Correspondence to A.E. Oro: oro{at}cmgm.stanford.edu
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Introduction |
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Recent work has shed light on how actin cross-linking proteins contribute to cell shape changes. The initial level of actin organization occurs through actin-nucleating proteins. These proteins assemble actin monomers into a fine meshwork of individual filaments that aid in cell shape changes (Pollard et al., 2001; Welch and Mullins, 2002). However, in vivo, actin filaments rarely exist as isolated single filaments, but instead associate into bundles or networks, in concert with actinbundling/cross-linking proteins at key cellular sites. Numerous studies have documented the wide range in elasticity of filaments with small changes in the concentration of actin-bundling proteins (Pollard et al., 2000; Gardel et al., 2004). Similar studies have shown the need for actin-bundling proteins to achieve mechanical rigidity at the leading edge of migrating cells (Xu et al., 1998; Shin et al., 2004). Loss-of-function studies demonstrate the importance of bundling proteins in processes such as cell migration, epithelial morphogenesis, and axon guidance during development (Mahajan-Miklos and Cooley, 1994; Zheng et al., 2000).
Structurally, actin-bundling proteins are modular proteins that are composed of multiple functional domains (Matsudaira, 1991; Puius et al., 1998; Revenu et al., 2004). Each consists of at least one F-actin binding domain that facilitates actin cross-linking and whose spacing and orientation determine the quality of the bundle formed. In addition to the sequences required for actin cross-linking, each protein also contains "activation" domains distinct from the bundling domain that help regulate the timing and location of bundle formation within the cell. Examples include calcium-binding domains that facilitate calcium-dependent functions (Bretscher and Weber, 1980) and protein interaction domains that allow association with microtubules or portions of the plasma membrane (Matsudaira, 1991; Stock et al., 1999; Tu et al., 2003). The existence of such a modular structure allows the rapid generation of enormous diversity in the actin cytoskeleton from a relatively small number of sequences.
In a screen for novel Shh-responsive genes, we have previously identified Basal cell carcinomaenriched gene 4 (BEG4)/Missing in Metastasis (MIM), hereafter called MIM, as a Shh-responsive gene in the developing hair follicle and in basal cell carcinomas of the skin (Callahan et al., 2004). MIM potentiates Gli-dependent transcription by forming complexes with the Gli transcription factor and the tumor suppressor Suppressor of Fused (Callahan et al., 2004). The previous identification of MIM binding to monomeric actin (Mattila et al., 2003; Woodings et al., 2003) suggests that MIM may be part of a growing family of cytoskeletal regulators that have effects on transcription. To help further understand the role of MIM in morphogenesis, we examined MIM function in actin cytoskeletal remodeling. Here, we show that MIM is a Shh-responsive modular protein that remodels the cytoskeleton by bundling actin filaments. We show that this activity requires self-association, F-actin binding, and an activation domain that associates with RPTP and is required for localizing it to the membrane. Our data suggest a mechanism by which MIM facilitates global and local cytoskeletal patterning events.
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Results |
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To characterize endogenous MIM subcellular localization, we stained ptch1/ fibroblasts with the anti-MIM antibody. MIM accumulates on stress fibers and actin-based structures in the cytoplasm and at the membrane (Fig. 2), but decorates only a subset of stress fibers, as seen by double staining with actin (Fig. 2, AC). In longer cytoplasmic projections, MIM decorates the length of the actin bundles but is excluded from the tips of membrane projections (Fig. 2, DG). The presence of numerous short MIM- and actin-containing structures in peripheral cell areas prompted us to try to determine whether MIM might colocalize with actin bundles at sites of focal adhesions. Consistent with this idea, double staining with markers of focal adhesion complexes such as paxillin, FAK, and phosphotyrosine epitopes confirmed that MIM is localized subjacent to focal adhesion complexes (Fig. 2, HJ). Examination of cells stained with MIM, paxillin, and F-actin demonstrated that MIM decorates actin bundles (Fig. 2 K) attached to focal adhesions (Fig. 2 L). From this data, we conclude that MIM is Shh inducible and localizes to actin bundles underlying focal adhesions.
