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Previous Article | Next Article 
Molecular and Cellular Biology, June 2006, p. 4041-4051, Vol. 26, No. 11
0270-7306/06/$08.00+0 doi:10.1128/MCB.01868-05
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Cell Biology Laboratory, Department of Biochemistry, BioSciences Institute, National University of Ireland, Cork, Ireland,1 Gallo Research Center, Department of Neurology, University of California, San Francisco, Emeryville, California 946082
Received 23 September 2005/ Returned for modification 10 November 2005/ Accepted 17 March 2006
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ABSTRACT |
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INTRODUCTION |
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Signaling cross talk between the IGF-IR and integrins can either enhance or regulate IGF-IR signaling in different tissues. In smooth muscle cells, IGF-I stimulation and integrin ligation facilitate recruitment of the phosphatase Shp2 via the Dok-1 scaffolding protein to limit IGF-IR signaling (2, 8). In transformed cells, integrin signaling cooperates with growth factor signaling to promote cell survival and migration (11, 23, 31, 42).
RACK1 is a scaffolding protein, which can mediate this cooperation between the IGF-IR and integrins. RACK1 has the ability to recruit signaling molecules to discrete cellular locations via protein-protein interactions at its seven WD repeats, and it has been shown to be involved in different cellular events at the cell membrane, ribosome, and nucleus (25, 32, 36). It is recruited into a complex with the IGF-IR and ß1 integrin (14, 18, 19) and can also promote integrin-mediated cell migration (5). RACK1 also regulates activation of the Ras pathway through its ability to sequester and control availability of Src (6, 7, 16).
Recently, we demonstrated that a mutant of the IGF-IR, Y1250/1251F, which is deficient in promoting cell survival (29) and anchorage-independent growth (15), does not associate with RACK1, and cells expressing this mutant are effectively deficient in adhesion signaling and cell migration (18). Lack of association of RACK1 with the Y1250/1251F mutant IGF-IR was accompanied by lack of association of Shc and Shp2 with RACK1, lack of IGF-I-mediated dissociation of Src from RACK1, and decreased IGF-I-mediated dephosphorylation of focal adhesion kinase (FAK). The characteristics of cells expressing this IGF-IR mutant suggest that integrin signaling is a necessary component of IGF-IR signaling in maintaining the transformed phenotype of these cells. An essential role for ß1 integrin in IGF-I-mediated tumor growth in prostate cancer was also recently proposed (10).
To determine mechanisms of cooperation between IGF-IR and ß1 integrin signaling in tumor cells, we searched for proteins that associate with RACK1 in an IGF-I- and adhesion-dependent manner. From this, we identified the serine threonine phosphatase protein phosphatase 2A (PP2A) as a RACK1-associated protein. PP2A has previously been shown to be activated by ß1 integrin ligation (17) and to negatively regulate activation of Akt in several cell types (2, 17, 40, 41). PP2A has also previously been shown to associate with Shc and to be released upon IGF-I stimulation, thus facilitating Shc phosphorylation (41). The dissociation of PP2A from Shc also requires adhesion (18).
In this study, we investigated whether PP2A contributes to RACK1-mediated regulation of IGF-IR signaling. We found that PP2A dissociates from RACK1 upon IGF-I stimulation and ligation of integrins. The association of RACK1 with PP2A and subsequent dissociation also required that RACK1 and IGF-IR be in one complex. Ligation of integrins with fibronectin or Matrigel was sufficient to cause dissociation of PP2A from RACK1, and this correlated with an IGF-I-mediated decrease in PP2A enzymatic activity. By using recombinant TAT fusion proteins of full-length RACK1, as well as N-terminal and C-terminal deletion proteins, we demonstrated that both ß1 integrin and PP2A associate with the C-terminal domain of RACK1 within WD repeats 4 to 7. Recombinant RACK1 interacted with recombinant PP2A in vitro and did not alter its enzymatic activity. However, RACK1 could restore activity in PP2A immunoprecipitated from IGF-I-stimulated cells. Inhibition of PP2A activity with okadaic acid reversed the increased cell migration caused by overexpression of RACK1, and cells with small interfering RNA (siRNA)-mediated RACK1 knockdown were deficient in cell migration and in IGF-I-mediated decrease in PP2A activity. Overall, the data indicate that the IGF-I-regulated association of PP2A with RACK1 is important for cell migration.
