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Vol. 17, Issue 10, 4564-4575, October 2006
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Department of Pharmacology, Vanderbilt University Medical Center, Nashville, TN 37232-6600
Submitted October 28, 2005;
Revised June 29, 2006;
Accepted July 31, 2006
Monitoring Editor: Martin A. Schwartz
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Martinez-Quiles et al., 2001), axonal outgrowth to target cells (Korey and Van Vactor, 2000), and the aggregation of Dictyostelium cells to form a multicellular organism (Devreotes and Parent, 1999; Chung et al., 2001). Chemotaxis also plays a role in disease states such as arthritis, cancer, and multiple sclerosis (Bagiolini, 2001). Directional cell movement requires a defined cell polarity in which components of the cytoskeleton are differentially localized at the leading edge of a migrating cell as well as its retracting posterior (Firtel and Chung, 2000; Chung et al., 2001). This polarization can be initiated by the chemoattractant binding of heterotrimeric G protein–coupled membrane-bound receptors, and subsequent activation of downstream signal transduction pathways direct reorganization of the actin and myosin cytoskeleton (Chung et al., 2001; Van Haastert and Devreotes, 2004). A significant focus of current research involves the elucidation of the pathways that mediate both cell polarization and directional sensing, the result of which makes up a cell's proper chemotactic response.
Wiskott-Aldrich syndrome (WAS) is a human X-linked immunodeficiency characterized by recurrent infections, hematopoietic malignancies, eczema, and thrombocytopenia (Aldrich et al., 1954; Derry et al., 1994; Snapper et al., 1998). These defects are caused by mutations in the gene encoding the Wiskott-Aldrich syndrome protein (WASP), a key adaptor protein that connects multiple signaling pathways to F-actin polymerization, a process essential for chemotaxis. The WASP family of proteins includes WASP, N-WASP, and SCAR or WAVE (Miki et al., 1996; Bear et al., 1998). We have previously confirmed the necessity of WASP during the chemotactic motility of Dictyostelium via targeted knockdown of this gene (Myers et al., 2005). Lowered expression of WASP significantly impaired the ability of cells to polarize, polymerize F-actin, and migrate toward the chemoattractant cAMP. Both WASP and N-WASP possess a WH1 (WASP homology 1) domain, a basic (B) domain, a Cdc42/Rac binding (GBD) domain, a polyproline (SH3-binding) domain, a G-actin binding WH2 (WASP homology 2) domain (N-WASP has two WH2 domains), a central domain, C, and a C-terminal acidic, A, segment (Symons et al., 1996; Zigmond, 2000). The importance of the WH1 domain has been suggested by the frequency of mutations found to cause WAS (Rong and Vihinen, 2000). Despite comprising only one-fifth of WASP, more than half of the known missense mutations causing WAS are present in the WH1 domain (Derry et al., 1995). The WH1 domain has been implicated in the recruitment of WASP family members to sites of actin polymerization during motility of vaccinia virus (Moreau et al., 2000), WASP interacting protein (WIP)-induced membrane protrusions (Vetterkind et al., 2002), and PI(4,5)P2-induced intracellular vesicles (Benesch et al., 2002).
