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Trask is a recently described transmembrane substrate of Src kinases whose expression and phosphorylation has been correlated with the biology of some cancers. Little is known about the molecular functions of Trask, although its phosphorylation has been associated with cell adhesion. We have studied the effects of Trask phosphorylation on cell adhesion, integrin activation, clustering, and focal adhesion signaling. The small hairpin RNA (shRNA) knockdown of Trask results in increased cell adhesiveness and a failure to properly inactivate focal adhesion signaling, even in the unanchored state. On the contrary, the experimentally induced phosphorylation of Trask results in the inhibition of cell adhesion and inhibition of focal adhesion signaling. This is mediated through the inhibition of integrin clustering without affecting integrin affinity state or ligand binding activity. Furthermore, Trask signaling and focal adhesion signaling inactivate each other and signal in exclusion with each other, constituting a switch that underlies cell anchorage state. These data provide considerable insight into how Trask functions to regulate cell adhesion and reveal a novel pathway through which Src kinases can oppose integrin-mediated cell adhesion.
Src family kinases (SFKs) are a family of nonreceptor protein tyrosine kinases with a domain structure consisting of a highly conserved kinase domain as well as an SH2 domain and an SH3 domain, a C-terminal negative-regulatory tyrosine residue, and an N-terminal myristoylation site. Three members of the family, Src, Yes, and Fyn, are ubiquitously expressed, while the expression of the other members is largely restricted to specific hematopoietic cell lineages. SFKs participate in numerous cellular pathways in association with growth factor receptors, G protein-coupled receptors, steroid hormones, STAT transcription factors, and integrin receptors (14, 17, 34). The role of SFKs in regulating cell adhesion signaling at sites of adhesion to the extracellular matrix (ECM) is particularly well established. Upon integrin engagement with the ECM and clustering of integrins at sites of cell adhesion to matrix, macromolecular complexes are assembled in association with the intracellular tails of activated integrins (30, 40, 48). Within these focal adhesions, focal adhesion kinase (FAK) is activated by autophosphorylation at tyrosine 397, creating a binding site for the Src SH2 domain (30, 37). Upon binding to FAK, Src is activated and phosphorylates a number of additional tyrosine residues on FAK, creating additional binding sites for SFKs and other proteins. Activated Src also phosphorylates a number of additional cytoskeletal proteins, including paxillin and p130Cas and proteins involved in regulating the RhoA, Rac1, and Cdc42 GTPases (23). These events function to stabilize focal adhesions, generating a force-induced mechanical link with the actin cytoskeleton, and regulate the surrounding membrane dynamics.
SFKs are required for proper establishment of focal adhesions, as fibroblasts deficient in Src kinases have significantly reduced tyrosine phosphorylation at focal contacts and defective cell adhesion to matrix (7, 26, 47). Although this loss-of-function model supports the current molecular models of focal adhesion establishment, the conclusions are not reciprocated by gain-of-function experiments. The constitutively activated v-src oncogene product interacts with focal contacts, phosphorylating target proteins within them (20, 33). However, the activities of the v-src product are destructive to focal adhesions, and in fact, v-src-transformed cells appear to have significantly reduced focal adhesions (11). Therefore, the evidence suggests that SFKs are capable of promoting both adhesive and antiadhesive functions, and most current models reconcile this by proposing that SFKs function in focal adhesion turnover (15). The mechanisms that mediate the antiadhesive functions of SFKs are less well understood. Some evidence suggests that SFKs can mediate focal adhesion disassembly through a RhoA- and mDia1-mediated pathway or through a calpain-mediated pathway (18, 51). In this paper, we describe a novel mechanism by which SFKs can negatively regulate focal adhesion assembly.
We have been studying a novel substrate of SFKs named Trask. Trask is a recently described 140-kDa transmembrane protein with little homology to known families of proteins. It has a large extracellular region containing CUB domains and a smaller intracellular region containing five tyrosines (5). Trask is widely expressed in epithelial cells and tissues as a variable blend of 140-kDa and 85-kDa forms, the latter due to proteolytic cleavage of its distal extracellular region by serine proteases, including the membrane bound MT-SP1 (5, 42). Trask is phosphorylated in vitro by SFKs, including Src and Yes, and is also phosphorylated by SFKs in cells, and its phosphorylation can be inhibited by all classes of SFK inhibitors (5). The phosphorylation of Trask is exclusively dependent on SFKs, since it fails to undergo phosphorylation in Src/Yes/Fyn knockout cells (SYF cells) unless transfected with an SFK member (50). Trask has also been independently identified as a cancer-associated gene by other groups. In a microarray analysis of colon cancers, it was identified as a transcript with increased expression in tumors compared with that in adjacent normal tissues and was named CDCP1 (39). In another line of study, a subtractive immunization screen designed to identify antibodies against more metastatic variants of HEp-3 carcinoma cells identified a surface protein that was named SIMA135, which is identical to Trask/CDCP1 (22).
The suggestion that Trask/CDCP1 is important in cancer progression has been further supported by correlative studies of human tumors, although the data are mixed and the nature of this association and the cellular role of Trask/CDCP1 in cancer is a matter of ongoing interest and investigation. In an extensive analysis of Trask expression and phosphorylation in human tissues, we found that Trask is widely expressed in most epithelial tissues; however, the SFK phosphorylation of Trask is restricted to physiological circumstances of detachment, such as in mitotically detached cells in the colonic crypts (42). However, in a large survey of human tumor sections, we found that Trask is phosphorylated in many epithelial cancers at all stages, including preinvasive cancers such as tubular adenomas, but not in their normal-tissue counterparts (50). In other studies, the elevated expression of Trask/CDCP1 has been linked with poorer prognosis in cancers of the lung, kidney, and pancreas (3, 24, 31) but with better prognosis in endometrioid cancer (28).
