAP domain independently of the PRR. However, this PRR was necessary to facilitate cell migration upon changes in cell morphology induced by both BCH and GAP domains (Shang et al., 2003 ). Because cortactin is known to be a key regulator for membrane dynamics (Weed and Parsons, 2001 ), we hypothesized that BPGAP1 could act in concert with cortactin to regulate cell migration.
We first set out to examine the subcellular localization of endogenous cortactin and the disposition of filamentous actin in HeLa cells expressing either the wild-type BPGAP1 or the PP mutant. These cells were made quiescent by serum removal or stimulated for 10 min with PDGF, followed by indirect immunofluorescence studies. Figure 6, A and B, show that under both types of condition, BPGAP1 and cortactin were mainly colocalized within cytoplasm and also with presence on the membrane periphery along with the filamentous actin (indicated by arrows). However, cells without BPGAP1 overexpression showed a strong absence of cortactin as well as reduced actin disposition on the membrane periphery. Consistent with this, cells expressing BPGAP1 PP mutant that did not interact with cortactin failed to display the presence of cortactin and actin on the membrane periphery or the protruding ends, despite the PP mutant being localized to those areas (Figure 6, C and D, arrowheads). Furthermore, we revealed that the actin staining in the protrusions formed by the PP mutant was also reduced compared with those formed by the wild-type counterpart. Similarly, when more cortactin was introduced upon cotransfection with wild-type BPGAP1, it was also markedly localized to the cell periphery with BPGAP1. However, such disposition was lost when the PP mutant was expressed in the cells (Figure 6E, arrows). These results suggest that the binding of BPGAP1 to cortactin facilitates the translocation of cortactin to the cell periphery and the assembly of actin there, in a manner independent of growth factor stimulation.
The localization of BPGAP1–cortactin-interacting complex along the membrane periphery strongly suggests a possible function for such complex in regulating cell migration, especially cortactin, which has been implicated in cell motility in both fibroblasts and endothelial cells (Huang et al., 1998 ; Patel et al., 1998 ), although little is known for its effects on cell motility in epithelial cells. We sought to further determine the effect of BPGAP1 with cortactin on cell migration in 293T cells, which when compared with HeLa cells, offered greater transfection efficiency required for the expression of both BPGAP1 and cortactin for subsequent reliable detection by FACS. Cells were transfected with the vector control, expression plasmids for HA-tagged wild-type BPGAP1 or the PP mutant, either alone or together with the FLAG-tagged cortactin and assayed for their effects on cell migration as described in MATERIALS AND METHODS. Figure 7 shows that 293T cells expressing BPGAP1–cortactin-interacting complex resulted in a significant twofold increase (p < 0.005) in the cell migration rate when cells were under quiescent or when PDGF was used as a chemoattractant. No effect was seen with the expression of either BPGAP1 or cortactin alone. Consequently, the PP mutant that did not interact with cortactin failed to stimulate any significant changes in the cell motility. These results therefore strongly support the notion that BPGAP1 interaction with cortactin facilitates cortactin translocation to cell periphery, leading to significant effect in the regulation of cell motility in epithelial cells. The significance of this is discussed and a model for such regulation is proposed below (Figure 8).
Our recent study showed that BPGAP1 modulates cell dynamics via an interplay of its multiple protein domains. The formation of pseudopodia by the BCH and GAP domains is coupled to enhanced cell motility that requires an intact proline-rich region. In the present study, we have provided molecular evidence that the cortical actin binding protein cortactin could mediate this missing link.
Being a multidomain protein itself, cortactin interacts with various signaling partners in different cell types and locations. It has the potential role as an adaptor/scaffold connecting other signaling proteins to regulate different cellular events. These include interaction with Shank proteins to organize the clustering of receptor complexes within the neuronal postsynaptic sites, binding with dynamin 2 to regulate receptor-mediated endocytosis at the membrane periphery and also involved with ZO-1 in the control of tight junction assembly of epithelial cells (Weed and Parsons, 2001 ). Besides, cortactin is also involved in the signaling pathway regulated by two small GTPases, Rac1 and Cdc42 (Weed et al., 1998 ; Vidal et al., 2002 ). The activation of Rac1 and Cdc42, together with the consequential Arp2/3 activity, is thought to be necessary for protrusive-based cell motility. In NIH3T3 fibroblasts and endothelial cells, cortactin was responsible for enhanced migration (Huang et al., 1998 ; Patel et al., 1998 ), whereas Rac1-induced signal transduction is required for the translocation of cortactin to the membrane periphery (Weed et al., 1998 ). Despite all these observations, and because cortactin itself is not directly associated with these GTPases, the actual molecular mechanisms linking cortactin to them remain unknown.
To date, no RhoGAP has been implicated in any of the cortactin-mediated signaling, although both have been widely studied for their separate cellular processes. We have established that BPGAP1 functions biochemically as a RhoA inactivator through its GAP domain while exerting its peculiar cell extensions via its BCH and GAP domains in a process that requires both active Cdc42 and Rac1 (Shang et al., 2003 ). Intriguingly, the BCH domain of BPGAP1, in addition to its ability to form homophilic and heterophilic complex as we previously identified for the BNIP-2 family proteins (Low et al., 1999 , 2000a ,b ), also selectively target Cdc42. In contrast, despite not being a substrate for the GAP domain, Rac1 is still a preferred binding partner for this domain in vitro and in vivo (our unpublished data). Coupled to the requirement of BCH and GAP domains in Cdc42/Rac-1 induced pseudopodia formation and the RhoA inactivation by GAP domain, the current identification of cortactin as the missing link for its induced cell migration would immediately implicate that BPGAP1 could serve to bridge cortactin and GTPases signaling for the concerted regulation of cell dynamics. The intricate nature of such regulation is now being investigated.