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MATERIALS AND METHODS |
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and anti-RACK1 monoclonal antibodies were obtained from BD Transduction Laboratories (Heidelberg, Germany). The antihemagglutinin (anti-HA) antibody 16B12 was obtained from Berkeley Antibody Company (Berkeley, CA), the antiactin monoclonal antibody was obtained from Sigma Ireland Ltd. (Dublin, Ireland), and the anti-ß1 integrin polyclonal antibody was obtained from Chemicon International (Temecula, CA). Factor Xa protease was purchased from NEB (Beverly, MA). Purified PP2A was purchased from Upstate Biotechnology (Milton Keynes, United Kingdom). MBP-RACK1 and in vitro protein interaction column assay and PP2A assays. Cloning and expression of the maltose-binding protein fused to RACK1 (MBP-RACK1) have been described previously (33). Amylose columns were prepared by loading 0.5 ml of amylase suspension (NEB) onto columns that were preequilibrated with 10 ml of column buffer (20 mM Tris-HCl, pH 7.4, 1 mM EDTA, 20 mM NaCl, 0.002% sodium azide, 10 mM ß-mercaptoethanol). The column was loaded with 800 µl of crude extract from bacterial cultures that expressed MBP or MBP-RACK1. This was allowed to flow through before being washed three times with column buffer (10 ml). A total of 0.5 µg of purified PP2A enzyme (a heterodimer of the 60-kDa [A] subunit and the 36-kDa [C] subunit) was then added to the column in overlay buffer and rotated at room temperature for 45 min. The solution was then allowed to flow through and washed extensively five times with overlay buffer (10 ml) to remove unbound material, before the proteins were eluted with 10 mM maltose in column wash buffer. The MBP or MBP-RACK1 eluate was then subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Western blotting to detect bound PP2A. The eluate was subjected to proteolysis with factor Xa (5 µl), at the protease cleavage site Ile-Glu/Asp-Gly-Arg, to produce the cleaved RACK1 for use in the PP2A assays.
Cell culture, transfection, and IGF-I-mediated stimulation of adherent and nonadherent cells. The MCF-7 breast carcinoma cell line and R– cell line (a mouse embryonic fibroblast cell line derived from the IGF-IR knockout mouse [35]) were maintained in Dulbecco's modified Eagle's medium (DMEM) (Biowhittaker, Verviers, Belgium) supplemented with 10% (vol/vol) fetal bovine serum (FBS), 10 mM L-Glu, and 5 mg/ml penicillin/streptomycin. PC-3 and DU145 cells were maintained in RPMI medium (Biowhittaker, Verviers, Belgium) supplemented with 10% (vol/vol) FBS, 10 mM L-Glu, and 5 mg/ml penicillin/streptomycin. Stable transfectants of MCF-7 HA-RACK1 cells have been previously described (19) and were maintained in DMEM supplemented with 10% FBS and 1 mg/ml G418.