The phosphorylation of Trask is linked with cell adhesion such that Trask is phosphorylated almost instantly when epithelial cells detach from matrix and is dephosphorylated when cells readhere to matrix (42). In the current study, we mechanistically studied the link between Trask phosphorylation and cell adhesion through loss-of-function and gain-of-function studies, looking at cell adhesiveness, at the affinity, binding, and clustering of integrins, and at focal adhesion assembly and signaling. We report that the SFK phosphorylation of Trask inhibits cell adhesion through the inhibition of integrin binding activity. This is mediated through the inhibition of clustering, but not regulation of affinity state, and consequent inhibition of focal adhesion assembly and signaling. We also found that the SFK phosphorylation of Trask functions in opposition to and in exclusion with focal adhesion signaling. These two signaling pathways oppose and inactivate each other, defining a switch that regulates cell anchorage state.
Cell lines were obtained from the American Type Culture Collection. To force cells into suspension, cells were washed in phosphate-buffered saline (PBS), exposed to a 2 mM solution of EDTA in Hanks buffer, and when fully detached, cultured in growth media in ULC plates (Corning) for 2 h. ULC plates are not permissive to cell adhesion.
Antiphosphotyrosine antibodies (PY99), anti-FAK, anti-p-Y397 FAK, antipaxillin, and anti-p130Cas antibodies were from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Generation of polyclonal and monoclonal anti-Trask and anti-p-Y743 Trask antibodies were previously described (50). PP1 and PP2 were from EMD-Calbiochem (San Diego, CA). Anti-integrin β1 antibodies were mouse monoclonal anti-integrin β1 (BD Biosciences) for immunoprecipitation and immunoblotting studies and anti-integrin β1 clone P5D2 for integrin-activating and flow cytometry studies (6, 13, 52).
For wound healing experiments, near-confluent MCF10A cells were mechanically scraped across the center of the well to generate a cell-free lane and observed over the next 24 to 48 h for wound closure by cell migration. For migration assays, the Transwell migration assay (Millipore) was used according to manufacturer's procedure. To the bottom wells, Dulbecco's modified Eagle's medium (DMEM)-10% fetal bovine serum (FBS) containing 20 ng/ml epidermal growth factor (EGF) was added. Cells were harvested, resuspended in serum-free medium at densities of 5 × 105 cells/ml, and added to the top well. When necessary, the cells were preincubated with anti-Trask or control antibodies (80 μg/ml) for 30 min at room temperature. After overnight incubation, the cells were stained with crystal violet and enumerated.
Total cellular lysates were harvested in modified radioimmunoprecipitation assay (RIPA) buffer (10 mM Na phosphate [pH 7.2], 150 mM NaCl, 0.1% SDS, 1% NP-40, 1% Na deoxycholate, protease inhibitors, 1 mM sodium orthovanadate). For Western blotting, 50 μg of each lysate was separated by SDS-PAGE, transferred to membrane, and immunoblotted using appropriate primary and secondary antibodies and enhanced chemiluminescence visualization. For immunoprecipitation studies, 300 μg of lysate was incubated overnight with specific antibodies, immune complexes were collected by protein G-Sepharose beads and washed, and the denatured complexes were immunoblotted as described above.
Wells of 96-well plates (not tissue culture treated) were incubated overnight at 4°C with 100 μl of matrix proteins (5 μg/ml fibronectin, 10 μg/ml collagen type IV [Sigma], and 10 μg/ml laminin). MDA-468TR/WT Trask MDA-468TR/pcDNA4-TO cells were induced for 16 h with or without doxycycline (Dox) and were plated at a density of 5 × 105 cells/ml in 100 μl per well. Cells were incubated at 37°C with 5% CO2 for 1 h, followed by three washes with PBS to remove unattached cells. Adhesion was assessed by a colorimetric assay using crystal violet (Sigma), a cytochemical stain that binds to chromatin. Cells were fixed in cold methanol for 15 min, and plates were then air dried and stained with 0.1% crystal violet in PBS for 5 min at room temperature. After removal of the crystal violet solution, the plates were washed extensively and dried, and the stain was released by using 2% SDS in PBS. Stain intensity was quantified by spectrophotometry (at 570 nm) using a plate reader.
Generation of MDA-468 cells expressing doxycycline-inducible Trask was previously described (5). Several clones were analyzed, all of which showed identical adhesion phenotypes when treated with doxycycline. These cells were named MDA-468TR/Trask. Tyrosines 707, 734, and 743 in Trask were mutated to phenylalanine in the pcDNA4-Trask vector by using the Stratagene Quik Exchange kit. The mutated insert was sequenced for confirmation and stably transfected into MDA-468TR cells. Several clones were expanded, and the doxycycline-inducible expression of mutant Trask was confirmed by myc immunoblotting. All clones had similar phenotypes. An individual clone (with tyrosine-to-phenylalanine [YΔF] mutations) was further studied and named MDA-468TR/YΔF Trask.
Small hairpin RNA (shRNA) sequences were cloned into pSico-RGFP and pSico-RNeo vectors. pSico-RGFP expresses green fluorescent protein (GFP) as a selectable marker, and pSico-RNeo contains a neomycin resistance cassette.