WIP has been shown to interact with human N-WASP, a ubiquitously expressed homolog, at the WH1 domain (Volkman et al., 2002). WIP is known to bind to both G- and F-actin (Martinez-Quiles et al., 2001). Depending on the system examined, WIP and other verprolin family members have been found to stimulate F-actin polymerization (Ramesh et al., 1997; Kinley et al., 2003), initiate filopodia formation (Martinez-Quiles et al., 2001; Vetterkind et al., 2002), and prevent F-actin depolymerization (Martinez-Quiles et al., 2001; Kato et al., 2002). WICH, a protein related to WIP, has been shown to induce microspike formation through cooperation with N-WASP (Kato et al., 2002). We have identified a homolog of mammalian WIP, WIPa, in Dictyostelium and examined the roles of WASP and WIP in the regulation of F-actin assembly and chemotaxis. The WH1 domain of WASP was found to bind WIPa. Overexpression of WIPa leads to an increase in microspike formation that is dependent on its WH2 domain, WASP, and VASP. WIPa translocates to the cortex of cells uniformly stimulated with cAMP in a time course that parallels F-actin polymerization. Lower levels of WIPa expression reduced prompt chemotactic responses of cells to a changing chemoattractant gradient. We conclude that WIPa is important for proper regulation of actin polymerization at sites of new pseudopod formation, which allows a chemotaxing cell the flexibility to respond to a spatially dynamic chemotactic gradient.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Molecular Biology
WASP and WIP expression constructs were produced by PCR amplification of Dictyostelium cDNA library or genomic DNA and subcloned into the pEXP4(+) vector in frame with eYFP at the N-terminus separated by the flexible linker GSGSG. The YFP fusion proteins were expressed under control of the actin-15 promoter. Point mutants were produced by cloning genes into pBS, using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA), and subcloned into pEXP4(+). All mutants were confirmed by sequence analysis. Protein expression constructs were produced using pGEX6P-1 (GST-tag) or pET-32a+ (6xHis-tag). Hairpin WIPa RNA construct was made by amplifying full-length WIPa and C-terminal half of WIPa and cloning them into pEXP4(+) in a tail to tail manner.
In Vitro Pulldown Assay
Protein expression constructs produced using pGEX6P1 or pET-32a+ were transformed into BL21 cells. Cultures were grown to a cell density of OD600 = 0.8, and 0.1 mM IPTG was added to induce protein expression for 1 h at 37°C. Proteins were purified according to the manufacturers' instructions (Sigma-GST proteins; Qiagen-His proteins). Equal quantities of recombinant protein were added to 500 µl TBS-T containing 60 µl of GST-tagged protein bound to glutathione-conjugated agarose beads and incubated at 4°C for 1 h. The agarose beads were pelleted by centrifugation at 14,000 rpm for 5 min and washed three times with 1 ml TBS-T. Protein was then stripped off the beads in 2x sample buffer by boiling for 4 min. Samples were resolved on a 12% SDS-PAGE gel, and proteins were identified via Western analysis using a mouse anti-6x His antibody (Cell Signaling, Beverly, MA). In vivo pulldown experiments were similarly performed, but agarose beads were incubated in 100 µl cell lysate, and proteins were identified using a mouse anti-GFP antibody.
In Vitro Actin Polymerization Assay
Experiments were performed as described by Zigmond, (1997). High-speed supernatants (HSS) were prepared by pulsing cells with 30 nM cAMP at 6-min intervals for 5 h, shaking with 3 mM caffeine for 30 min, lysing cells by passage through a 5-µm pore filter in PM buffer (12 mM Na/K phosphate buffer, pH 6.1, 2 mM MgSO4, plus 20 mM KCl and 1 mM EGTA), spinning for 20 min at 2.8e5 x g, and freezing in liquid nitrogen. One hundred nanomolar purified RacC was charged with either 100 µM GTP
S or 100 µM GDP
S for 10 min at 30°C. Supernatants were then stimulated at room temperature with the nucleotide-bound RacC at time 0. Aliquots of 50 µl were taken at each time point and stopped by dilution into 850 µl of actin buffer containing 6% formaldehyde, 0.15% Triton X-100, and 1 µM TRITC-phalloidin. Actin was stained for 1 h at room temperature and spun down at 14,000 rpm for 10 min. Rhodamine-phalloidin was extracted with 300 µl of 100% methanol, and fluorescence was measured (540 excitation/575 emission).
For visualization of actin filaments, supernatants were allowed to polymerize in the presence or absence of 0.4 µM phalloidin on a slide coverslip for 5 min at 37°C before stopping reactions with actin buffer containing formaldehyde and TRITC-phalloidin. Images were taken using fluorescent microscopy with a 60x objective lens and Metamorph software (Universal Imaging).
F-Actin Pulldown Assay
Experiments were performed the same as in vivo actin polymerization assays, except 100 µl of cells were mixed at each time point in Dictyostelium lysis buffer (25 mM Tris, pH 7.6, 100 mM NaCl, 1 mM EDTA, 1% NP-40, 10% glycerol, 1 mM DTT, 3 µM phalloidin). Lysates were spun down at 14,000 rpm for 10 min. Pellets were boiled in 2x loading buffer, resolved on 10% SDS-PAGE, and blotted onto nitrocellulose, and YFP-WIPa was detected using an anti-GFP antibody.