R– cells were transiently transfected with pcDNA3/HA-RACK1, IGF-IR wild type (WT), IGF-IR S1248A, IGF-IR Y1250/1251F, or empty pcDNA3 vector (8 µg of DNA), using Lipofectamine transfection reagent (Life Technologies). After 24 h in culture, the transfected cells were seeded onto 10-cm plates and cultured for an additional 18 h. For analysis of signaling responses in adherent cells, the cells were washed with phosphate-buffered saline (PBS) and starved from serum for 4 h before being stimulated with IGF-I for the indicated times. For analysis of signaling responses in nonadherent cells, confluent MCF-7 cells were detached with trypsin and then washed with PBS. Cells were resuspended in serum-free medium and maintained in suspension for 4 h before stimulation with IGF-I for the indicated times. For ligation of integrins, fibronectin (5 µg/ml) or Matrigel (5%) was added to the nonadherent cultures, followed by incubation in serum-free medium for 4 h and stimulation with IGF-I for the indicated times. Where indicated, okadaic acid (1 nM) was added to serum-starved cells 30 min before stimulation with IGF-I.
Preparation and transduction of TAT-RACK1 proteins.
cDNAs encoding full-length RACK1, amino acids 1 to 317 (TAT-RACK1), the N-terminal fragment of RACK1, amino acids 1 to 180 (RACK1
C), or the C-terminal fragment of RACK1, amino acids 138 to 317 (RACK1N), were cloned into pTAT-HA (13) and expressed in BL21 Escherichia coli cells (Promega, Madison, WI). The recombinant proteins were isolated using Ni-nitrilotriacetic acid agarose resin (Invitrogen, CA) and purified under denaturing conditions using dialysis cassettes and decreasing concentrations of urea-PBS. Transduction of the TAT fusion proteins into MCF-7 cells was achieved by incubating the cells with 1 µM TAT-RACK1, RACK1C, or RACK1N proteins in medium containing 5% serum in DMEM at 37°C for 2 or 4 h, after which cells were prepared for lysis.
Preparation of cellular protein extracts and immunoprecipitation. Cellular protein extracts were prepared by washing cells with PBS and then scraping them into lysis buffer consisting of Tris-HCl, pH 7.4, 150 mM NaCl, 1% NP-40 plus the tyrosine phosphatase inhibitor Na3VO4 (1 mM), and the protease inhibitors phenylmethylsulfonyl fluoride (PMSF) (1 mM), pepstatin (1 mM), and aprotinin (1.5 mg/ml). After incubation at 4°C for 20 min, nuclear and cellular debris were removed by microcentrifugation at 14,000 rpm for 15 min at 4°C.
For immunoprecipitation of endogenous or transfected IGF-IR, ß1 integrin, or HA-tagged proteins, extracts from stimulated or unstimulated cells were initially precleared using bovine serum albumin-coated protein G agarose beads (15 µl beads per 400 µg of total protein in 700-ml lysis buffer) by incubation at 4°C for 1 h with gentle rocking. The lysates were recovered from the beads by centrifugation at 3,000 rpm for 3 min and transferred to fresh tubes for incubation with primary antibody (3 µg of each antibody) for 18 h at 4°C with gentle rocking. Immune complexes were obtained by adding 20 ml of protein G agarose beads for 3 h at 4°C. The beads were washed (three times) with ice-cold lysis buffer, and the immune complexes were then removed from the beads by boiling for 5 min in 20 µl of 2x SDS-PAGE sample buffer for electrophoresis and Western blot analysis.
For RACK1 immunoprecipitations, 700 µg protein was immunoprecipitated with 1 µg anti-RACK1 immunoglobulin M antibodies, 5 µg goat anti-mouse immunoglobulin M Fab fragment, 30 µl protein G agarose beads, and 500 µl immunoprecipitation buffer (1 mM EGTA, 1 mM EDTA, 10 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1% deoxycholate plus the tyrosine phosphatase inhibitor Na3VO4 [1 mM], and the protease inhibitors PMSF [1 mM], pepstatin [1 mM], and aprotinin [1.5 mg/ml]) as described previously (19).