To create the shTrask-1 construct, the oligonucleotide 5′-TGAATGTTGCTTTCTCGTGGCAGTTCAAGAGACTGCCA-CGAGAAAGCAACATTTTTTTTGGATCC-3′ was annealed to 5′-TCGAGGATCCAAAAAAAATGTTGCTTTCTCGTGGCAGTCTCTTGAACTGCCACGAGAAAGCAACATTCA-3′ and cloned into HpaI and XhoI sites of the pSicoR vector. To create the shTrask-2 construct, the oligonucleotide 5′-TGATAGATGAGCGGTTTGCAATGCTGATTCAAGAGATCAGCATTGCAAACCGCTCATCTATTTTTTTTGGCGCGCC-3′ was annealed to 5′-TCGAGGCGCGCCAAAAAAAATAGATGAGCGGTTTGCAATGCTGATCTCTTGAATCAGCATTGCAAACCGCTCATCTATCA-3′ and cloned into HpaI and XhoI sites of the vector. For generation of the nonsilencing construct, the oligonucleotides 5′-TGTCTCGCTTGGGCGAGAGTAAGTTCAAGAGACTTACTCTCGCCCAAGCGAGATTTTTTTGGCGCGCC-3′ and 5′-TCGAGGCGCGCCAAAAAAATCTCGCTTGGGCGAGAGTAAGTCTCTTGAACTTACTCTCGCCCAAGCGAGACA-3′ were similarly annealed and cloned. The shRNA constructs were transfected into 293T cells along with the appropriate packaging vectors. The resulting lentiviral particles were used to infect cells. Stably integrated GFP-expressing MCF10A cell infectants were purified by flow sorting, and Trask knockdown was confirmed in the GFP-expressing cell population by Western blotting. MDA-468 cells were infected with pSico-RNeo shRNA constructs and selected with G418 (300 μg/ml).
To measure the integrin signaling in suspended cells in the absence of cell spreading, polystyrene beads (Polysciences) were used. These beads were precoated with 125 μg/ml fibronectin (Sigma) or 50 μg/ml anti-β1 integrin antibody (P5D2) (Santa Cruz Biotechnology) by rotation for 1 h at room temperature in PBS. A total of 106 beads were incubated with 106 cells in a 2-ml suspension in ULC plates for 3 h. At this time, aggregation between beads and cells was observed under a microscope, and the cells remained with rounded morphology. The cell-bead suspensions were pelleted and cells lysed in RIPA buffer.
To measure bead-cell aggregation, fluorescent beads and cells were used. The beads were FluoSphere polystyrene microspheres, which had yellow-green fluorescence (505/515), from Molecular Probes. Beads were precoated with 125 μg/ml fibronectin (Sigma) or anti-β1 integrin antibody (P5D2) at 50 μg/ml (Santa Cruz Biotechnology) by rotation for 1 h at room temperature in PBS. Cells were fluorescently labeled with Vybrant DiD cell-labeling solution (Molecular Probes) according to the manufacturer's instructions. For aggregation, 1 × 106 beads were incubated with 1 × 106 cells in a 2-ml suspension in ULC plates for 3 h. The percent aggregation was determined by flow cytometry.
Cell surface β1 integrin expression was assayed by flow cytometry. MDA-468TR/Trask cells were left uninduced or induced with doxycycline overnight and subsequently stained with anti-β1 integrin antibodies (conformation independent) and secondary fluorescent antibodies and quantified by fluorescence-activated cell sorter (FACS) analysis.
Recombinant human fibronectin fragment 3, containing type III domains 8 to 13 and the amino acid sequence between Glu1266 and Pro1908 (R&D Systems), was fluorescently labeled with the Alexa Fluor 488 microscale protein labeling kit (Invitrogen) according to the manufacturer's instructions. For binding, cells were detached with EDTA, washed with serum-free medium, and incubated with the fluorescent fibronectin fragment for 30 min at room temperature. Cells were washed with serum-free medium, fixed with paraformaldehyde, and immediately analyzed by FACS analysis. Where indicated, samples were incubated and washed in the presence of 2 mM MnCl2. Integrin conformation was also assayed using the conformation state-specific antibody 9EG7 (BD Biosciences) as follows. EDTA-detached cells were washed with 1× Tris-buffered saline (TBS)-5% bovine serum albumin (BSA) and incubated with 9EG7 in the presence of 1 mM calcium or 2 mM manganese in 1× TBS-5% BSA for 1 h on ice. Cells were washed with 1× TBS-1% BSA and consequently incubated with fluorescein isothiocyanate (FITC)-conjugated anti-rat secondary antibodies for 30 min on ice. Cells were washed with 1× TBS, resuspended, and immediately analyzed by FACS.
Tissue cultured cells were viewed and imaged by phase-contrast microscopy using a Nikon TS-100F inverted microscope equipped with a Nikon D100 digital camera attached to the photo port. Images were imported into Photoshop software, converted to grayscale, and gamma adjusted for optimal representation. Fluorophore-stained cells were imaged by fluorescence microscopy using an Axioplan 2 Zeiss microscope at the appropriate excitation wavelengths. Zeiss Fluar objective lenses were used with Immersol 518N as the imaging medium. Pictures were taken at room temperature with an Axiocam MRm camera and Axiovision 4.5 acquisition software, imported into Photoshop software, and gamma adjusted for optimal representation.
Cells were grown on number 1.5 coverslips, and induced with or without Dox for 12 h and fixed with 4% paraformaldehyde. Cells were stained with anti-integrin β1, anti-FAK, or antipaxillin antibodies (Santa Cruz Biotechnology) and secondary Alexa Fluor 546-conjugated antibodies (Invitrogen). Samples were mounted with water, illuminated with a Nikon laser total internal reflection (TIRF) illuminator, and observed under a Nikon TE2000E inverted microscope. Images were taken with the NIS-Elements Advanced Research software and a 100× objective.