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Actin filament binding of MIM in the absence of the WH2 domain led us to attempt to determine whether F-actin bundling, an activity seen in other actin-associated proteins (Loomis et al., 2003), could explain the cellular phenotype. Initially, in a low-speed F-actin cosedimentation assay, purified GST-MIM markedly induced actin filament pelleting (Fig. 4 C, lanes 36) compared with actin filaments alone (Fig. 4 C, lanes 1 and 2), GST control (Fig. 4 C, lanes 1922), or in the presence of MIM mutants (Fig. 4 C, lanes 718). Although MIMWH2 pelleted actin efficiently, MIMN277 and MIM
N399 displayed markedly reduced bundling activity in this study. We also found that MIM-dependent actin bundling was inhibited by phosphoinositol diphosphate (PIP2), a hallmark of other actin cross-linking proteins (Stock et al., 1999; Fig. 4 D). Finally, we directly visualized by transmission electron microscopy (TEM) the ultrastructure of the actin bundles induced by MIM (Fig. 4 E). Purified MIM mixed with actin filaments led to the formation of thick actin bundles. Similar bundles were seen with MIM
WH2, which shows that MIM is sufficient to cross-link actin filaments into ordered bundles independently of the WH2 domain. Consistent with the sedimentation assays, MIMN277 and MIM
N399 mutants demonstrated few if any of the bundles seen with full-length MIM (unpublished data). Our results from these biochemical and cell biological assays suggest that MIM bundling activity is responsible for the actin-based projections induced by MIM.
The MIM self-association domain is required for cytoskeletal remodeling
At least two classes of actin cross-linking proteins exist, one that forms bundles through antiparallel homodimers, such as -actinin, and another that cross-links directly through multiple F-actin binding domains on the same molecule (Matsudaira, 1991). The presence of a conserved coiled-coil domain (Fig. 5, A and B), used by actin cross-linking proteins such as the plakins (Fontao et al., 2001), suggests that MIM might fit into the former category. Based on domain analysis programs (Lupas et al., 1991), the predicted coiled-coil domain lies between amino acid residues 100 and 160 and results in a surface of hydrophobic residues opposed by a surface of highly charged residues (Fig. 5 B). We examined MIM self-association using GST pull-down assays from lysates of 293T cells transfected with myc-tagged MIM constructs (Fig. 5 C). Indeed, MIM associated with itself, supporting the homodimer model. The NH2-terminal fragment MIMN277 was sufficient to bind to full-length MIM, indicating that it contained the self-association domain. Moreover, MIM
N159, a mutant lacking the coiled-coil region, failed to bind to GST-MIMN277 or wild-type MIM (Fig. 5 C). These results were confirmed genetically using a GAL4-based yeast two-hybrid interaction assay (Fig. 5 D). Cells coexpressing a MIM bait plasmid and a prey plasmid containing either MIM or MIMN277 grew on selective media, whereas cells expressing MIM plus MIM
N159 or vector alone did not grow. These results confirm and extend previous findings (Yamagishi et al., 2004) using NH2-terminal peptides and strongly argue for a specific self-association through the coiled-coil domain.
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The presence of endogenous MIM in actin bundles supporting focal adhesions in ptch/ MEFs suggests that targeting to these structures may be required for activation. To determine whether relocalization to lipid-rich areas could restore MIM activity, we fused a GAP43 membrane localization domain to the COOH terminus of MIMN408 (Fig. 6 D). The GAP43 localization domain from neuromodulin localizes proteins to cholesterol-enriched focal adhesions at the tips of cytoplasmic projections (Laux et al., 2000). Cells transfected with GFP-MIMN408-GAP43 showed increased staining on the plasma membrane and dramatically increased cell projections (Fig. 6, C and D). This mutant also increased ruffle formation, but had only a minimal effect on stress fiber reduction or microspikes, locations not targeted by the GAP43 tag. This effect was not caused by GFP overexpression in membrane compartments, as the YFP-GAP43 control gave no detectable phenotype (Fig. 6, C and D). These data support the notion that MIM's presence at lipid-rich membrane areas is necessary, in addition to self-association and F-actin binding, for generating membrane projections.
MIM binds to RPTP and relocalizes it to the membrane
To further define the components that activate MIM, we searched for candidates that associate with the MIM activation domain. The MIM COOH terminus has previously been shown to interact with RPTP (Woodings et al., 2003). Cell and developmental studies with RPTP
and other Type IIa RPTPs indicate that they assemble a signaling complex at focal adhesions and are crucial in correctly organizing the cytoskeleton (for review see Johnson and Van Vactor, 2003). Initially, we examined the significance of tyrosine phosphatase activity for MIM function. We treated ptch/ cells with the phosphatase inhibitor orthovanadate (Heffetz et al., 1990) and examined MIM localization and activity. Treated cells exhibited a dramatic reduction in MIM-associated actin cables and cytoplasmic projections (Fig. 7 A), which is consistent with a role for phosphatase activity in MIM function.
