1 Glaxo-IMCB Group, Institute of Molecular and Cell Biology, 30 Medical Drive, Singapore 117609
2 Membrane Biology Laboratory, Institute of Molecular and Cell Biology, Singapore 117609
3 Institute of Neurology, 1 Wakefield Street, London, WC1N 1PJ, UK
*Author for correspondence (e-mail: mcbkohcg{at}imcb.nus.edu.sg)
Accepted August 11, 2001
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SUMMARY |
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Key words: GEF, GTPase, Rho family, PIX, PAK
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INTRODUCTION |
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Some of the downstream effectors of the Rho GTPases and the pathways they regulate are well studied. In particular, the formation of stress fibres, which is downstream of RhoA-GTP, requires the function of Rho-kinase/ROK (Amano et al., 1997; Leung et al., 1995). ROK can phosphorylate and inactivate the myosin-binding subunit of the myosin light chain (MLC) phosphatase (Kimura et al., 1996). This results in an increase in phosphorylated MLC, which has enhanced actin binding and bundling activity, and hence an increase in stress fibre formation. The effector proteins downstream of Rac1 in lamellipodia formation are not as well characterized although POR1 might be involved in this process (Van Aelst et al., 1996). N-Wasp mediates the link between Cdc42 and the Arp2/3 proteins in actin polymerization, which might participate in the formation of filopodia (Miki et al., 1998; Rohatgi et al., 1999). The ROK-related target MRCK is directly involved in the formation of focal complexes (FCs) and filopodia, as demonstrated by observations that a kinase inactive mutant can block these processes downstream of Cdc42 (Leung et al., 1998).
A less well characterized morphological effect of Rho GTPases is microvillus formation. These apical membrane protrusions, found on polarized epithelial cells, fibroblasts and lymphocytes, are important for the cells to sense extracellular signals. At least one member of the ERM (ezrin, moesin and radixin) family of proteins is required to drive microvillus formation by specifically cross-linking actin filaments to the plasma membrane. The activation of ERM proteins is linked to phosphorylation, phosphoinositide binding and RhoA signalling pathways (Bretscher et al., 1997; Hirao et al., 1996; Matsui et al., 1999; Oshiro et al., 1998; Shaw et al., 1998; Tsukita and Yonemura, 1999). It has also been reported that, by activating Cdc42, RhoG causes the formation of microvilli (Gauthier-Rouviere et al., 1998).
PAK is an effector kinase of Cdc42 and Rac1 (Manser et al., 1994) in promoting the breakdown of Rho-dependent actin stress fibres and focal adhesion complexes (Manser et al., 1997; Sells et al., 1997). We have isolated a PAK-interacting exchange factor (PIX) that exhibits exchange activity towards both Cdc42 and Rac1 in vitro (Manser et al., 1998). The identification of PIX demonstrates a GTPase activator being directly coupled to an effector, thereby providing specificity to the signalling pathway. In the case of T-cell receptor activation the PIX-PAK interaction is indeed required for GTPase-mediated kinase activation (Ku et al., 2001). The complex might also provide the link for cross-talk between Cdc42 and Rac1 pathways because elevated levels of ßPAK (which can be recruited by Cdc42) drive a Rac phenotype in PC12 cells (Obermeier et al., 1998; Sells et al., 1999). The PAK-PIX interaction, mediated by the SH3 domain of PIX, plays a key role in these two cell systems.
Other identified domains of PIX include the calponin homology (CH), Dbl homology (DH), pleckstrin homology (PH) and a GIT1 binding domain. Here, we demonstrate that a discrete coiled-coil C-terminal domain appears to regulate PIX function via intermolecular interactions. Although the universal pairing of DH and PH domains in Dbl-family RhoGEFs suggests that the PH domain modulates the activities of the DH domain, the solution structure of ß1PIX DH and PH domains does not reveal such a functional coupling (Aghazadeh et al., 1998).