To interrogate changes in signaling associated with the anchorage state, we compared adherent epithelial cells with nonadherent (suspended) epithelial cells. The unanchored state can be induced by brief exposure to EDTA and subsequent culture in nonadherent plates. Trask was phosphorylated upon loss of anchorage in MCF10A immortalized epithelial cells (Fig. (Fig.1A).1A). Trask phosphorylation was similarly induced if adhesion is severed by brief exposure to trypsin (Fig. (Fig.1B).1B). Trypsin also cleaves p140Trask to its smaller 85-kDa form. The cleavage is not linked with the state of anchorage or with the phosphorylation of Trask, since EDTA disrupts cell adhesion, inducing Trask phosphorylation without cleavage. The phosphorylation of Trask is tightly linked with the state of anchorage, occurring immediately upon loss of anchorage and continuing for as long as the cells are maintained in suspension and rapidly reversing upon respreading and adhesion (Fig. (Fig.1B).1B). The association between Trask phosphorylation and the loss of adhesion is not unique to these cells and was reproducible in other epithelial cell lines, including immortalized keratinocytes (Fig. (Fig.1C)1C) and epithelial cancer cell lines (Fig. (Fig.1D).1D). Primary keratinocytes also showed no phosphorylation of Trask when cultured in the adherent state but immediately phosphorylated Trask when deprived of anchorage (data not shown). The SFK phosphorylation of Trask in these cells is not an artifactual finding unique to circumstances of tissue culture detachment, but rather it experimentally reproduces physiological conditions of cell detachment in vivo. In human tissue sections, the phosphorylation of Trask is rarely seen, since almost the entire normal epithelium is anchored. However, Trask phosphorylation can be seen in physiologically detached mitotic cells (Fig. (Fig.1E).1E). To look at a physiologic circumstance of experimentally induced adhesion disruption in vivo, we studied skin wounding in mice. Traumatic scalpel-inflicted wounding and disruption of the dermal tissue architecture induced the phosphorylation of Trask in mouse skin to levels comparable to those in detached MCF10A cells (Fig. (Fig.1F).1F). Trask phosphorylation is also frequently seen in human cancers at all stages in vivo (50). Therefore, the SFK phosphorylation of Trask is inversely linked with the state of anchorage in all in vivo and in vitro circumstances that we have observed. Anchorage deprivation in cultured cells provides a highly relevant experimental model system for mechanistic exploration of this physiologic finding, which we have pursued in the following studies.
The loss of anchorage is associated with the phosphorylation of Trask as well as the concomitant dephosphorylation of focal adhesion proteins, consistent with the dismantling of focal adhesions (Fig. (Fig.2A).2A). To study the function of Trask in cell detachment, we generated MCF10A cells lacking Trask expression due to shRNA knockdown (MCF10A/shTrask) along with control cells (MCF10A/shControl) (Fig. (Fig.2B).2B). While control cells fully inactivated integrin signaling in the unanchored state, MCF10A/shTrask cells showed persistent phosphorylation of FAK when detached, indicating a failure to properly inactivate integrin signaling in the unanchored state (Fig. (Fig.2C).2C). The phenotypic consequence of this is evident during the process of cell detachment. Treatment of control MCF10A cells with trypsin resulted in the rapid loss of cell adhesions with consequent cell retraction and rounding (Fig. (Fig.2D).2D). However, Trask knockdown cells (MCF10A/shTrask) were considerably more persistent in adhesion to matrix than were controls and were significantly retarded in their rates of cell detachment, retraction, and rounding (Fig. (Fig.2D).2D). Dissociation from the underlying matrix was impaired in MCF10A/shTrask cells despite the effective disruption of cell-cell associations (Fig. (Fig.2E).2E). With prolonged exposure to trypsin, MCF10A/shTrask cells eventually did go into suspension, but the process was much slower than that for control cells. Similar results were seen with MDA-468 breast cancer cells and when cell detachment was induced by EDTA (discussed below; see Fig. 4D). Similar results were also seen with transient transfection of Trask small interfering RNA (siRNA) (data not shown).
Since proper focal adhesion turnover is important in cell migration, we studied the role of Trask in cell migration in wound healing assays. When a scraping wound was generated in a monolayer of MCF10A cells, cells migrated and eventually filled the wound over a period of 40 h. However, MCF10A/shTrask cells failed to migrate into and close a wound (Fig. (Fig.3).3). This effect is not due to clonal variation, as similar results were seen with transiently transfected shRNA knockdown cells not subjected to subcloning (not shown).
The reduced abilities of Trask knockdown cells to detach from matrix and to effectively inactivate integrin signaling suggest that the SFK phosphorylation of Trask may function to oppose cell adhesion. To further study the hypothesis that the SFK phosphorylation of Trask functions to oppose cell adhesion, we moved to a cell model with which we could conduct both loss-of-function and gain-of-function experiments. For this, we chose MDA-468 breast cancer cells. In contrast to untransformed MCF10A cells, MDA-468 cells have some basal SFK-dependent phosphorylation of Trask, possibly due to the activated state of SFKs in these cancer cells, and the overexpression of Trask in these cells leads to its constitutive phosphorylation at much higher levels, similar to the detached state. For loss-of-function experiments, MDA-468 cells were engineered with near total knockdown of Trask (MDA-468/shTrask) along with nonsilencing controls (MDA-468/shControl) (Fig. (Fig.4A).4A). Forcing MDA-468 cells into suspension led to hyperphosphorylation of Trask (Fig. (Fig.4B)4B) and the concomitant loss of focal adhesion signaling as shown by the dephosphorylation of FAK and paxillin (Fig. (Fig.4C,4C, lanes 2 and 4). However, Trask knockdown cells failed to effectively inactivate focal adhesion signaling, as seen by persistent FAK and paxillin phosphorylation in the suspended state (Fig. (Fig.4C,4C, lanes 6 and 8). This failure to inactivate integrin signaling in detached Trask knockdown cells suggests that the phosphorylated Trask (p-Trask) functions to oppose integrin signaling. Consistent with this, MDA-468/shTrask cells showed increased adhesiveness compared with control cells. This could be seen both during cell detachment and during cell reattachment. When induced to detach by treatment with EDTA, MDA-468/shTrask cells had delayed detachment compared with control cells (Fig. (Fig.4D),4D), and when detached MDA-468/shTrask cells were replated on fibronectin-coated (FC) plates, they had more rapid attachment and spreading than control cells did (Fig. 4E and F).