F-Actin Staining
For phalloidin staining, cells were pulsed with 30 nM cAMP at 6-min intervals for 5 h and placed on glass coverslips in 12 mM sodium phosphate buffer (pH 6.2) + 200 µM each of MgCl2 and CaCl2 for at least 30 min. Cells were fixed with 3.7% formaldehyde for 10 min. Cells were permeabilized with 0.5% Triton X-100, washed, and incubated with TRITC or Alexa594-phalloidin (Sigma) in PBS containing 0.5% BSA and 0.05% Tween-20 for 30 min. Cells were washed in PBS containing 0.5% Tween-20. Images were captured with a Roper Coolsnap camera (Tucson, AZ) and Metamorph software (Universal Imaging).
For labeling barbed-ends, cells were permeabilized with 100 mM PIPES, pH 6.9, 1% Triton X-100, 4% PEG, 1 mM EGTA, 1 mM MgCl2, 3 µM phalloidin for 3 min and 0.4 µM rhodamine-labeled G-actin in 1 µM ATP solution was added. After 5 min staining, cells were washed three times with PIPES buffer and fixed with 3.7% paraformaldehyde. For quantitation of leading edge F-actin content, the amount of F-actin at the leading edge of these cells was quantitated by measuring the integrated phalloidin signal (area x average intensity) at the leading edge divided by the average intensity of phalloidin staining throughout the cell body.
Chemotaxis Assay
Cells were pulsed with 30 nM cAMP at 6-min intervals for 5 h. These conditions maximally induce expression of aggregation-stage genes, including the cAMP receptor cAR1 and the coupled G protein 
subunit G2. The chemotaxis assays were performed as previously described (Chung and Firtel, 1999; Meili et al., 1999). After pulsing, cells were plated on glass-bottomed microwell dishes (MarTek, Ashland, MA). A micropipette filled with 100 µM cAMP was positioned to stimulate cells by using a micromanipulator (Eppendorf, Hamburg, Germany), and the response and movement of cells were recorded by using Metamorph software (Universal Imaging; 1 image per every 5 s). Cell movement was examined by tracing the movement of a single cell in a stack of images.
Subcellular Localization and Fluorescent Microscopy
Cells were pulsed as normal and plated onto glass-bottomed microwell dishes. Cells were stimulated with 100 µM cAMP either using a micropipette or globally with an Eppendorf pipetman. YFP localization and translocation were recorded by using Metamorph software (Universal Imaging; 1 image per every 3 s) and a YFP emission filter. Quantitation of YFP signal was performed using line scan of lines transecting the cell body for each time point before and after stimulation.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), some have been shown to affect WASP–WIP interaction (Stewart et al., 1999). A previous study identified an aromatic triad of residues within the EVH1 domain of Mena necessary for its interaction with a ligand containing the consensus binding sequence FPPPP (Prehoda et al., 1999). The WH1 domain of N-WASP has been shown to bind a proline-rich motif (DLPPPEP) of WIP similar to that recognized by the EVH1 domain of Mena (Volkman et al., 2002). Sequence alignment of the WH1 domain of Dictyostelium WASP with other EVH1/WH1-containing proteins revealed that the triad of residues is conserved (Figure 1A). To investigate the role of the WH1 domain in the function of WASP, we chose to mutate the three aromatic residues that correspond to the aromatic triad: tyrosine 33, tryptophan 39, and phenylalanine 91 (Figure 1A) to alanine. A fourth residue, phenylalanine 90, was mutated because of its proximity to F91. A construct for full-length WASP carrying all four mutations (WASPYWFF) was made. This construct and wild-type WASP were expressed in WASPTK cells. This strain has been shown to express very low levels of WASP and exhibits strong chemotaxis defects (Myers et al., 2005), although random movement appears not to be significantly affected (Table 1). Western analysis revealed equivalent expression of each protein, and a lack of degradation indicated that the aromatic residue mutations do not affect protein stability (unpublished data). Expression of either wild-type or mutant WASP rescued the ability of cells to polarize in response to cAMP pulsing; however, the WASP mutant exhibited a reduced ability to polymerize F-actin at the leading edge (Figure 1B). Figure 1D shows that the leading edge F-actin content of WASPTK cells expressing wild-type WASP (WTWASP) is threefold greater than that of wild-type cells alone, presumably because of ectopic overexpression of WASP. However, YWFFWASP mutant exhibit less than a twofold increase in F-actin accumulation over wild type. Consistent with this data, YWFFWASP-expressing WASPTK cells exhibited a reduced level of F-actin polymerization in response to global cAMP stimulation, indicating that the aromatic triad residues are required for full WASP function (Figure 1C). Chemotactic parameters such as mean velocity, angular deviation, and chemotactic index were measured. In each measurement, WASPTK cells expressing YWFFWASP exhibited an intermediate level of stimulation between that of WTWASP-expressing cells and WASPTK cells alone, indicating WH1 domain is required for the full function of WASP (Table 1).