The intracellular localization of RhoGEFs is often achieved by specific domains which associate with other proteins or phospholipids at the cell membrane. The PH domain of Dbl mediates the oncogenic activities of the protein by targeting it to specific cytoskeletal components (Zheng et al., 1996). The N-terminal PH domain of Tiam1 localizes it to the plasma membrane allowing Rac-mediated membrane ruffling and JNK activation (Michiels et al., 1997; Stam et al., 1997). The PH domain of the Ras/Rac GEF Sos is involved in membrane targeting and is preferentially localized to the leading edge of the motile cells (Chen et al., 1997). By contrast, other Dbl-family proteins Lfc and GEF-H1 are localized to the microtubule network (Glaven et al., 1999; Ren et al., 1998). The proper presentation of the exchange factors to their respective GTPases is thus critical to their biological activities.
In this paper, we report a ßPIX splice variant that is enriched in the brain. All PIX isoforms are substrates of PAK but their exchange activities are not affected by the phosphorylation. ß1PIX, but not ß2PIX, translocates to the cell periphery, where it drives formation of ruffles and microvillus-like structures. The coiled-coil region in ß1PIX appears to be responsible for its localization to the cell periphery and for mediating its cellular activities.
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MATERIALS AND METHODS |
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Full length PIX was cloned into pAS2-1 vector (Clontech) as the bait in a two-hybrid screen for PIX-interacting proteins. The N-terminal (1-360 bp) and C-terminal (2014-2331[stop-codon] bp) of
PIX were amplified by PCR with restriction sites incorporated for the convenience of cloning. Most of the cDNA fragment was cloned by insertion of the original cDNA from the internal BamHI (174 bp) to NcoI (2025 bp) sites. The cDNA library used was a human brain cDNA Matchmaker library in pAct2 from Clontech. The bait recombinant plasmid was first transformed into a reporter yeast strain (Y190), which contained the HIS3 and lacZ genes under the control of a Gal4-responsive element. The matchmaker cDNA library was then transformed into Y190 containing the bait plasmid. The His3 gene allowed a positive growth selection for clones that were then screened using the blue/white ß-galactosidase (ß-Gal) filter assay to confirm the protein interactions. An estimated 1x106 transformants were screened. After putative positive yeast clones were identified, recombinant plasmids in pAct2 were extracted and retransformed into Y190 containing the bait plasmid. The ß-Gal filter assay was repeated and positive clones were sequenced.
Generation of anti-PIX antibodies
Polyclonal anti-PIX SH3 antibodies were raised by injection of glutathione-S-transferase (GST) fusion protein of PIX (amino acids 155-545) into rabbit. The rabbit antisera obtained were tested for specificity by western blot analysis using protein lysate of rat tissues and cell lysates containing transfected Flag-tagged PIX isoforms. It was found that the antisera was able to recognize specific bands (used at 1:500 dilution). Only a single band was observed with the Flag-PIX transfected cell lysates. The antibodies were affinity purified using MBP-PIX-SH3 column. A similar method was used to purify antibodies against the serine-rich C terminus of ß2PIX. The peptide used was amino acids 556-625 of ß2PIX.
Cell fractionation
COS-7 cells were lysed in hypotonic buffer (50 mM HEPES (pH 7.3), 1 mM MgCl2) without any detergent. NaCl and PMSF were added to 0.3 M and 1 mM, respectively, to the cell lysates. The cell lysates were spun at 100,000 g to separate the S100 soluble fraction and P100 pellets. The P100 pellets were then extracted with buffer containing TritonX-100 (50 mM HEPES (pH 7.3), 1 mM MgCl2, 0.3 M NaCl, 1% Triton X-100 and 1 mM PMSF) to extract the membrane embedded proteins. After centrifugation at 100,000 g, the pellets were extracted with buffer containing 1% SDS to extract the cytoskeletal proteins. Extracts from different fraction were resolved on SDS-PAGE gels and analysed by western hybridization.
Guanine nucleotide exchange assay
The guanine nucleotide exchange activity was measured using Rac1 assay as previously described (Manser et al., 1998). The GEF activity was determined by the incorporation of [35S]GTPS (NEN) into Escherichia coli expressed and purified GST-Rac1. The PIX proteins used in these assays were purified from transiently transfected COS-7 cells using anti-Flag M2 beads (Sigma) and quantified by Coomassie staining of duplicates. Bacterially expressed GST-PAK was used to phosphorylate a set of the immunoprecipitated PIX (
2 µg) prior to the exchange assays. 8 µg of GST-PAK protein was used per reaction, which also contained 500 µM ATP in the kinase buffer (50 mM Hepes pH 7.3, 10 mM MgCl2, 2 mM MnCl2). The reaction was incubated at 30°C for 30 minutes. The reactions were stopped by addition of EDTA. The Flag-beads were washed twice with PBS buffer before proceeding with the exchange assay. In each exchange assay about 2 µg of Flag-PIX fusion protein and 5 µg of GST-Rac1 were used. The reaction mixture was incubated at 30°C. Bound [35S]GTP
S was assessed by absorption of protein onto nitrocellulose membranes and liquid scintillation counting. In each experiment, two aliquots were taken at each time. Each experiment was repeated.