For gain-of-function experiments, MDA-468 cells were engineered to express the tetracycline (Tet) repressor and to overexpress myc-tagged Trask (MDA-468TR/Trask) or vector control (MDA-468TR/vector) when exposed to Dox (Fig. (Fig.5A).5A). Dox induces the overexpression and constitutive SFK phosphorylation of Trask at high levels (Fig. (Fig.5A),5A), and this serves as a model of Dox-induced SFK phosphorylation of Trask. The phosphorylation of Trask induced by its overexpression is specifically due to SFKs, since it can be inhibited by Src-selective tyrosine kinase inhibitors, and we have previously shown that it fails to undergo phosphorylation in Src/Yes/Fyn knockout (SYF) cells unless an SFK member is cotransfected (50). The constitutive phosphorylation of overexpressed Trask is likely due to the saturation of dephosphorylation mechanisms (not shown). When control MDA-468 cells were seeded onto tissue culture plates, they spread and adhered within several hours of seeding (Fig. (Fig.5B),5B), coincident with the dephosphorylation of Trask seen with the onset of adhesion in these and other cells. However, cells with Dox-induced constitutively phosphorylated Trask failed to spread onto matrix and remained in the suspended state indefinitely (Fig. (Fig.5B,5B, lower-right image). When Trask phosphorylation was induced by doxycycline in already adherent cells, this also resulted in the loss of adhesion to the underlying matrix, although with a longer latency. A disruption of focal adhesions in already adherent cells was evident before the eventual loss of adhesion when observed under TIRF microscopy (discussed further below; see Fig. 11). The antiadhesive phenotype of Trask-overexpressing cells was not due to clonal selection, since 4 different MDA-468TR/Trask clones showed identical biological phenotypes (data not shown). Trask-overexpressing cells failed to adhere, whether they were initially detached with trypsin (Fig. (Fig.5B)5B) or EDTA (Fig. (Fig.5C).5C). The failure to adhere is not due to reduced expression of cellular matrix proteins (Fig. (Fig.5D).5D). The failure to adhere is also not specific to tissue culture-treated plates and was similarly seen with plates coated with fibronectin, laminin, or collagen (Fig. (Fig.5E).5E). This suggests that the inhibition of cell adhesion by phosphorylated Trask is due to the inhibition of integrin function. Consistent with this, when Trask phosphorylation was experimentally induced in adherent cells by doxycycline treatment, FAK and paxillin were dephosphorylated (Fig. (Fig.5F).5F). This led to a rapid reduction in the number of focal adhesions (shown below; see Fig. 11) and eventual loss of cell adhesion. The doxycycline induction of Trask phosphorylation in MDA-468TR/Trask cells also inhibited cell migration in Transwell chamber assays (Fig. (Fig.5G).5G). These findings were not unique to MDA-468 cells. Trask overexpression and constitutive phosphorylation were also studied in HEK293 cells and similarly inhibited cell adhesion and dephosphorylated FAK and paxillin in these cells (data not shown).
The loss-of-function experiments discussed above show that Trask knockdown cells are unable to properly inactivate integrin signaling, and the gain-of-function experiments show that the SFK phosphorylation of Trask inactivates integrin signaling. Taken together, these data show that phosphorylated Trask negatively regulates integrin-mediated cell adhesion. The fact that both loss-of-function and gain-of-function experimental models of Trask impair cell migration (Fig. (Fig.33 and and5G)5G) is consistent with the fact that both adhesive and antiadhesive mechanisms must function for proper focal adhesion turnover in migrating cells.
A limitation of the MDA-468TR/Trask overexpression model is that we cannot specifically attribute the results to the SFK phosphorylation of Trask rather than the overexpression of Trask. Therefore, we generated a full-length Trask construct with tyrosine-to-phenylalanine (YΔF) mutations at Y707, Y734, and Y743. This YΔF Trask mutant correctly localizeed to the membrane but showed no detectable phosphorylation when overexpressed (Fig. (Fig.6A).6A). However, in contrast to the case with wild-type Trask, overexpression of YΔF Trask did not lead to dephosphorylation of the focal adhesion proteins FAK, paxillin, and p130Cas (Fig. (Fig.6B),6B), and it did not inhibit cell adhesion (Fig. (Fig.6C)6C) or cell migration (Fig. (Fig.6D).6D). These data support the conclusion that the antiadhesive effects seen in Dox-induced MDA-468TR/Trask cells are specifically due to the phosphorylation of Trask and not the overexpression of Trask.