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; Vaduva et al., 1997; Ho et al., 2001; Martinez-Quiles et al., 2001).
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We next attempted to identify the region of WIPa that interacts with the WH1 domain. Unlike human WIP, WIPa does not have a prominent WH1-binding domain (Volkman et al., 2002; Zetti and Way, 2002) at its C-terminus. One potential binding region lies in the middle of the protein. This region has preserved the "LPPPP"-binding motif, as well as conserved residues that have been suggested, using modeling data, to form electrostatic interactions with conserved charged residues within the WH1 domain (Volkman et al., 2002). We mutated two of these conserved residues (K93 and S95) to alanine and performed GST pulldown experiments. Figure 2D shows that the mutant WIPa (KSWIPa) exhibits a significant reduction in affinity to the WTWH1 domain. This reduction in binding averaged to 41.7 ± 8.9% of wild type over three independent experiments.
WIPa Is Enriched at the Leading Edge and Membrane Protrusions
We previously showed localization of WASP at the leading edge of chemotaxing cells (Myers et al., 2005). YFP-WIPa expressed in vivo also localized to the leading edge of polarized, chemotaxing cells (Figure 3A). YFP-WIPa colocalizes with CFP-WASP at the leading edge (Figure 3C), although they do not colocalize within intracellular compartments. WIPa is transiently enriched at membrane protrusions, which appears to be an early stage of pseudopod formation (Figure 3B). WIPa enrichment, however, seems to be lost during retraction of these protrusions, indicating that WIPa might be required for the onset of extension or maintenance of protrusion (Figure 3, A and B).
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). The K36AWIPa mutant lost its ability to localize to the leading edge of chemotaxing cells. Overexpression of K36AWIPa did not exhibit a significant increase in either microspike formation or F-actin accumulation (Figure 4, B–D), suggesting that actin binding is also required for WIPa activity.
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). A construct designed to express hairpin WIPa (hp WIPa) RNA was electroporated into AX3 cells to reduce endogenous WIPa expression. RT-PCR analysis revealed significant reduction in the expression of WIPa mRNA, but not the messages of unrelated genes involved in F-actin regulation such as WASP, RacC, and VASP (Figure 4A). Northern analysis showed similar results (unpublished data). Phalloidin staining of hp WIPa cells demonstrated moderately reduced F-actin accumulation at the leading edge; however, these cells are more polarized and elongated (average length 18 ± 0.49 µm) compared with wild type (14.83 ± 0.68 µm; Figure 4B). These cells exhibit a lower steady state number of microspikes along the cell body (Figure 4D), indicating that these cells may be defective in their ability to extend lateral membrane protrusions.