In vitro kinase assay
Immunoprecipitated Flag-PAK or GST-PAK was washed with kinase buffer and then incubated in 30 µl of the same buffer containing 500 µM ATP and 10 µCi [-33P]ATP (Amersham) and 10 µg of myelin basic protein as substrate. The reaction mixtures were incubated at 30°C for 30 minutes and the reactions were stopped by the addition of the SDS sample buffer.
Cell transfection and microinjection
Relevant cDNAs were cloned into the pXJ-Flag vector for mammalian cell expression (Manser et al., 1997). pXJ-Flag-PIX was cloned similarly as described above. pXJ-Flag-ß2PIX was cloned replacing the C terminus of ß1PIX by a PCR fragment at the internal KpnI (962 bp) site. The N-terminal deletion mutant, pXJ-Flag-
N80ß1PIX, was generated by splicing a PCR fragment covering 241 bp to the KpnI site (962 bp) of ß1PIX to vector containing DNA fragment C-terminal to the KpnI site. The C-terminal truncation mutant pXJ-Flag-ß1PIX1-555 was generated by splicing a PCR fragment containing the truncated C terminus to the N terminus of ß1PIX at the internal KpnI site. pXJ-GST-ß1PIX-C-ter was constructed by cloning a PCR fragment containing nucleotides 1612-1941 of ß1PIX. The cDNA constructs were transfected into COS-7 or HeLa cells using DOSPER (Boehringer Mannheim) or SuperFect (Qiagen) using the suppliers protocol. In most cases, 8 µg of plasmid DNA was used per 100 mm dish of 80% confluent cells and 1.5 µg of plasmid DNA was used for 20x20 mm two-well chamber slide (Nalge Nunc International). Cell staining was done as previously described (Manser et al., 1997).
HeLa cells were seeded onto 20x20 mm glass cover slips and injected using an Eppendorf microinjector (no number given on the apparatus) and a Zeiss axiovert microscope. Plasmid (50 ng ml1) encoding green fluorescent protein (GFP) was injected with the cDNA of interest (50 ng ml1). The injected cells were returned to the incubator for 2-4 hours prior to fixation in 3% paraformaldehyde as described previously (Manser et al., 1997).
Immunoprecipitation
Relevant cDNAs were cloned in pXJ-Flag or pXJ-GST mammalian expression vectors (Manser et al., 1997). The cells were harvested in protein lysate buffer (50 mM Hepes (pH7.5), 0.3 M NaCl, 1 mM MgCl2, 1 mM EGTA, 10 mM ß-glycerophosphate, 10 mM NaF, 1 mM sodium vanadate, 5% glycerol, 5 mM DTT, 0.5% Triton X-100). The cell lysates were passed through a 30G syringe (x3) and were cleared by centrifugation at 10,000 g for 5 minutes. The Flag-tag proteins were isolated using anti-Flag Mab M2-beads (Sigma) and GST fusions with glutathione-Sepharose beads (Pharmacia). Protein complexes were dissociated from the beads by heating to 100°C in 1x SDS buffer for 3 minutes.
Scanning electron microscopy
HeLa cells were first plated on glass coverslips and microinjected with the DNA constructs of interest in pXJ-Flag or pXJ-HA expression vectors (Manser et al., 1997) together with pXJ-GFP plasmid DNA for identifying the injected cells. Injected cells were incubated for 2-4 hours to allow protein expression. Cells that failed to express the GFP marker were removed and remaining cells expressing the protein of interest were fixed with 1% glutaraldehyde. Samples were then gradually dehydrated using increasing ethanol, followed by critical point drying and gold sputtering. The samples were analysed with a Phillips XL30-FEG scanning electron microscope. About 30-50 cells were examined for each experiment.