The evidence that the SFK phosphorylation of Trask functions in opposition to another SFK target function, outside-in integrin signaling, prompted us to study these seemingly conflicting roles more directly. To do this, we moved to a more direct experimental model of integrin signaling. One of the limitations of studying cell adhesion on a flat surface is that this experimental system involves the concomitant activation of the interdependent but mechanistically distinct processes of cell adhesion (mediated through integrin activation and focal adhesion signaling) and cell spreading (mediated through remodeling of the actin cytoskeleton). In order to more specifically study the functional link between the SFK phosphorylation of Trask and integrin function without the compounding effects of cell spreading, we set up a model of integrin activation using FC beads in contact with cells under conditions of anchorage deprivation. The beads are much smaller than the cells and do not provide a surface for spreading. However, the beads provide sufficient immobilization of fibronectin to cluster and activate integrins, allowing fibronectin-integrin engagement and signaling to be studied in the absence of cell spreading. In this experimental model, integrin outside-in signaling can be assayed biochemically by analysis of p-FAK and p-paxillin, and integrin physical binding activity can be studied by quantitative assays of cell-bead adhesion using flow cytometry. Fibronectin immobilization on beads is necessary, since soluble fibronectin in media is unable to activate outside-in integrin signaling (Fig. (Fig.7A).7A). When uninduced MDA-468TR/Trask cells came into contact with FC beads in the suspended state, Trask was dephosphorylated (Fig. (Fig.7B,7B, lane 2). This effect was reproduced by using beads coated with antibodies that activate integrin β1 (IAbC beads) similar to fibronectin (Fig. (Fig.7B,7B, lane 3). Therefore, the activation of integrins leads to the dephosphorylation of Trask, and the dephosphorylation of Trask is not a consequence of cell spreading.
FC beads or IAbC beads failed to dephosphorylate Trask in cells with Dox-induced constitutive SFK phosphorylation of Trask (Fig. (Fig.7B,7B, lanes 5 and 6); therefore, we looked to determine whether integrin signaling and Trask signaling occur simultaneously or whether they are mutually exclusive. In MDA-468TR/vector cells or in uninduced MDA-468TR/Trask cells, FC beads induced integrin signaling (assayed by p-FAK/p-paxillin) (Fig. (Fig.7C,7C, compare lane 1 with lane 3 and lane 5 with lane 7) concomitant with the dephosphorylation of Trask (7B, compare lane 1 with lane 2). If Trask could not be dephosphorylated (due to doxycycline-induced expression/phosphorylation), then FC beads failed to induce integrin signaling (7C, compare lane 6 with lane 8). This indicates that the two signaling processes oppose each other. Similar results were obtained when integrins were activated by IAbC beads rather than FC beads (Fig. (Fig.7D7D).
This inhibition of integrin signaling is specifically due to the phosphorylation of Trask and not its overexpression, since the doxycycline-induced overexpression of the phosphorylation-defective YΔF Trask mutant failed to block integrin signaling (Fig. (Fig.7C,7C, compare lane 10 with lane 12). In fact, the YΔF Trask mutant itself promoted an exaggerated integrin signaling response. This is due to a partial dominant-negative effect of this mutant, evidenced by the reduction of endogenous Trask phosphorylation upon induction of the YΔF Trask mutant (Fig. (Fig.7E,7E, lane 2). Trask knockdown cells similarly showed an enhancement of integrin signaling. In Trask knockdown cells, baseline integrin signaling was elevated (Fig. (Fig.7F,7F, compare lane 1 with lane 3) and FC beads induced an exaggerated integrin signaling response (7F, lane 4).
Therefore, we found that Trask signaling (i.e., phosphorylation) was inhibited by the experimental induction of integrin signaling. However, we also found that integrin signaling was blocked by the experimental induction of Trask signaling and was enhanced by eliminating Trask signaling. These results were consistent and reproducible across all experiments done either in monolayer adhesion models or in suspended cell-bead models as described above. Taken together, these data reveal that Trask signaling and integrin signaling functionally inactivate each other and signal in a mutually exclusive fashion.
To determine whether phospho-Trask inhibits integrin signaling through a physical interaction with integrin complexes, we looked for the presence of Trask in integrin immune complexes. Trask is indeed seen in β1 integrin immune complexes after Dox induction in MDA-468TR/Trask cells (Fig. (Fig.88 A, lane 2). The interaction is specific for phosphorylated Trask and is not seen in uninduced cells or in cells induced to express the phosphorylation-defective YΔF Trask mutant (Fig. (Fig.8A,8A, lane 5). Immunostaining experiments confirm the expression of Trask at the cell membrane within the adhesion plane (Fig. (Fig.8B).8B). This is consistent with its physical presence within integrin immune complexes and its functional role in disrupting integrin signaling in adherent cells. The Trask-integrin interaction is not an artifact of the overexpression model and is also seen with endogenous Trask when phosphorylated by SFKs in the suspended state (Fig. (Fig.8C,8C, lane 2). The interaction is specific for phosphorylated Trask and is disrupted if Trask is dephosphorylated by brief treatment with a Src inhibitor (Fig. (Fig.8D).8D). Immunostaining experiments confirm that p-Trask and β1 integrin both localize to the membrane in suspended MDA-468 cells (Fig. (Fig.8E).8E). This is consistent with the physical presence of Trask within integrin immune complexes in suspended cells and its functional role in inactivating integrin signaling during anchorage deprivation.