WIPa Facilitates F-actin Polymerization via Promoting Elongation of Actin Filaments
Increased level of F-actin and number of microspikes in cells overexpressing WIPa suggest that WIPa stimulates F-actin polymerization. To address more specifically the role of WIPa in actin regulation during membrane extension, the barbed ends of newly formed F-actin were labeled with TRITC-G actin in cells expressing WIPa. Figure 5A shows that wild-type WIPa, but not K36AWIPa, induces an almost twofold increase in leading edge barbed-end staining, suggesting that WIPa stimulates new actin filament formation. To support this data, in vitro F-actin polymerization assays were performed using HSS prepared from cAMP-pulsed Dictyostelium cells as described previously (Zigmond et al., 1997). GTP-bound RacC binds the GTPase-binding domain (GBD) domain of WASP and relieves its autoinhibitory interaction with the WASP acidic (A) domain (Han et al., 2006), presumably allowing WASP to activate the Arp2/3 complex. GTP-bound RacC initiated a roughly threefold stimulation of actin polymerization in wild-type supernatants (AX3), but not in supernatants prepared from WASPTK cells (Figure 5B). Consistent with Zigmond et al. (1997), this small G protein activation of actin polymerization depended on its binding to GTP, as the inactive GDP-bound conformation of RacC failed to stimulate polymerization (AX3+GDPS RacC). Addition of 100 nM recombinant WIPa increased F-actin levels in wild-type supernatant (AX3+WIPa), but not in supernatants of WASPTK cells (WASPTK+WIPa). WIPa knockdown cell supernatants (hp WIPa) exhibited a decrease and delay in the polymerization of F-actin relative to wild-type supernatant. Addition of 100 nM recombinant WIPa to hp WIPa supernatants (hp WIPa+WIPa) rescued this defect (Figure 5B). To visualize assembled F-actin, these experiments were repeated on glass coverslips and viewed with fluorescent microscopy. RacC appears to increase the number of F-actin nucleation sites, presumably via activation of WASP and the Arp2/3 complex (Figure 5C). Addition of recombinant WIPa increases the length of these actin filaments (HSS+RacC + 100 nM WIPa), suggesting that WIPa promotes the elongation of actin filaments. We considered the possibility that WIPa stabilizes F-actin by preventing depolymerization of F-actin. If this were true, addition of WIPa in the presence of phalloidin stabilizing F-actin would be unlikely to lead to a further increase in the length of actin filaments. These reactions were repeated in the presence of 0.4 µM unlabeled phalloidin. The same WIPa-induced increase in the length of actin filaments was observed (unpublished data), suggesting that WIPa's activity does not depend solely on prevention of F-actin depolymerization. WIPa addition to the supernatant in the absence of GTPS-charged RacC does not stimulate filament formation, indicating WIPa alone does not stimulate nucleation of F-actin (unpublished data).
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). Similarly, two phases of WIPa translocation to the cell cortex after cAMP stimulation were observed. The first phase of translocation appeared to be more uniform along the cortex and peaked after 6 s (Figure 6A). In contrast, the smaller second peak of WIPa translocation is less uniform and more localized to sites of membrane protrusions as PHCRAC domain does. WIPa translocation appears to require its WH2 domain, as K36AWIPa showed a significantly lower level of translocation (Figure 6A). This is consistent with the inability of this mutant to localize to the F-actin–enriched leading edge of chemotaxing cells (Figure 3A). Similarly, WIPa expressed in WASPTK cells exhibited a reduced ability to translocate to the cortex. These data suggest that new F-actin polymerization at the cell cortex drives WIPa translocation. To confirm this, we determined the amount of WIPa that cosedimented with the detergent-insoluble F-actin pellet after centrifugation of lysate from cells stimulated with cAMP. Consistently, two peaks of WIPa translocation to F-actin pellet were observed. However, either mutation of the WIPa WH2 domain or WASP deletion significantly reduced the levels of WIPa in the F-actin pellet (Figure 6B).
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WIPa Is Important for the Ability of Cells to Promptly Respond to a New Gradient
To determine the role of WIPa in controlling cell motility, we performed micropipette chemotaxis assays (Chung and Firtel, 1999). Polarized cells were plated on a glass slide and allowed to chemotax toward a cAMP-releasing micropipette for at least 5 min before taking images. Table 1 shows that WIPa overexpression reduces the mean velocity and chemotactic index of cells migrating toward the micropipette, whereas angular deviation is significantly increased, indicating these cells make more directional changes presumably due to the increase of microspike/pseudopod protrusion. These chemotactic defects are not seen in cells overexpressing K36AWIPa (Table 1). Reduction of endogenous WIPa message, however, appears to have little effect on chemotaxis efficiency. Cells expressing the hp WIPa construct are well polarized and exhibit little change in mean velocity and chemotactic index when migrating in a single direction toward a stably localized micropipette (Table 1), suggesting that WIPa may not be essential for the stabilization or maintenance of a mature leading edge.