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RESULTS |
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PIX isoforms are associated with different cellular fractions
DNA constructs of Flag-ß1PIX and Flag-ß2PIX were transfected into COS-7 cells to determine the distribution of these two isoforms in the cell. From the analysis of fractionated cell extracts, most Flag-ß1PIX was present in the Triton-X-100-soluble fraction (Fig. 2A). Some Flag-ß1PIX was also found in the water-soluble extract and in the SDS-soluble fraction. However, most of the Flag-ß2PIX was found in the water-soluble fraction (Fig. 2A). Very little if any was found in the Triton-X-100-soluble fraction or in the SDS fraction.
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The coiled-coil domain affects ß1PIX localization
To determine the intracellular distribution of the various PIX isoforms and the potential role of their various domains, cDNAs encoding different Flag-tagged PIX proteins were transiently transfected into HeLa cells (Fig. 2B). Both PIX and ß1PIX were distributed in the cytoplasm but prominent at the cell periphery, whereas ß2PIX was found primarily in cytoplasm and nucleus. When the coiled-coil domain of ß1PIX was deleted, the ß1PIX1-555 protein showed cytoplasmic and nuclear localization similar to that of ß2PIX. By contrast, removal of N-terminal SH3 domain (
N80ß1PIX) did not affect the peripheral membrane localization of ß1PIX. By itself, the coiled-coil C-terminus of ß1PIX could localize GST to the cell periphery. These results indicated that the coiled-coil domain is important for the peripheral localization of
PIX and ß1PIX, and might represent the key targeting sequence for certain
PIX and ß1PIX proteins.
ß1PIX but not ß2PIX can form homodimers via the coiled-coil domain
In a two-hybrid screen for PIX partners, full-length PIX was found to interact with a construct containing a C-terminal portion of
PIX (residues 662-776). This C-terminal region of
PIX also interacted with ß1PIX, indicating that PIX homo- and heterodimers can be formed (data not shown). Because ß1PIX coiled-coil domain could dimerize but did not interact with ß1PIX1-555 (Fig. 3A), we conclude that ß1PIX does not interact in a head-to-tail manner. When GST-ß1PIX538-646 (ß1PIX-C-ter) was used to pull down various Flag-ß1PIX constructs, only those constructs containing the complementary C terminus of ß1PIX were precipitated (Fig. 3A). Full-length Flag-ß1PIX also brought down GST-ß1PIX. By contrast, no co-precipitation was observed between GST-ß1PIX and Flag-ß2PIX or between GST-ß2PIX and Flag-ß2PIX (Fig. 3B). Thus, the coiled-coil region is implicated in dimerization. ß2PIX appears to be monomeric because it behaves differently from the other two isoforms.
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Because PIX can form dimers and also binds tightly to PAK via its SH3 domain, we investigated whether PIX and PAK can exist as multimeric complexes in the cell. GST-PAK, HA-ß1PIX and Flag-N80ß1PIX were transfected together into COS-7 cells. Flag-
N80ß1PIX could be detected in the GST-PAK complex. The results indicated that PAK binds to PIX in dimeric form (Fig. 3C, lane 2). GST-PAK itself does not precipitate with
N80ß1PIX because the SH3 domain that binds PAK is missing in this mutant (Fig. 3C, lane 1). When DNA constructs of GST-PAK, HA-PAK and Flag-ß1PIX1-459 (lacking the dimerization domain) were transfected together into COS-7 cells, Flag-ß1PIX1-459 was found in the complex but HA-PAK could not be detected in the GST-PAK complex (Fig. 3C, lane 3). Neither could HA-PAK be recovered from the GST-PAK complex (
N80ß1PIX was included to drive non-productive dimers with endogenous ß1PIX) (Fig. 3C, lane 4). However, GST-PAK could complex to HA-PAK when ß1PIX (wild type) was present. The results suggest that a GST-PAK-(Flag-PIX)2-HA-PAK tetramer could be formed (Fig. 3C, lane 5). Hence, PAK and PIX proteins associate as multimeric complexes.