Western blotting. All protein samples for Western blot analysis were resolved by SDS-PAGE on 4 to 20% gradient gels or 12% gels and then transferred to nitrocellulose membranes, which were blocked for 1 h at room temperature in Tris-buffered saline (TBS) containing 0.05% Tween 20 and 5% milk (wt/vol). All primary antibody incubations were performed for 18 h at 4°C. Secondary antibody incubations were carried out at room temperature for 1 h. Where indicated, membranes were stripped by incubation in 62.5 mM Tris-HCl, 1% SDS, and 0.7% 2-mercaptoethanol for 30 min at 50°C, followed by extensive washing in 0.2% and 0.05% TBS-Tween 20. Secondary antibodies conjugated with horseradish peroxidase were used for detection with enhanced chemiluminescence (SuperSignal; Pierce, Rockford, IL) or ECL Plus (Amersham Pharmacia Biotech, Buckinghamshire, United Kingdom) as per the manufacturer's instructions.
Phosphatase assays for PP2A activity. Cellular PP2A activity was measured, using para-nitrophenyl phosphate (p-NPP) as the substrate, with the serine/threonine phosphatase assay kit (Upstate Biotechnology). Cells (R+ or MCF-7) were serum starved for 4 h (or when required, preincubated with okadaic acid [1 nM] for 1 h) prior to stimulation with IGF-I (100 ng/ml) for the indicated times. Cells were lysed in ice-cold lysis buffer (50 mM Tris, pH 7.5, 10% glycerol, 1% NP-40, 137 mM NaCl, 1 µM pepstatin, 1 mM PMSF, 1 µM aprotinin) for 20 min on ice. Nuclear and cellular debris were removed by centrifugation at 14,000 x g at 4°C for 15 min. Clarified supernatants were incubated with anti-PP2A (3 µg) or anti-PP1 (4 µg) antibody for 18 h at 4°C, followed by the addition of 30 µl precoated protein G agarose beads for 3 h. Precoated beads were prepared by incubating beads with 1% (wt/vol) bovine serum albumin in TBS-Triton X-100 for 2 h at room temperature. Immunoprecipitates were washed twice with ice-cold lysis buffer and once with assay buffer (50 mM Tris, pH 7.0, 0.1 mM calcium chloride) by centrifugation at 3,000 rpm for 3 min and resuspended in assay buffer containing 2.5 mM nickel chloride and 900 µg of p-NPP/ml and incubated for 45 min at 37°C. Where indicated, RACK1 cleaved from MBP or MBP alone was added to the PP2A reactions for the duration of the incubation period. The amount of para-nitrophenol produced was determined by measuring the absorbance at 405 nm. Assays were performed with triplicate samples.
Transwell and wound healing migration assays. MCF-7 cells were cultured in 10-cm plates, and fresh medium was added 18 h prior to performing each assay. Cells were serum starved for 4 h, and where indicated, okadaic acid (1 nM) was added at 3.5 h. Cells were trypsinized, washed twice, and then resuspended in serum-free medium. The final cell density was determined using a hemocytometer. The lower wells of the Transwell chamber apparatus were loaded with serum-free DMEM supplemented with 100 ng/ml IGF-I (final concentration) or DMEM supplemented with 10% serum. A 50-µl volume of cell suspension containing 100,000 cells was added to each upper well (10,000 cells in the case of DU145 cells). The loaded chamber was incubated for 16 or 24 h, at which time the chamber was removed from the incubator and disassembled. Cells on the upper surface of the membrane were removed by scraping so that only cells that had migrated through the membrane remained. The membrane was then fixed with methanol, stained with 0.1% crystal violet, and air dried. Cell counts were obtained by counting all cells, and data are presented as an average of counts from five fields of triplicate wells for each test condition.
To assess cell migration, monolayer wound repair assays were carried out by seeding 0.3 x 105 MCF-7 cells/well in multiple wells of a 24-well plate and grown to 70% confluence. The medium was then removed from the wells, and a wound was scored in each well by using a sterile 1-ml pipette tip. The cultures were washed once in DMEM to remove debris and reincubated with DMEM supplemented with 10% FBS. Movement across the wound was examined microscopically, and after 24 h, the cells were stained with Giemsa dye and multiple fields were photographed with a Nikon TE300 inverted microscope equipped with a SPOT digital camera and a magnification of x10.