Cortactin is normally distributed diffusely in the cytoplasm of quiescent cells and upon stimulation or its overexpression, it is translocated to the periphery (McNiven et al., 2000 ; Weed et al., 2000 ; Van Rossum et al., 2003 ). However, we did not observe this translocation phenomenon in our system by using the HeLa cells, despite stimulation with PDGF, unless BPGAP1 also was coexpressed. Likewise, different from previous reports on fibroblasts and endothelial cells where cortactin alone promoted cell motility (Huang et al., 1998 ; Patel et al., 1998 ) overexpression of cortactin alone in the epithelial 293T cells did not elicit enhanced cell migration unless BP-GAP1 also was overexpressed. Such cell type specificity is likely due to the absolute requirement of BPGAP1 as a primary inducer for cortactin translocation in these cells, leading to the stimulation on cell migration. Although not measured in the current study, we believe that such inducer of cortactin translocation might already exist in great abundance to readily elicit their effects in fibroblasts and endothelial cells. Interestingly, others have shown that cortactin lacking its SH3 domain was also sufficient to translocate to the periphery upon activation of Rac pathway (Weed et al., 2000 ). Perhaps, there exists an “inhibitory” system in the cells where unknown protein(s) binding to cortactin at its SH3 domain could prevent it from migrating to the periphery. However, upon overexpression of cortactin, this inhibition could be released simply by the lack of sufficient inhibitors to sequester its translocation. Consistently, in our system where BPGAP1 was overexpressed, it is equally plausible that BPGAP1 could have displaced such unknown protein(s), thus allowing translocation probably by recruiting at the same time other cellular proteins through its BCH and/or even the GAP domain. The BPGAP1-cortactin would then be available for subsequent complex to form, leading to protrusive-based motility (Figure 8, see model). In this regard, it is interesting to note that the PP mutant that still elicited protrusions via BCH and GAP domains failed to promote significant assembly of cortactin and actin on the cell periphery or the leading edge of protrusions. These results suggest that different cytoskeletal networks could be used for cell protrusions leading to different physiological outcomes.
Our findings that cortactin binding to BPGAP1 via its SH3 domain could therefore provide a new mechanism for the translocation of cortactin to the periphery, consistent with the observations that subcellular localization of cortactin may be regulated by mechanisms independent of its tyrosine phosphorylation upon growth factor stimulation (Weed et al., 1998 ) and further supporting the notion that tyrosine phosphorylation of cortactin itself could not account fully for its role in conferring cell migration (Van Rossum et al., 2003 ).
Although the PRR of BPGAP1 (182-PPPRPPLP-189) forms a constitutive complex with cortactin via its SH3 domain, this region is unique compared with the widely regarded cortactin SH3 domain binding consensus sequence +PPΨPXKP (Sparks et al., 1996 ; Figure 4C). Apart from the conserved prolines at + 2, +5, and +8 positions, the requirement for basic residue in +1, hydrophobic residue in +4 and the lysine residue in +7 are all notably missing. Nonetheless, mutations of the two conserved prolines at +3 and +5 (P184A and P186A, respectively; Figure 5A) were sufficient to abolish the interaction. For comparison, it also had been indicated that the substitution of the alternating two prolines with alanines in cortactin binding protein 1 reduced its binding to the cortactin SH3 domain (Du et al., 1998 ). Other than that, the BPGAP1 target sequence shows little resemblance to the target sequence of several other “nonconforming” proteins (Figure 4C). For examples, for dynamin both 826PPQIPSRP833 and 849APAAPSRP856 are used (McNiven et al., 2000 ); CBP90 uses 540PPIPPKKP547 (Ohoka and Takai, 1998 ) and most recently Fgd1 with 158KPQVPPKPSYL168 (Hou et al., 2003 ). This implies that the SH3 domain of cortactin could have different degrees of substrate specificity, perhaps useful for its roles as a modulator in multiple signaling pathways.
In summary, by using proteomics, molecular and cellular tools, our current work delineates functional interaction between a novel RhoGAP and cortactin in cell dynamics control. We have identified BPGAP1 as an important regulator of cell dynamics that requires cortactin for a concerted effect on cell migration. The challenge now lies on how the PRR of BPGAP1 and the SH3 domain of cortactin could distinguish between several other known or potential partners in the confines of the living cell. We believe that this issue can be partly resolved by identification of many more other cellular interacting partners in this BPGAP1–cortactin complex or complex where these two proteins exist as mutually exclusive entities. Although many cortactin binding partners have been identified to date, our results provide the first evidence that a RhoGAP functionally interacts with cortactin and represents a novel determinant in the regulation of cell migration. It also could potentially provide a link between the small GTPases and cortactin in regulating the spatial and temporal context of cell dynamics.
We thank Sashikant Joshi and Wang Xian Hui at the Protein and Proteomics Centre for technical advice and help on MALDI-TOF analyses; Dr. Mohan K. Balasubramania at Temasek Life Sciences Laboratory for providing the facility for triple-labeling confocal microscopy; Tan Li Hui for technical help in FACS; and Dr. Jan Buschdorf and Soh Fu Ling for reviewing our manuscripts. This work is supported by a Graduate Research Scholarship awarded to B.L.L. and a grant from Academic Research Fund, the National University of Singapore and Biomedical Research Council of Singapore.
Article published online ahead of print. Mol. Biol. Cell 10.1091/mbc.E04-02-0141. Article and publication date are available at www.molbiolcell.org/cgi/doi/10.1091/mbc.E04-02-0141.