Because WIPa overexpression caused frequent directional changes, we examined whether WIPa might be responsible for the ability of a cell to sense and respond to changes in the chemoattractant gradient, the micropipette was moved to a new location during chemotaxis. The instantaneous velocity of cells was measured immediately after the pipette was shifted to a location approximately perpendicular to the cell. Figure 7A shows that wild-type cells redirect themselves toward the new chemoattractant source quickly. However, hp WIPa cells exhibit a delayed response to the new direction. Although wild-type cells resume normal velocity in <60 s, hp WIPa cells took more than 300 s to resume normal speed (Figure 7B). We speculated that the delayed response might be due to a delay in making a new pseudopod toward the new directional cue. To test this, we measured the time required for each cell type to initiate pseudopod protrusion, in clear response to the new chemoattractant source, which eventually developed into the new leading edge (Figure 7C). The average time required for new pseudopod formation toward the new cAMP source was 48 s for wild-type cells, whereas hp WIPa cells took 145 s (Figure 7D). These results suggest that WIPa stimulates lateral membrane protrusion and prompt formation of a new pseudopod when cells respond to a change in the direction of a chemoattractant source.
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; Schirenbeck et al., 2006). It is of great interest to determine if both WIPa and VASP are required for microspike formation. We expressed WIPa in a vasp null cell line. F-actin was stained to determine the steady state level of microspikes in each cell type. Consistent with a previous study (Han et al., 2002), vasp null cells showed a significantly lower number of microspikes. Overexpression of VASP without a membrane-targeting myristoylation tag does not significantly increase the number of fliopodia/microspikes per cell. vasp– cells overexpressing WIPa failed to exhibit an increase in the steady state level of microspikes as seen in the wild-type background (Figure 8A). Coexpression of VASP and WIPa in vasp null cells restored increased microspike formation, suggesting that VASP might be required for microspike formation by WIPa.
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), we considered the possibility that VASP may be involved in the translocation of WIPa to the cortex where it could then initiate microspike formation. Figure 8B shows a reduced WIPa translocation in the vasp null background, raising the possibility that a lack of WIPa at the plasma membrane partially explains the lack of microspikes in the vasp null background. VASP has been shown to nucleate and bundle actin filaments in vitro (Schirenbeck et al., 2006), and as we observed a dependence of WIPa translocation upon F-actin, we measured the actin polymerization response of VASP null cells to cAMP. An increase in whole cell levels of F-actin in response to cAMP was diminished in vasp null cells (Figure 8D). Consistently, expression of the F-actin–binding protein ABP in vasp null cells revealed a reduced level of actin accumulation at the cell cortex (Figure 8E). Significantly less WIPa was pelleted with F-actin after cAMP stimulation of vasp null cells, presumably due to a decrease in levels of actin polymerized and/or bundled (Figure 8C). These data suggest that, like WASP, VASP plays a role in the actin polymerization response to global cAMP. Diminished cortical actin levels in these cells lead to a decrease in the translocation of WIPa to the plasma membrane, where it might normally facilitate microspike formation with VASP. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Our data suggests that WIPa stimulates microspike formation and lateral membrane protrusion in a polarized cell. In these processes, WIPa may facilitate elongation of actin filaments already nucleated by WASP's activation of the Arp2/3 complex as shown in our in vitro polymerization assay. It is also possible that WIPa may provide WASP with an additional actin monomer during Arp2/3 complex activation and enhances its rate of filament assembly. The number of WH2 domains in a complex of proteins including the Arp2/3 complex may determine the degree to which the complex enhances actin polymerization. WIP enhances F-actin polymerization of cortactin, which has no WH2 domain (Kinley et al., 2003). Similarly, WASP, which has one WH2 domain and is known to be a better Arp2/3 activator than cortactin, is also stimulated by WIP in a manner that depends on WIP's own WH2 domain (Ramesh et al., 1997). Consistently, the VCA domain of N-WASP, which possesses two WH2 domains, was found to be a better activator of the Arp2/3 complex and microspike formation than VCA domains possessing a single WH2 domain (Yamaguchi et al., 2000).
