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Department of Microbiology, Mount Sinai School of Medicine, New York, New York 10029,1 Department of Oncology-Lombardi Cancer Center, Georgetown University Medical Center, Washington, D.C. 200072
Received 19 November 2001/ Returned for modification 7 December 2001/ Accepted 27 December 2001
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ABSTRACT |
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INTRODUCTION |
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(17), and suppressor of cytokine signaling 2 (20). However, information regarding the functional roles of many of these molecules in IGF-IR signaling and biological functions is currently limited.
While it has been shown that the protein tyrosine kinase FAK may function as a substrate for IGF-IR (3), its role in mediating integrin signaling is better understood (11). FAK becomes rapidly tyrosine phosphorylated after engagement of integrins with extracellular matrix (ECM) proteins or antibody (Ab)-induced clustering of integrins, leading to the formation of focal adhesion plaques, areas of the plasma membrane which make contact with and connect the actin cytoskeleton to the ECM (11). In addition to mediating cell adhesion with the ECM, integrins also function to regulate the reorganization of the cytoskeleton and the transduction of adhesion-dependent proliferative and survival signals. ECM- and integrin-initiated signals are mediated by focal adhesion protein complexes, which contain actin cytoskeleton-binding proteins, such as 
-actinin, talin, tensin, vinculin, paxillin, and p130CAS and protein kinases such as FAK, protein kinase C (PKC), c-Src, and ß1 integrin-linked kinase (ILK) (30). The activation of pathways mediated by ERK, c-Jun NH2-terminal kinase (JNK), and ILK lead to the activation of transcription factors c-Fos, c-Jun, and ß-catenin/TCF, respectively, and the subsequent induction of cyclin D1 expression and activation of cyclin-dependent kinase (Cdk) 4/6 activity, which are required for progression through the G1 phase of the cell cycle (26). Cellular adhesion results in downregulation of the Cdk inhibitors (CKIs) p21Cip1/WAF1 and p27Kip1 and induction of cyclin E-Cdk2 activity, events necessary for transit through G1 (26). The mechanism of integrin-mediated downregulation of p21Cip1/WAF1 and p27Kip1 is currently unknown. Adhesion-dependent signaling appears to be necessary for the optimal activation of the MAPK pathway by growth factor-dependent mitogenic signals (2). Additionally, growth factors such as IGF-I may cross-regulate integrin-dependent signaling pathways, since recent reports have demonstrated the tyrosine phosphorylation of paxillin and p130CAS and association of Crk with p130CAS after IGF-I stimulation (12, 24), but the significance of this cross-regulation remains unknown.
To identify signaling molecules that are involved in mediating IGF-IR-dependent functions, we used the cytoplasmic domain of IGF-IR as bait in a yeast two-hybrid interaction trap. We identified here an IGF-IR-interacting molecule called RACK1, which was originally named for its ability to bind to activated PKC in vitro (46). Subsequently, RACK1 was shown to interact with the cytoplasmic tail of ß integrins after treatment with phorbol ester (33) and to interact with and inhibit the kinase activity of c-Src when overexpressed in NIH 3T3 cells (13), and this interaction was recently shown to be enhanced by activation of PKC and tyrosine phosphorylation of RACK1 (14). In addition, RACK1 was suggested to be an anchor for Ran1 (Pat1) kinase, a regulator for meiotic development in yeast (36). Very recently, RACK1 was shown to associate with PTPµ at cell-cell contacts (38) and to associate with the alpha interferon receptor and signal transducer and activator of transcription 1 (Stat1) to facilitate its activation (61).