The data discussed above show that p-Trask inhibits outside-in integrin signaling. In these experiments, integrin function was assayed through the phosphorylation of focal adhesion proteins, which occurs downstream of activated integrin receptors. The exact point of inhibition can actually be at several events that occur prior to activation of focal adhesion signaling. p-Trask could inhibit the adoption of high-affinity integrin conformations (typically referred to as inside-out integrin signaling), it could inhibit the engagement of activated integrins with ECM, or it could inhibit the clustering of activated integrins. To more specifically determine the point at which p-Trask interferes with integrin activation and signaling, we performed several additional experiments. We first repeated the FC bead assays with the goal of assaying cell-bead binding activities rather than cellular signaling events. In this experiment, cell-bead binding was quantitatively assayed by two-color flow cytometry using fluorescent cells and fluorescent FC beads (schematically described in Fig. Fig.9A).9A). The loss-of-function and gain-of-function cell models of Trask, along with their controls, were assayed in this experiment. The Dox-induced overexpression and phosphorylation of Trask did not alter the surface expression of β1 integrin (Fig. (Fig.9B);9B); therefore, the cell-bead aggregation data from these experiments reflect the binding activity of integrins and not their surface expression. Relative to the basal state, shRNA knockdown of Trask increased FC bead binding by >2-fold (Fig. (Fig.9C,9C, compare column 1 with columns 2 and 3). Conversely, the Dox-induced SFK phosphorylation of Trask significantly reduced FC bead binding (Fig. (Fig.9,9, compare column 6 with column 7). The suppression of binding activity is due to the phosphorylation of Trask and not its overexpression, since it was not seen with overexpression of the phosphorylation-defective YΔF Trask mutant (Fig. (Fig.9,9, columns 8 and 9). The apparent partial increase in binding activity with the YΔF Trask mutant is consistent with a partial dominant-negative effect of this mutant on Trask signaling (discussed previously). This experiment shows that p-Trask interferes with integrin binding activity.
p-Trask may interfere with integrin binding activity either through the inhibition of integrin conformational activation and ligand binding or the inhibition of integrin clustering. To differentiate between these possibilities, we looked more specifically at the affinity state and ligand binding of integrins and the clustering of integrins. We assayed the affinity state of integrins by two methods. With the first method, we studied cell binding to a labeled monovalent soluble fibronectin fragment. This assay showed that the expression or phosphorylation of Trask had no effect on the fibronectin binding activity of integrins (Fig. 10A). Furthermore, phosphorylated Trask did not block the Mn-induced conformational activation of integrins (Fig. 10A). With a second method, we studied the activation state of β1 integrin by using an activation state-specific antibody. This assay similarly showed that the expression or phosphorylation of Trask had no effect on the conformation of β1 integrin and that phosphorylated Trask did not block the Mn-induced conformational activation of β1 integrin (Fig. 10B). These assays show that p-Trask does not affect the affinity state of integrins, or their ligand binding activity, and taken together with all the previous data, suggest that phosphorylated Trask inhibits cell adhesion through the inhibition of integrin clustering. We looked more directly at integrin clustering and focal adhesion complexes by total internal reflection fluorescence (TIRF) microscopy. The induction of Trask hyperphosphorylation in MDA-468TR/Trask cells led to a marked reduction in integrin clustering and focal adhesion complexes (Fig. (Fig.11).11). Continued Trask hyperphosphorylation eventually leads to the loss of all focal adhesions and loss of cell adhesion, as described earlier. Therefore, p-Trask inhibits cell adhesion by inhibiting the clustering of activated integrins and negatively affecting focal adhesion assembly.
Much is known about the signaling mechanisms that regulate focal adhesions, although at this time we know much more about the mechanisms that promote focal adhesion formation upon integrin activation and clustering and much less about the mechanisms that disrupt them. In particular, SFKs are critically involved in regulating focal adhesion formation and signaling downstream of integrin receptors. Activated Src localizes to focal adhesions, and this is an event that is required for cell spreading on fibronectin (16, 25, 26, 43). Fibroblasts deficient in Src kinases have focal contacts but reduced tyrosine phosphorylation at focal adhesions and defective cell adhesion to matrix (7, 26, 27, 47). Upon integrin clustering, FAK is recruited to the cytoplasmic tails (c-tails) of β-integrins, where it undergoes autophosphorylation at Y397 followed by binding and conformational activation of Src and reciprocal full activation of FAK by Src through phosphorylation of several of its tyrosine residues (30, 35). Src can also be directly activated by its interaction with β integrins (1, 40). The FAK-Src complex phosphorylates a number of substrates, including paxillin and p130Cas, further solidifying and expanding the focal adhesion complex and ultimately linking it with the actin cytoskeleton (4, 10, 21, 38, 44).
In this study, we describe an effector of SFKs that negatively regulates focal adhesion formation, thereby mediating an antiadhesive function. When phosphorylated by SFKs, Trask appears in complexes containing β1 integrin, interfering with integrin clustering and preventing the mechanical and signaling events that link the intracellular cytoskeleton with the ECM. Future studies will focus on determining the molecular mechanisms through which p-Trask prevents integrin clustering. It is now well established that the activities of the integrin heterodimer are regulated by intracellular proteins interacting with the β integrin cytoplasmic tail. In particular, proteins that can induce the separation of the intracellular tails of the α and β integrins promote an extended integrin conformation in the integrin extracellular domains (ECDs) with high affinity for ligand binding (2, 32). The binding of several intracellular proteins, including talin, kindlins, Dok1, tensin, and Numb, is mediated through the integrin NPxY motif (9, 29, 49). While the c-tail interactions described to date have provided great insight into how intracellular proteins can regulate the affinity state of the integrin heterodimer, much less is known about inside-out signaling mechanisms that may regulate integrin clustering. p-Trask does not regulate the affinity state of the integrin heterodimer, as we have shown using assays that measure monovalent ligand binding and integrin conformation. Rather, it inhibits integrin clustering and establishment of focal adhesions. Future studies will seek to more specifically determine how Trask phosphorylation by SFKs inhibits integrin clustering. One possibility is that phosphorylation of Trask by SFKs promotes the SFK phosphorylation of the β integrin c-tail, promoting or disrupting integrin c-tail interactions that are important in clustering. In preliminary attempts thus far, we have not been able to identify p-Trask-induced tyrosine phosphorylation of β1 integrin, but the interactions of the β integrin c-tail are known to be regulated through tyrosine phosphorylation. In particular, the integrin NPxY motif binds some proteins specifically when it is phosphorylated and other proteins preferentially in its unphosphorylated state (29). The tyrosine kinases that phosphorylate these sites have not been well established. v-Src has been shown to phosphorylate the NpxY motif within the β integrin c-tail, disrupting certain interactions and partly explaining the adhesion defects in v-src-transformed cells (36). However, this appears to be one of many promiscuous functions of v-src, and an analogous physiological role for c-Src or other SFKs has not been described. It remains possible that this phosphorylation activity of cellular SFKs is induced specifically in the unanchored state, facilitated by the appearance of p-Trask. The inhibitory effects of p-Trask on integrin clustering and binding activity are almost surely mediated through the phosphorylation of its intracellular tyrosines and not through any functions attributed to its ECD, since Trask truncation mutants lacking the entire ECD inhibit cell adhesion when overexpressed and phosphorylated identically to what is seen with full-length Trask (unpublished data).