RNA interference. The siRNAs targeted to human RACK1 were purchased from Ambion. The sequences are as follows: siRNA identification (ID) no. 135615 (siRNA1), 5'-CCAUCAAGCUAUGGAAUACTT-3' (sense) and 5'-GUAUUCCAUAGCUUGAUGGTT-3' (antisense); siRNA ID no. 135616 (siRNA2), 5'-GCUAUGGAAUACCCUGGGUTT-3' (sense) and 5'-ACCCAGGGUAUUCCAUAGCTT-3' (antisense); and siRNA ID no. 135617, 5'-CCUUUACACGCUAGAUGGUTT-3' (sense) and 5'-ACCAUCUAGCGUGUAAAGGTG-3' (antisense). The siRNA negative control used was purchased from Ambion (negative control no. 1, catalog no. 4611). MCF-7 or Du145 cells were seeded at 50% confluence and were transfected with 10 nM or 50 nM oligonucleotide, using the Oligofectamine transfection reagent (Invitrogen). Protein expression was assessed by Western blotting at 24, 48, and 72 h posttransfection. The Transwell migration assays were performed at 48 h posttransfection.
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RESULTS |
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Association of RACK1 with IGF-IR is required for association of RACK1 with PP2A. Since IGF-I rapidly promoted dissociation of PP2A from RACK1 and since RACK1 associates with the IGF-IR (14, 19), we next asked whether the IGF-I-mediated dissociation of PP2A from RACK1 required association of RACK1 with the IGF-IR. To test this, we used two C-terminal mutants of the IGF-IR (serine mutant S1248A and the double tyrosine mutant Y1250/1251F) which do not associate with RACK1 (Fig. 2A) (19). As shown in Fig. 2B, in cells expressing the WT IGF-IR, PP2A was associated with RACK1 in serum-starved cells and dissociated 5 min after IGF-I stimulation. However, in cells expressing either the S1248A or the Y1250/1251F mutants, PP2A was associated with RACK1 in serum-starved cells but did not dissociate in response to IGF-I stimulation. This indicates that association of RACK1 with the IGF-IR is necessary for IGF-I-mediated release of PP2A from RACK1.
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Ligation of integrins is necessary and sufficient for IGF-I-mediated dissociation of PP2A from RACK1. Cell adhesion is necessary for association of RACK1 with the IGF-IR, and we have previously shown that integration of IGF-IR and adhesion signaling are defective in cells expressing the Y1250/1251F mutant IGF-IR (18). Thus, we reasoned that IGF-I-mediated regulation of the RACK1-PP2A complex might be disrupted in nonadherent cells. To investigate this, we measured the association of PP2A with RACK1 in MCF-7 cells stimulated with IGF-I in both adherent and nonadherent cultures. In adherent cells, as expected, PP2A was associated with RACK1 in serum-starved cells but dissociated rapidly in response to IGF-I stimulation. In contrast, in nonadherent MCF-7 cells, PP2A was associated with RACK1 in serum-starved cells but did not dissociate from RACK1 in response to IGF-I stimulation (Fig. 3A). This indicates that adhesion is necessary for IGF-I-mediated dissociation of PP2A from RACK1.
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Next, we asked whether ligation of integrins is sufficient to promote IGF-I-mediated dissociation of PP2A from RACK1. To test this, nonadherent MCF-7 cells were stimulated with IGF-I in the presence or absence of either fibronectin (Fig. 3C) or the synthetic extracellular matrix material Matrigel (Fig. 3D) and were compared to IGF-I-stimulated cells in adherent cultures. In both fibronectin- and Matrigel-treated cells, PP2A was associated with RACK1 under serum-starved conditions, and IGF-I stimulation induced almost complete dissociation of PP2A from RACK1 (Fig. 3C and D). This was in contrast to what was observed in nonadherent cultures without fibronectin or Matrigel, where PP2A remained associated with RACK1 (Fig. 3C and D).