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We assayed to determine whether the RPTP D2MIM interaction is required for cytoskeletal remodeling. If binding to the D2 domain at the membrane were required for MIM activity, then a soluble D2 peptide should prevent RPTP-mediated MIM membrane activation. In fact, coexpression of soluble D2 with GFP-MIM inhibited the cell extension phenotype in C3H10T1/2 cells (Fig. 7 C). This inhibition occurred in 70% of the cases. In these doubly transfected cells, MIM was distributed uniformly throughout the cytosol, similar to the
N159 or
N399 mutants. This suggests that interaction with the membrane-associated RPTP-D2 domain is required for MIM activation.
Although the RPTPMIM interaction activates MIM-dependent cytoskeletal remodeling at the membrane, it also appears to be required for the subcellular localization of RPTP
to the membrane. Available anti-RPTP
antisera could detect endogenous expression by Western blot analysis but not by cell staining, so we examined RPTP distribution using transfected protein. Staining of expressed RPTP
in the absence of MIM using a monoclonal anti-RPTP
antibody (Pulido et al., 1995) revealed a dotlike pattern as well as cytosolic staining in C3H10T1/2 cells (Fig. 8). In the presence of MIM, the two proteins colocalized and RPTP
distribution was dramatically enhanced at the membrane and at sites of cytoplasmic projections. MIM did not alter the localization of, or colocalize at the membrane with, RPTP
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D2, a mutant lacking the D2 domain, supporting the in vivo specificity of the interaction with RPTP
. Because of the effects on the subcellular distribution of MIM in vanadate-treated cells, we attempted to determine whether the phosphatase activity of RPTP
was required for its relocalization. RPTP
containing a cysteine-to-serine mutation in the catalytic domain functioned similarly to the wild-type protein, suggesting that such activity is not required for MIM-induced relocalization.
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Discussion |
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The data reported here point out a crucial role for the coiled-coil domain in MIM-dependent bundling activity. In vitro, the dimerization domain aligns two actin filament binding domains to allow bundling to occur, just as it does in -actinin and other bundling proteins. The biochemical and genetic data presented in this work with full-length MIM, in conjunction with previous biochemical data using the MIM NH2 terminus (Yamagishi et al., 2004), suggest that there is a specific interaction between the MIM dimer and actin filaments, although the exact stoichiometry, affinity, and orientation of binding of the protein on the filament will require more careful biophysical studies. However, the importance of this domain is illustrated by the observation that having the activation domain without the coiled-coil domain (MIM
N399 or MIM
N159) is not sufficient for membrane association, strong bundling, or RPTP relocalization. This suggests that in the cell, recognition of MIM by RPTPs at the membrane requires a three-dimensional surface provided by the alignment of the dimerization domain. Interestingly, a search of GenBank sequences reveals two other proteins that have related dimerization sequences, the recently identified ABBA (Yamagishi et al., 2004) and IRSp53 (Miki et al., 2000; Nakagawa et al., 2003), which share 90% and 25% identity, respectively. The similarity between MIM and these dimerization domains suggests that MIM may form heterodimers with other family members, much like members of the plakin or ezrin/radixin/moesin subfamilies of cytoskeletal regulators. Preliminary data suggest that MIM can form heterodimers (unpublished data) with ABBA, which points to additional diversity in the ability to generate cytoplasmic projections.
Our data suggest that MIM belongs to a growing family of cytoskeletal regulators that have transcriptional effects. Previously reported data indicate that MIM forms a cytoplasmic complex with Suppressor of Fused and the transcription factor Gli to regulate transcription (Callahan et al., 2004). This nuclear effect is in direct contrast to the cytoplasmic and membrane effects of actin bundling shown here. Because of recent data suggesting a role for actin binding in transcription (Olave et al., 2002), we considered the possibility that transcription was dependent on the MIM bundling domain. However, we observed that MIM potentiates transcription even without the self-association or WH2 domains that are required for actin bundling or monomeric actin binding. This supports the idea that actin bundling and transcriptional potentiation are mediated through distinct domains. Other proteins have been identified and suggested to regulate the cytoskeleton and transcription, including the Wnt pathway regulators ß-catenin and plakoglobin (Moon et al., 2002; Maeda et al., 2004). Interestingly, the identification of separable domains differs from other regulators such as ß-catenin that use the same domain (armadillo repeats 38) to bind to either adherens junctions or to TCF transcription factors (Rubinfeld et al., 1993; Su et al., 1993; Hulsken et al., 1994; Sadot et al., 1998).