Phosphorylation of PIX has no effect on its GEF activity
PIX was first identified as a protein that both binds to and is phosphorylated by PAK (Manser et al., 1998). We immunoprecipitated various Flag-tagged isoforms and mutants of PIX from transfected COS-7 cells and subjected them to in vitro phosphorylation by GST-PAK. We found that a major PAK phosphorylation site(s) was located between residues 459 and 555 of ß1PIX (Fig. 4A, lanes 3,4). A deletion mutant termed ß1PIXpro (deletion of 460-495) was still phosphorylated by PAK (Fig. 4A, lane 6), suggesting that a prominent phosphorylation site(s) resides in ß1PIX 496-555. We have mapped the major phosphorylation sites to S525 and T526 of ß1PIX (data not shown). This region is conserved in
PIX and therefore phosphorylation might regulate a common activity among PIX proteins. One testable function is regulation of Rac1 or Cdc42 GEF activity.
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ßPIX can negatively regulate PAK
Although a truncated form of ßPIX suppresses PAK activation (Bagrodia et al., 1998) a truncated PIX was reported to enhance PAK activity (Daniels et al., 1999). We therefore compared the effects of ßPIX isoforms on PAK activity in vivo. PIX constructs were expressed with GST-
PAK (Pak1) and Cdc42 in COS-7 cells. Wild-type Cdc42 stimulates PAK activity to much more limited degree than Cdc42V12 (Fig. 4C) and allows an assessment of potential activation and suppression. GST-PAK was isolated on glutathione-Sepharose beads and assayed using myelin basic protein (MBP) as substrate (Fig. 4C). Consistent with previous observations,
PAK was inactive in the absence of Cdc42 (
2% of that in the presence of Cdc42V12) (Manser et al., 1997) and ß1PIX had no activating effect on PAK in these cells (data not shown). However, with Cdc42, ß1PIX inhibited
PAK activity. Similar inhibition did not occur with the ß1PIX1-555 mutant lacking the coiled-coil domain, and ß2PIX showed an intermediate effect (Fig. 4C). Because inhibition was also observed in a ß1PIX mutant (
N80) lacking the PAK-binding SH3 domain, it seems these effects are not mediated by the direct binding of PIX to PAK.
ß1PIX but not ß2PIX drives the formation of membrane ruffles
We previously demonstrated that PIX causes morphological changes in HeLa cells consistent with Rac1 activation. However, these are somewhat different from the morphology of Rac1V12-producing cells (Manser et al., 1998). Microinjection of
PIX and ß1PIX plasmid DNA caused similar phase-dark ruffles at the cell periphery (Fig. 5A). This morphological change was not observed in cells microinjected with plasmids encoding ß2PIX, ß1PIX1-555,
N80ß1PIX, ß1PIX-DHm (an exchange-activity-deficient mutant) or the ß1PIX C-terminal domain. Thus, the ability of ß1PIX to generate ruffles was dependent on the integrity of domains involved with PAK binding as well as dimerization. These observations are consistent with the dependence of lamellipodia formation on PAK-PIX interactions in PC12 cells (Obermeier et al., 1998). The formation of these ruffles was blocked by co-injection with dominant negative Rac1N17 but not with dominant negative Cdc42N17 (Fig. 5B). Rac1N17 by itself did not result in any obvious change in the cell morphology (data not shown). It has been shown that PIX has exchange activity towards Cdc42 in vitro (Manser et al., 1998) and in vivo (Yoshii et al., 1999). However, ß1PIX neither induces filopodium-like peripheral structures nor drives the cell rounding that is characteristic of other Cdc42 GEFs, such as hPem2 (Reid et al., 1999) (Fig. 5C). Co-injection of ß1PIX with wild-type Cdc42 gave a phenotype not seen with Cdc42 alone, and more similar to cells overexpressing Cdc42V12 (Fig. 5C). Co-injection of ß1PIX with wild-type Rac1 elicited ruffle-like structures and enhanced cell spreading, a phenotype associated with overexpression of Rac1V12 but not wild-type Rac1 in these cells (not shown). Similarly, co-injection of ß2PIX with Cdc42 (Fig. 5C) produced an activated Cdc42 phenotype. A Rac1V12 phenotype was observed when ß2PIX was co-injected with wild-type Rac1 (data not shown). Thus, overexpression of ß1PIX or ß2PIX elicits phenotypes associated with activated Rac1 and Cdc42 only when levels of the wild-type GTPase are increased, unlike with Tiam1 or hPEM2. This activity was apparent even though a proportion of ß2PIX was targeted to the nucleus.