In adherent cells, ß1 integrin was not associated with RACK1 in the absence of IGF-I but was recruited in response to IGF-I (Fig. 3D). In nonadherent cells, ß1 integrin and RACK1 were constitutively associated at a low level, and additional ß1 integrin was not recruited in response to IGF-I, whereas in nonadherent cells treated with Matrigel, ß1 integrin was constitutively associated with ß1 integrin, but more ß1 integrin was recruited in response to IGF-I. Overall, these data demonstrate that ligation of integrins is sufficient to promote IGF-I-mediated dissociation of PP2A from RACK1 and suggest that cell spreading and other signals are not required. The data also indicate that as PP2A is released from RACK1, ß1 integrin is recruited to RACK1 in response to IGF-I.
Binding of both PP2A and ß1 integrin occurs within WD repeats 4 to 7 of RACK1. Our data indicate that ligation of integrins is both necessary and sufficient to induce IGF-I-mediated dissociation of PP2A from RACK, and the kinetics of this dissociation is similar to that for recruitment of ß1 integrin into a complex with RACK1 and the IGF-IR. Based on this, we hypothesized that integrins and PP2A may associate with RACK1 in a mutually exclusive manner in response to IGF-I. To test this, we sought to identify the domains of RACK1 required for association with PP2A and ß1 integrin using recombinant TAT fusion proteins of RACK1 purified from E. coli. These TAT fusion proteins refold when transduced into cells and have been used previously to identify the sites of interaction of RACK1, Fyn, and the NR2B subunit of the N-methyl-D-aspartate receptor (13) and to investigate the dimerization of RACK1 (39).
MCF-7 cells were transduced with TAT fusion proteins encompassing the entire RACK1 protein (WT), WD repeats 1 to 4 of RACK1 (RACK1C), or WD repeats 4 to 7 of RACK1 (RACK1N) (Fig. 4A). Each had the HA epitope tag at the N terminus. RACK1 has been previously shown to form homodimers via the fourth WD repeat (39). Therefore, to check that the Tat fusion proteins were folded correctly, we first demonstrated that endogenous cellular RACK1 could be coprecipitated with each TAT-RACK1 protein (Fig. 4B). Both ß1 integrin and PP2A could be coprecipitated with both the WT TAT-RACK1 and TAT-RACK1N proteins but not with the TAT-RACK1C protein (Fig. 4C). This is in agreement with the results of a previous study which showed that ß1 integrin associates with WD repeats 5 to 7 of RACK1 (22).
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N protein in response to IGF-I stimulation. This showed that for both WT TAT-RACK1 and TAT-RACK1N, as PP2A was released from RACK1, ß1 integrin was recruited in response to IGF-I (Fig. 4D). Taken together, the data support the conclusion that ß1 integrin and PP2A associate with the WD repeat 4-to-7 domain of RACK1 and that there is a mutually exclusive association of PP2A and ß1 integrin with RACK1 in response to IGF-I stimulation. This suggests that integrin ligation may displace PP2A from RACK1.
RACK1 can restore activity to IGF-I-stimulated cellular PP2A in vitro. We next asked whether RACK1 can alter PP2A enzymatic activity. To do this, we incubated either MBP or RACK1 that had been cleaved from MBP-RACK1, using factor Xa, with PP2A that had been immunoprecipitated from either serum-starved or IGF-I-stimulated MCF-7 cells. We also incubated MBP or RACK1 with purified PP2A. RACK1 did not alter the phosphatase activity of PP2A from serum-starved cells and did not alter the activity of purified PP2A. However, in the presence of RACK1, the activity of PP2A immunoprecipitated from cells that had been stimulated with IGF-I was restored to that observed in serum-starved cells (Fig. 5). This indicates that if PP2A is already maximally active, RACK1 does not alter its activity, but if PP2A activity has been reduced, RACK1 can restore it. This suggests that RACK1 has a role in stabilizing or regulating the activity of PP2A in cells and supports the conclusion that PP2A associated with RACK1 is active.