Another aspect of the modular nature of MIM is the identification of distinct sequences outside the actin bundling domain that regulate bundling activity at sites of cytoplasmic projections. Colocalization studies, together with binding and cell biological experiments with a blocking polypeptide, support an important interaction domain between the RPTP D2 domain and MIM amino acids 408538 (Figs. 79). RPTPs are known to assemble into large complexes of proteins that regulate the subjacent cytoskeleton during retinal and motor neuron axon pathfinding (for review see Johnson and Van Vactor, 2003). Recent data indicate that some associated proteins function to localize RPTPs to focal adhesions and neuronal synapses. For example, liprin binds to the D2 domain of another type IIa RPTP, LAR, and is required for LAR function at the synapse, in part by localizing LAR to the synapse (Serra-Pages et al., 1995, 1998; Kaufmann et al., 2002). Our data suggest a similar function for the activation domain of MIM on RPTP to assemble both at the membrane into specialized membrane domains. Future experiments will address whether liprin and MIM are part of the same complex and direct the RPTPs to similar or different compartments at the membrane.
The activation domain of MIM greatly enhances MIM cytoskeletal remodeling in vivo through interaction with RPTP. Because the cross-linking activity of many bundling proteins is activated by dephosphorylation (Zhai et al., 2001), it is tempting to speculate that MIM activity could be controlled via a competition between tyrosine phosphatases and tyrosine kinases, such as Abl or Src. This is consistent with the known association of Abl kinase with Type IIa RPTPs (Wills et al., 1999). Supporting this idea is the strong effect of phosphatase inhibitors on MIM localization and cytoskeletal activity. However, the fact that MIM408-GAP43 rescues much of the cytoskeletal phenotype by localizing MIM to focal adhesions (Fig. 6) suggests that RPTP may be playing a localizing, rather than a catalytic, role with MIM. This is supported by the ability of MIM to localize a catalytically dead RPTP to the membrane (Fig. 8) and our observation that the apparent size of MIM protein does not change in vanadate-treated cells (unpublished data). Similar results have been seen with the fly LAR protein, in which catalytically inactive LAR can rescue LAR-null animals (Krueger et al., 2003). We speculate that modification of non-RPTP accessory proteins may be required to activate MIM-dependent actin bundling activity at the membrane.
Our data provide a framework for how actin bundling proteins like MIM may coordinate effects of both global and local signaling pathways on the cytoskeleton during development. Morphogens such as Shh induce cytoskeletal regulators such as MIM and then rely on MIM's interaction with RPTPs to localize actin bundles. Interestingly, in the neural tube, MIM localizes to Shh-dependent and Islet-1positive motor neurons, which have been shown to express RPTP in rats (Sommer et al., 1997). This suggests that Shh signaling and RPTP may cooperate to control motor neuron morphogenesis through MIM during spinal cord development. Future studies to examine how the activation domain of MIM regulates precise cytoskeletal changes in vivo will enhance our understanding of how morphogens such as Shh control organogenesis.
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Materials and methods |
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F-actin binding coprecipitation assays
High-speed (155,000 g) cosedimentation assays were performed according to the manufacturer's instructions (Cytoskeleton, Inc.). In brief, polymerized actin was incubated with GST-MIM recombinant proteins, BSA, or -actinin. Aliquots of pellet and supernatant were run on SDS-PAGE gels and stained with Coomassie blue. For the F-actin binding curve, we incubated a fixed amount (1 µg) of purified GST-MIM with increasing amounts (05 µM) of polymerized actin. Fractions of inputs (F-actin added) and pellets (GST-MIM bound to F-actin) were subjected to SDS-PAGE and blotted with our purified rabbit anti-MIM antibody. Bands were quantified by densitometry. For low-speed assays, 5 µM F-actin (diluted in G buffer from 23-µM stock) was subjected to centrifugation for 1 h at RT at 10,000 g. After 1 h of incubation with GST-MIM recombinant proteins, samples were spun for 1 h at 10,000 g. Aliquots of pellet and supernatant were run on SDS-PAGE gels and stained with Coomassie blue.
Electron microscopy
F-actin was polymerized as described in the previous paragraph in polymerization buffer at 23 µM and further diluted into the same buffer to 1 µM. Filaments were mixed with the recombinant proteins at ratios of 1:60 to 1:240 (actin: 6XHis-MIM proteins) and adsorbed onto glow-discharged carbon-coated copper grids for 30 s. The grid was washed with two drops of water before being stained with 1% uranyl acetate for 15 s. Electron micrographs were taken in a transmission electron microscope (model JEM-1230; JEOL) at 120 kV.