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DISCUSSION |
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PIX contains an additional CH domain at its N terminus that is not present in ßPIX. Although the CH domain is found in many actin-binding proteins and signalling molecules, a single CH domain is not sufficient to bind actin (Gimona and Mital, 1998). It has been proposed that actin binding requires two CH domains (Stradal et al., 1998), and so dimerization might confer actin binding on
PIX.
PIX isoforms containing coiled-coil C-termini might form heterodimers. None of the cell lines we have tested contain multiple PIX isoforms (Fig. 1C), although both PIX and ß1PIX are present in Jurkat cells (Ku et al., 2001). Membrane localization could certainly facilitate dimer formation, as in the case of Ras (Inouye et al., 2000), where dimerization of the GTPase is essential for the activation of Raf-1. Yet another example is the N terminus of amphiphysin II, which contains sequences responsible for both plasma membrane targeting and dimerization (Ramjaun et al., 1999). It has been reported recently that the DH domain of the Dbl oncoprotein forms oligomers and that oligomerization is essential for Dbl-induced transformation (Zhu et al., 2001). It was suggested that oligomerization of Dbl could result in a signalling complex that further augments and co-ordinates the GEF activities of Dbl. Clearly, dimerization of PIX provides the possibility of forming multimeric complexes with PAK, GIT1/p95PKL (Bagrodia et al., 1999; Turner et al., 1999; Zhao et al., 2000a) and associated proteins (Fig. 7). Interestingly, PAK has also recently been reported to form dimers via the Cdc42/Rac1-binding domain (Lei et al., 2000).
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Although PAK can bind PIX tightly, their subcellular location confirms our previous observations of a dynamic association between the pair. Thus, essentially all of the PAK and ßPAK are found in the cytosol, whereas
30% of PIX is membrane associated and also present in a detergent-insoluble fraction (Fig. 2A). Because active (autophosphorylated) PAK dissociates from PIX (Zhao et al., 2000b), PAK will only transiently complex to PIX at the membrane owing to the presence of kinase activators at this site (Lu et al., 1997). However, introducing a PAK inhibitor into cells allows PAK to be stabilized within FCs (Zhao et al., 2000a) (where it is usually not visible). Thus, we propose a model in which PAK can be recruited to the membrane and FCs through binding to membrane-associated PIX (Fig. 7). Indeed addition of a CAAX box to PAK has a similar effect of driving
PAK to FCs (Manser et al., 1997). This localization is further enhanced by PIX binding to GIT1, which unmasks the cryptic paxillin binding site in the GIT1 C-terminal region (Zhao et al., 2000a). Upon activation, PAK is immediately released back to the cytosol while PIX-GIT1 remains at the membrane (and FCs). This might explain why most of PAK fractionates into the cytosolic fraction, whereas PIX is distributed between the cytosol and the membrane-cytoskeleton fraction.
Localized morphological changes produced by PIX
The interactions that lead to PIX-mediated cell shape changes are complex. Although the in vitro and in vivo GEF activities of PIX are very low compared with other Dbl family members (Manser et al., 1998), PIX can apparently still cause morphological changes but in a more restricted manner. This restriction reflects PIXs localization to the membrane, where bound phosphatidylinositol-3-kinase reportedly co-operates with PIX in activating Cdc42/Rac1 (Yoshii et al., 1999). Here, we observe that only PIX isoforms that dimerize cause ruffle and microvillus-like structure formation. Nonetheless, the binding of PAK to PIX is important because the SH3 deletion mutant (
N80) is ineffective. This is consistent with previous data showing that PAK-induced lamellipodium formation is dependent upon PIX binding (Obermeier et al., 1998). We suggest that the ß2PIX isoform would become active by recruitment through as yet unidentified membrane-associated partners.
The induction of phase dark ruffles by full-length ß1PIX was not observed with ß2PIX, the exchange deficient mutant ß1PIX-DHm or ß1PIX lacking either the SH3 domain or the coiled-coil region. These results suggest that the C-terminal sequences target PIX to the cell periphery, where its exchange activity is closely linked to PAK association. These ruffles were unlike those induced by microinjection of a more active Rac1-specific GEF, Tiam1 (Michiels et al., 1995) (data not shown), which gives a similar phenotype to cells injected with Rac1V12, as reported previously (Manser et al., 1997). This suggests that ß1PIX drives a more localized production of Rac1-GTP, although co-injection of ß1PIX with Rac1 did result in cell spreading, resembling that generated by expression of Rac1V12 (data not shown). The phase-dark ruffles are not blocked by Cdc42N17, indicating that Rac1 activation is direct.