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Taken together, these data indicate that when cells express more RACK1, it preferentially associates with ß1 integrin, and the IGF-I-stimulated complex of the IGF-IR and ß1 integrin is more abundant. This is consistent with the increased migratory capacity and a rapid IGF-I-mediated dephosphorylation of FAK in these cells and suggests that increased RACK1 expression enhances IGF-I-promoted turnover of focal adhesions to facilitate cell migration.
Okadaic acid reverses the ability of RACK1 to enhance MCF-7 cell migration. We next investigated the role of PP2A in the enhanced migratory activity of cells overexpressing RACK1. MCF-7/Neo and MCF-7/RACK1 cells were pretreated with 1 nM okadaic acid for 30 min before we assessed cell motility toward IGF-I in Transwell assays. As can be seen in Fig. 7A, okadaic acid almost completely reversed the increased migration of the MCF-7/RACK1 cells but had no effect on the modest migration or the viability of MCF-7 or Neo cells. This suggests that PP2A activity contributes to the function of RACK1 in enhancing IGF-I-mediated cell motility.
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PP2A activity was higher in serum-starved MCF-7/RACK1 cells than in Neo cells but was still reduced as effectively by IGF-I stimulation as in Neo cells (Fig. 7C). This may explain why okadaic acid has a dramatic effect on cell migration and also suggests that the complex of PP2A with integrin and RACK1 results in increased PP2A activity. RACK1 may also act to sequester PP2A from integrins in nonadherent cells.
RACK1 siRNA ablates IGF-I-mediated regulation of PP2A activity and suppresses cell migration. Our data indicate that RACK1 associated with PP2A acts to promote cell migration in transformed cells. We were interested in determining whether suppression of RACK1 expression would affect IGF-I-mediated regulation of PP2A activity and cell migration. Three siRNA oligonucleotides targeted to RACK1 were tested in MCF-7 and DU145 cells, and all three had the ability to almost completely suppress RACK1 expression. PP2A activity was measured in MCF-7 cells that had been transfected with a control oligonucleotide or two siRNAs directed toward RACK1. PP2A activity was suppressed in response to IGF-I stimulation of control cells but was not reduced in response to IGF-I stimulation of the siRNA-expressing cells that had no detectable RACK1 expression (Fig. 8A). Surprisingly, basal levels of PP2A were higher in the siRNA-expressing cells. These data indicate that although PP2A activity is high, it is not responsive to IGF-I in the absence of RACK1.
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DISCUSSION |
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RACK1 had no effect in vitro on enhancing the activity of purified constitutively active PP2A or PP2A that had been immunoprecipitated from starved MCF-7 cells. However, RACK1 restored the activity of PP2A immunoprecipitated from IGF-I-stimulated cells to levels observed in unstimulated cells. This suggests that although RACK1 cannot directly stimulate further activation of already-active PP2A, it may stabilize or enhance PP2A activity through its scaffolding function. These observations also suggest that PP2A is active when associated with RACK1.
The IGF-I-mediated decrease in cellular PP2A activity requires RACK1 to be associated with the IGF-IR. In all cases where RACK1 was not associated with the IGF-IR, such as in nonadherent cells and in cells expressing the Y1250/1251F mutant, or where RACK1 expression was depleted by siRNA, IGF-I did not suppress PP2A activity. However, total basal cellular PP2A activity in serum-starved cells could not be correlated with RACK1 expression levels or with the association of RACK1 with the IGF-IR. When RACK1 was overexpressed, less PP2A was associated with RACK1 than in control cells, more RACK1 was associated with ß1 integrin, and cellular PP2A activity was increased. In this scenario, PP2A activity was acutely responsive to IGF-I stimulation and rapidly decreased to levels in control cells. When RACK1 was knocked down by siRNA, PP2A levels were constitutively higher than in controls but did not decrease in response to IGF-I. These cells were deficient in cell migration, which suggests that RACK1 is required for the IGF-I-mediated decrease in PP2A activity that is required for cell migration.