Antibody generation
Anti-MIM antibodies were generated by injecting purified GST-MIM amino acids 1277 into rabbits. On day 224, sera were affinity purified over a MIM column, generated by covalently attaching purified 6XHIS-MIM 1277 with the Amino Link kit (Pierce Chemical Co.).
Immunocytochemistry
C3H10T1/2, Neuro-2a, and PC12 cells were cultured as indicated by American Type Culture Collection and transfected with FuGENE (Roche). ptch MEFs were maintained as described previously (Taipale et al., 2000) and were serum-starved overnight before processing. Neuro-2a cells were grown on collagen 1coated chamber slides and in 10% serum. Cells were fixed and permeabilized in 4% PFA and 1% Triton X-100 before primary antibody application. For staining of RPTP, cells were fixed for 10 min in 4% PFA, and then washed twice in PBS and permeabilized in 0.5% Triton X-100. Immunoreactivity was visualized with anti-GFP Alexa Fluor 488 and Alexa Fluor secondary antibodies (Molecular Bioprobes). Paxillin and FAK monoclonal antibodies (BD Biosciences), phosphotyrosine monoclonal antibody (Cell Signaling), phalloidin-TRITC (Sigma-Aldrich), anti-RPTP
(Pulido et al., 1995), and monoclonal anti-Myc 9E10 (Sigma-Aldrich) were used as recommended by the manufacturers. Anti-MIM antibody specificity by cell staining was tested in C3H10T1/2 cells transfected with GFP-MIM and immunostained with anti-MIM antibody, preimmune serum or with the anti-MIM antibody that competed with 600 ng of immunizing peptide, followed by a secondary goat antirabbit Alexa Fluor 546 and Hoechst (Molecular Bioprobes) staining. For quantitation of loss of stress fibers, induction of microspikes, or cell projections, cells with projections were defined as either having three or more dendritic cell projections longer than one cell diameter or having many shorter ones with a filopodium appearance. Statistical analysis was performed using GraphPad software. Treatment of ptch/ cells with 10 µM sodium orthovanadate was performed for 9 h. Cells were mounted in Vectashield (Vector Laboratories).
Imaging
Cells in Figs. 13 and 57 were visualized at RT using a confocal laser scanning system (model MRC 1024; Bio-Rad Laboratories) with a Krypton/Argon laser. Lens used was 100x oil NA 1.40 Plan Apo (Nikon). Images were acquired using Lasersharp 2000 and further processed with the Image J software. Neural tube images and Figs. 8 and 9 were collected using a confocal laser scanning microscope (model LSM 510; Carl Zeiss MicroImaging, Inc.) equipped with a Coherent Mira 900 Tunable Ti:Sapphire laser for two-photon excitation. Lasers used were Argon at 458/488/514 nm, HeNe at 543 nm, and Ti:Sapphire at 780 nm. Objectives used were Fluor 20x NA 0.75, Plan Apo 63x NA 1.40, and Plan Apo 100x oil NA 1.40. Images were acquired using the LSM 510 W.S software, version 2.5. All color images were created using Adobe Photoshop 6.0 software.
Neural tube immunofluorescence
Mouse embryos were collected at 11.5 d post coitum, fixed for 7 h in 4% PFA, and processed for paraffin embedding. Deparaffinized sections were blocked for 30 min in 10% sheep serum and 0.1% Tween 20, incubated for 2 h in primary antibody, and then incubated for 45 min in secondary antibody. The antibodies used were rabbit anti-MIM at 1:65 and mouse antiIslet-1 at 1:2 (40.2D6; Developmental Studies Hybridoma Bank). The secondary antibodies used were goat antirabbit Alexa Fluor 488 and goat antimouse Alexa Fluor 546 at 1:250. The last wash included Hoechst at 1:20,000 to stain nuclei.
Online supplemental material
Fig. S1 shows the specificity of the anti-MIM antibody. Fig. S2 more fully characterizes the MIM-induced cytoskeletal changes in C3H10T1/2 cells. Additional methods are also included. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200409078/DC1.
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Acknowledgments |
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The work was supported by National Institutes of Health grants P01AR44012 and AR046786 (to A.E. Oro) and grants from the Secretaria de Estado de Educacion y Universidades cofunded by the European Social Fund (Ministerio de Educacion, Cultura y Deporte) and a Dean's Fellowship (to R. Gonzalez-Quevedo).
Submitted: 14 September 2004
Accepted: 17 December 2004
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