We show here for the first time that PIX also promotes localized changes on the cell membrane resembling microvilli. Microvilli and filopodia do share many similarities and components. Although villin has been recognized as a tissue-specific component, these structures can form in its absence, which explains why microvilli are seen in epithelial cells and fibroblasts that do not express villin. The structures induced by PIX are microvillus-like, based on their position, number and ultrastructure by SEM (i.e. they are not visible by light microscopy). Such short microvillus-like structures have been reported with RhoA and RhoG, although, with the latter, they are thought to occur indirectly via activation of Cdc42 (Gauthier-Rouviere et al., 1998; Shaw et al., 1998). Consistent with this, ß1PIX-induced microvillus-like structures are blocked by Cdc42N17 but not Rac1N17. That ß2PIX and ß1PIX derivatives (ß1PIX-DHm, ß1PIX1-555, N80) do not induce microvillus-like structures confirms that the localization of (functional) PIX is important in its induction of microvillus-like structures, which is potentiated by an association with PAK.
Recent genetic evidence implicates the large GEF Trio in a pathway that includes DPak and the Drosophila Nck homologue Dock. (Newsome et al., 2000). The N-terminal GEF domain of Trio, TrioGEF1, can activate both RhoG and Rac1 (Bellanger et al., 1998; Blangy et al., 2000; Debant et al., 1996). Whether Trio and RhoG lie upstream of PIX remains to be addressed. We have not ruled out the possibility that PIX activates other Rho GTPases such as TC10 (Neudauer et al., 1998). Apart from the induction of long filopodia, TC10 also caused the formation of microvilli (Vignal et al., 2000). Although TCL is very similar to TC10, it elicits different effects on cell morphology, including long, thin extension at the cell periphery and large dorsal protrusions (Vignal et al., 2000). Hence, it is unlikely that PIX preferentially activates TCL.
Ezrin/radixin/moesin (ERM) proteins are essential for microvillus formation and their connection to PIX signalling is of interest. Antisense phosphorothioate oligonucleotide mixtures against ERM mRNAs induce disappearance of microvilli in epithelial cells (Takeuchi et al., 1994). The breakdown of microvilli is commonly observed in the early stage of apoptosis, when the ERM proteins are found to translocate from the microvilli to the cytoplasm (Kondo et al., 1997). Moesin has been reported to be phosphorylated by the Rho kinase and the related myotonic-dystrophy-kinase-related Cdc42-binding kinase (MRCK), which is an effector of Cdc42 (Leung et al., 1998; Oshiro et al., 1998). Thus, PIX at least can play a role in the activation of the ERM proteins by recruitment of MRCK.
In conclusion, we show that a C-terminal domain of PIX and ß1PIX plays a key role in the dimerization and localization of PIX, which are required to drive changes in cell morphology. A proposed model of how the PAK, PIX and GIT1 function (Fig. 7) suggests that the FC provides an important docking site, which is consistent with PAK playing a key role in these structures (Manser et al., 1997). When PAK is activated by Cdc42-GTP or Rac1-GTP, subsequent phosphorylation of PIX and GIT1 must modulate some as-yet-unidentified function. PIX isoforms are substrates of PAK but their in vitro exchange activity was unaffected by phosphorylation. By contrast, the Rac1 GEF Vav1 is potently activated upon tyrosine phosphorylation by Src-family members (Crespo et al., 1997). Although ß1PIX associates with the membrane via the coiled-coil domain, we do detect a significant amount of soluble ß1PIX, suggesting this process is regulated. As membrane association is critical for PIX to drive the local formation of microvillus-like structures and membrane ruffles, it will be important to determine what other molecular interactions play a role in this process.
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ACKNOWLEDGMENTS |
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(Kim, S., Lee, S. H. and Park, D. (2001). Leucine zipper-mediated homodimerization of the p21-activated kinase-interacting factor, beta Pix. Implication for a role in cytoskeletal reorganization. J. Biol. Chem. 276, 10581-10584)
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