It is not immediately obvious why both overexpression of RACK1 and knockdown of RACK1 resulted in increased basal activity of PP2A in serum-starved adherent cells. This may be due to the disruption of the scaffolding functions of RACK1 and an increase or decrease in the targeting or location of other RACK1-associated proteins that may regulate PP2A activity. Another possibility is that integrin function, which has previously been shown to promote activation of PP2A, is deregulated (17). This possibility is supported by our observations that in nonadherent cells and in cells expressing the Y1250/1251F mutant IGF-IR, total cellular PP2A activity was low.
It is likely that components of the RACK1 scaffolding complex with the IGF-IR and ß1 integrin cooperate with PP2A to promote cell migration. We previously found that phosphorylated Shc, Src, IRS-1, IRS-2, and Shp2 can all either transiently or constitutively associate with RACK1 (18). However, although IGF-I and other growth factors have previously been shown to promote dissociation of PP2A from Shc, which facilitates Shc phosphorylation (41), we found that RACK1 can associate only with phosphorylated Shc. This suggests that PP2A is not in the Shc-RACK1 complex at the same time. Protein kinase C alpha (PKC) is also constitutively associated with RACK1 in MCF-7 cells (data not shown). PKC has been shown to promote ß1 integrin-mediated chemotaxis (30) and vascular endothelial growth factor-induced migration of multiple myeloma cells (31). Since PKC is in an active conformation when associated with RACK1 (32) and since PP2A and PKC can physically associate (4), it is possible that PKC may play a role in regulating PP2A activity.
Our data indicate that RACK1 enhances cell migration, in particular chemotaxis toward IGF-I or serum. This is in contrast to the reported suppression of cell migration mediated by RACK1 in CHO cells (5, 9). In these studies, the effects of RACK1 were attributed to PKC activity (5) or RACK1-associated Src (9), but effects on IGF-I-mediated migration were not assessed. The differences observed in our study in transformed cell lines are very likely due to the enhanced IGF-IR signaling in transformed cells, the constitutive association of RACK1 with the IGF-IR, and the formation of the complex with RACK1 and ß1 integrin, which facilitates IGF-I-mediated migration. This is supported by the observation that overexpression of RACK1 increases the migratory capacity of MCF-7 cells toward IGF-I, but not in the absence of IGF-I.
Okadaic acid reverses the increased cell migration due to overexpression of RACK1, which indicates a necessity for PP2A activity in this cell migration. MCF-7 cells overexpressing RACK1 had increased abundance of the IGF-IR-ß1 integrin complex, increased basal cellular PP2A activity, and increased IGF-I-mediated dephosphorylation of focal adhesion kinase. Thus, enhanced RACK1 expression can increase the cooperative signaling between the IGF-IR and integrins that could contribute to cancer cell growth and migration. Indeed, RACK1 expression has been found to be increased in metastatic cancers and in angiogenesis (3).
In summary, we have identified PP2A as a new interacting protein for RACK1 and have shown that RACK1 promotes IGF-I-mediated cell migration through its ability to exclusively associate with ß1 integrin and PP2A. Higher RACK1 expression implies enhanced integrin signaling and confers an increased capacity for cell migration, growth, and invasion. In this way, RACK1 may be an important biomarker for IGF-IR function in cancer cells.
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ACKNOWLEDGMENTS |
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We are grateful to our colleagues in the Cell Biology Laboratory for helpful discussions and to Kurt Tidmore for preparing the illustrations.
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FOOTNOTES |
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REFERENCES |
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