NMR spectroscopy and computer modeling studies have provided insight into the pathogenesis of WAS by revealing structural details of the WASP (or N-WASP) WH1 interaction with WIP (Rong and Vihinen, 2000; Volkman et al., 2002; Kim et al., 2004). Consistently, we showed that Dictyostelium WIPa binds to the WASP WH1 domain and that the aromatic triad residues are essential for this interaction. We identified a potential WH1-binding region of WIPa that lies within the middle of the protein. This region has preserved a polyproline binding motif, as well as conserved residues that have been suggested to form electrostatic interactions with conserved aspartic acid residues within the WH1 domain (Volkman et al., 2002). Mutation of two of these residues reduced the affinity of WIPa for the WH1 domain. This region, however, is oriented in a C- to N-terminal manner. As proline-rich binding modules like the WH1 domain are known to isoenergetically dock with PPII ligands in two possible orientations due to the twofold rotational pseudosymmetry of the ligand, it is plausible that this region of WIPa interacts with the WH1 domain of WASP.
When cAMP-pulsed Dictyostelium cells are globally stimulated with chemoattractant, two distinct phases of actin polymerization are triggered. Both phases have been shown to correspond to a translocation of an F-actin marker and a PI(3,4,5)P3 marker to the cell cortex (Chen et al., 2003). Cytosolic WIPa also appears to translocate to the cortical membrane in two distinct phases that parallel this actin polymerization. The reduction in translocation caused by the K36A mutation would imply that WIPa translocation is partially mediated by a WH2-F-actin interaction. Interestingly, the second phase of translocation of WIPa is localized to sites of pseudopod/microspike formation. Overexpression of WIPa would result in increased WIPa translocation to the cortex, leading to more F-actin assembly (Figure 6C) and more lateral membrane protrusion and thus lower chemotactic efficiency (Table 1). WIPa translocation to the membrane also appears to require PI3 kinase activity, suggesting that PI(3,4,5)P3 might trigger WIPa translocation. We do not fully understand how this translocation is dependent on PI3 kinase yet. In our previous study, WASP localization was significantly changed upon the inhibition of PI3 kinase by LY294002 (Myers et al., 2005). Sequestration of WIPa by WASP in LY294002-treated cells might result in defective translocation of WIPa to the cortical membrane. We suggest that upon cAMP stimulation, membrane-bound WASP stimulates actin polymerization at the cell cortex presumably via activation by PI(3,4,5)P3 and RacC. The polymerized actin recruits cytosolic WIPa to the cortex, where it facilitates elongation/polymerization of F-actin with WASP, resulting in membrane extension and protrusion. Cell cortical actin accumulation in response to cAMP also appears to require VASP, perhaps via its nucleation or bundling activity (Schirenbeck et al., 2006). A loss of this actin accumulation in VASP null cells appears to lead to a reduction in WIPa translocation similar to that seen in WASPTK cells. The reduction in cortical WIPa may be responsible for the reduced ability of WIPa overexpression to increase the steady state level of microspikes in the absence of VASP.