Of particular interest is the fact that the inhibitory activities of p-Trask and focal adhesion signaling are reciprocal in nature, as each of these two signaling mechanisms turns off the other, and they function in a mutually exclusive fashion. The SFK phosphorylation of Trask, whether induced physiologically by loss of anchorage or experimentally by Trask overexpression, inactivates focal adhesion signaling. The reverse also holds true, as the activation of integrin signaling, whether induced physiologically by cell adhesion to matrix or experimentally by fibronectin-induced activation of integrin receptors, promotes the dephosphorylation of Trask. It is apparent from all our experimental model systems and from all observed physiological states that Trask phosphorylation and focal adhesion signaling inactivate each other. Therefore, integrin signaling and Trask signaling inhibit each other and constitute mutually exclusive and opposing signaling programs, defining a switch that determines the cell anchorage state. Although there are five tyrosines within the intracellular domain of Trask, in extensive tyrosine mutation studies, we have found that the phosphorylation of Trask occurs as an all-or-none event consistent with its function as a switch (unpublished data). The mechanisms that mediate the dephosphorylation of Trask almost surely involve the regulation of specific protein tyrosine phosphatases (PTPs). It is likely that adhesion promotes the activation of a PTP or the interaction of a PTP with Trask, thereby shifting the phosphorylation state equilibrium toward the dephosphorylation of Trask.
Interest in Trask (also known as CDCP1) has grown because of a growing body of evidence suggesting that it may have cellular functions that are particularly important in tumor progression. A number of studies have documented increased expression of Trask/CDCP1 in some cancers of the lung, kidney, and pancreas, with a poorer prognosis conferred by its elevated expression (3, 24, 31). Some studies have suggested that Trask/CDCP1 promotes anoikis resistance in cancer cells (45). This may be a cell-specific finding, since anoikis resistance was not affected when Trask was knocked down in MDA-468 breast cancer cells (data not shown). Other studies suggest that Trask/CDCP1 may be more important for tumor metastasis. Consistent with this, a number of experimental studies using overexpression, knockdown, or monoclonal antibody targeting approaches show that the functions of Trask/CDCP1 are important in tumor invasion and metastasis (12, 19, 41, 46).
The link between Trask/CDCP1 and cell adhesion has been reported by others as well. Brown et al. reported that Trask/CDCP1 undergoes tyrosine phosphorylation upon loss of adhesion, but they reported that the phosphorylation event is specifically linked with the proteolytic cleavage of the Trask/CDCP1 ECD and does not occur with EDTA-induced detachment (8). However, this study was done without the benefit of anti-Trask/CDCP1 antibodies, and the data were only indirectly inferred from the cross-reactive activities of an anti-p-FAK antibody. Our current data obtained using specific anti-Trask/CDCP1 antibody reagents as well as myc-tagged Trask/CDCP1 constructs now clearly show that the phosphorylation of Trask/CDCP1 is tightly linked with the state of anchorage and is not a consequence of proteolytic cleavage (Fig. 1A, C, D, and F, ,4B,4B, ,5A,5A, and and6A).6A). In panels of the cell types studied, we found that the relative expression levels of the 85- and 140-kDa forms of Trask vary considerably among different cell types but that both forms undergo phosphorylation when anchorage is lost. Both examples were seen in this study. MDA-468 cells express predominantly the 85-kDa form of Trask, which undergoes phosphorylation upon detachment (Fig. (Fig.4B),4B), mouse keratinocytes express predominantly the 140-kDa form of Trask, which undergoes phosphorylation upon scraping (Fig. (Fig.1F),1F), and MCF10A cells express a mixture of the 85- and 140-kDa forms of Trask, both of which undergo phosphorylation upon detachment (Fig. (Fig.1A).1A). The evidence is now clear that the phosphorylation of Trask/CDCP1 is not a consequence of the proteolytic cleavage of its ECD.
It should be noted that overexpressed Trask does not follow the same cleavage ratio as endogenous Trask. Although MDA-468 cells express predominantly the 85-kDa form of Trask, its overexpression by transfection produces both forms. This is likely due to the fact that the overexpression exceeds the cellular capacity for full cleavage. However, both forms are phosphorylated and both forms are found in integrin immune complexes when expressed (Fig. (Fig.8A).8A). Therefore, Trask cleavage does not appear to play a role in the regulation of integrins. The adhesion functions of Trask appear to be entirely mediated through the tyrosine phosphorylation of its intracellular domain.
This work was funded by the National Institutes of Health (grant CA113952 to M.M.M.). D.S.S. is funded by a Susan G. Komen for the Cure Postdoctoral Fellowship. C.H.W. was funded by a California Breast Cancer Research Program Postdoctoral Fellowship.
We thank Michael McManus and the UCSF Sandler Lentiviral RNAi core facility. Data for the TIRF microscopy analysis were acquired at the Nikon Imaging Center at UCSF/QB3.
We have no conflicts of interest to declare.
Published ahead of print on 28 December 2010.