We have provided evidence that WIPa regulates chemotaxis. Although WIPa overexpression increases microspike formation, decreases speed, and increases angular deviation of a cell toward a chemoattractant source, reduction of WIPa expression decreases microspike formation, slightly improves the chemotactic index, but decreases the ability of cells to reorient toward a new chemoattractant source. The effects of the Dictyostelium formin dDia2 on cell migration are similar (Schirenbeck et al., 2005). Overexpression of dDia2 increases filopodium formation and leads to a decrease in cell motility. Deletion of the dDia2 gene resulted in cells with fewer filopodia but higher cell velocity. Because WIPa stimulates onset of early pseudopod at the plasma membrane, WIPa may allow cells to respond to new gradient promptly by initiating localized bursts of actin polymerization and/or elongation, whereas WASP may provide the main driving force of lamellipodia protrusion by stimulation of F-actin polymerization at the leading edge. The severe chemotactic defects exhibited by the WH1 domain WASP mutant and the lack of significant defects of WIPa knockdown cells during chemotaxis toward a single direction would imply that another polyproline-containing protein might participate in the stabilization or maintenance of WASP-generated actin assembly during lamellipodia protrusion at the leading edge. We have shown in vitro that WIPa increases the length of RacC-induced, WASP-dependent actin filaments and, in vivo, WIPa localizes to sites of newly forming pseudopodia on the cortical membrane. When a chemotaxing cell senses a change in direction in cAMP input, a localized increase in WIPa activity might provide the cytoskeletal framework for the formation of a new pseudopod and, eventually, a new leading edge by facilitating actin polymerization. A decrease in WIPa expression, then, would result in cells with a reduced ability to change direction promptly upon change of chemoattractant gradient due to delayed formation of pseudopodia/microspikes. It is likely that chemotaxis of these cells already committed toward a single chemoattractant source would not be altered or even faster due to less angular deviation. An increase in WIPa expression would produce more lateral pseudopodia resulting in a corresponding increase in the number of turns, and thus a greater angular deviation. A role for both WIP and WASP in Dictyostelium chemotaxis is consistent with the severe impairment of T-cell homing to spleen and lymph nodes from WASP–/–WIP–/– double knockout mice (Gallego et al., 2006).
A species-specific difference, however, between the importance of the WASP family of proteins may exist. Although lamellipodia and filopodia formation appear to be more dependent on SCAR in Drosophila BG2 cells (Biyasheva et al., 2004) and WAVE2 in mammalian cells (Yan et al., 2003), WASP deficiency appears to have a greater effect on membrane protrusion and actin assembly than SCAR deficiency in Dictyostelium. SCAR deficiency does lead to decreased actin assembly at the leading edge (Bear et al., 1998), but these cells polarize well and form visible streams, unlike WASPTK cells, suggesting the possibility of rather normal chemotactic movement despite abnormal F-actin organization. It has been suggested that SCAR is not required for the generation of actin protrusions, but may be more involved in extending and reshaping them (Blagg et al., 2004). This would imply the evolutionary acquisition of WASP-dependent roles of actin assembly during chemotaxis by WAVE/SCAR family members sometime after the divergence of the protozoans. Alternatively, it is conceivable that WASP function is more essential in highly motile cells such as Dictyostelium, neutrophils, and macrophage to provide highly dynamic regulation of the actin cytoskeleton than in slower moving cells like fibroblasts. It has been demonstrated that the VCA domain of SCAR shows different potency in stimulating Arp2/3 than that of WASP or N-WASP, at least in in vitro systems. The VCA domain of N-WASP and WASP induce rapid actin polymerization, whereas SCAR1 induces the slowest rate of nucleation (Zalevsky et al., 2001). It is conceivable to expect WASP or N-WASP to be the primary regulators of dynamic F-actin assembly. Thus, lack of WASP function might cause more severe phenotypes in highly motile cells.
In conclusion, WIPa translocates from the cytoplasm to the plasma membrane upon cAMP stimulation. The regulation of the F-actin cytoskeleton by WIPa at sites of membrane protrusion and pseudopod formation appears necessary to allow chemotaxing cells the flexibility of responding to new sources of chemoattractant and to prevent premature commitment to a single chemoattractant source.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Chang Y. Chung (Chang.Chung{at}Vanderbilt.edu)
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). WIPa, 100 nM, was added 15–30 s after RacC addition where indicated. Reactions were stopped at the indicated time points with actin buffer containing TRITC-phalloidin, and F-actin was pelleted by centrifugation. The amount of F-actin at time 0 was standardized as 100%. (C) Visualization of polymerized F-actin. Supernatants were incubated with or without 100 nM GTP
5 min. Time 0 indicates the first frame after which the micropipette was moved to its new position. Time-lapsed images were taken at 5 s intervals for a total of 