In this study, we demonstrated the IGF-I-inducible intracellular interaction of RACK1 with IGF-IR, PKC, and ß1 integrin. To address the role of RACK1 in IGF-IR-mediated functions, we introduced exogenous full-length RACK1 into an NIH 3T3 cell line that expressed an elevated level of IGF-IR (NIH 3T3-IGFR) and showed that overexpression of RACK1 resulted in the inhibition of IGF-I-dependent cell growth in both anchorage-dependent and anchorage-independent conditions. Overexpression of RACK1 altered the cell morphology of NIH 3T3-IGFR fibroblasts by enhancing the formation of stress fibers and focal adhesions concomitant with increased cell spreading, which were correlated with the increased tyrosine phosphorylation of FAK and paxillin. The major IGF-IR mitogenic signaling pathways mediated by IRS-1, Shc, PI3K, and MAPK were unaffected. In contrast, IGF-I-inducible signaling by ß1 integrin and focal adhesion molecules, including p130CAS, paxillin, and Crk, was perturbed in these cells. RACK1 overexpression, with resultant cell cycle delay in G1 or G1/S, was correlated with increased levels of p21Cip1/WAF1 and p27Kip1 and reduced IGF-I-dependent cyclin E-associated kinase activity. Introduction of antisense oligonucleotides into Swiss 3T3 cells reduced RACK1 expression, inhibited cell spreading, and suppressed IGF-I-dependent monolayer cell proliferation. This report provides initial evidence for a role of RACK1, a novel IGF-IR-interacting molecule, in the regulation of focal adhesion-based cytoskeletal reorganization and IGF-I-dependent cell growth, cell transformation, and integrin-mediated signaling.
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MATERIALS AND METHODS |
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Mammalian expression plasmids. Primers 5'-GCTGGTACCGAGGCACGAGGCGGCGTG-3' (sense [primer 1]), 5'-TGCTCTAGACTTCTAGCGTGTGCCAAT-3' (antisense [primer 2]), 5'-CCCAAGCTTACCATGGACTACCCTTATG-3' (sense primer with hemagglutinin [HA] sequence [primer 3]), and 5'-ACATGCATCTAGGCATAATCTGGCACATCATAAGGGTATGCGCCATCTGCGCCGCGTGTGCCAATGGTCACCTG-3' (antisense primer with HA sequence [primer 4]) and the template pJG4-5-RACK1 were used to generate RACK1 PCR fragments containing an HA epitope sequence at either the 3' end (primer pair 1 and 4) or the 5' end (primer pair 2 and 3). The PCR-generated fragments were cloned into the mammalian expression vector phEF-Neo, a plasmid containing the human elongation factor 1 promoter and the neomycin resistance gene, to generate phEF-HA-RACK1 (5' HA) and phEF-RACK1-HA (3' HA). The phEF-RACK1 plasmid, encoding full-length RACK1, was generated by combining a fragment of phEF-RACK1-HA containing the 5' RACK1 sequence with a fragment of phEF-HA-RACK1 containing the remaining 3' RACK1 sequence, which effectively eliminated the HA sequence. All constructs were verified by dideoxy sequencing. The IGF-IR expression plasmids, pMX-IGFR, and phEF-IGFR have been described previously (34, 68).
Cell culture. Cells were maintained at 37°C and 5% CO2 in a humidified incubator except where noted. Swiss 3T3, NIH 3T3, R2-IGFR, NIH 3T3-IGFR, 3T3-RasV12, rat intestinal epithelial (RIE)-NM1, RIE-T6, and 3Y1-Src cells were grown in Dulbecco’s modified Eagle medium (DMEM; Gibco) supplemented with 5% heat-inactivated bovine calf serum (BCS; Gibco) and routinely used antibiotics. Human embryonic kidney (HEK) 293T epithelial cells were grown in the same conditions as described above except with 10% heat-inactivated fetal bovine serum (FBS; Gibco). Primary chicken embryo fibroblasts (CEF) and virus-infected CEF were grown in DMEM containing 5% heat-inactivated BCS and 1% heat-inactivated chicken serum and antibiotics. NIH 3T3-IGFR and R2-IGFR cells, which stably overexpress IGF-IR, were generated by transfection of NIH 3T3 and Rat2 fibroblasts with pMX-IGFR (encodes the full-length human IGF-IR under the control of the Moloney murine leukemia virus long terminal repeat) by the calcium phosphate method and subsequent selection with neomycin. The generation of RIE-NM1 and RIE-T6 cells, which are rat intestinal epithelial cells stably expressing NM1 and T6, oncogenic gag-human IGF-IR and gag-human IR fusion receptors, respectively, was described previously (40). 3Y1-Src rat fibroblasts, which express v-Src, were generated previously in our laboratory by infection of 3Y1 cells with SR-A Rous sarcoma virus and clonal selection. 3T3-RasV12, an NIH 3T3 cell line expressing an activated H-Ras mutant, was kindly provided by Andrew Chan at Mount Sinai School of Medicine.
DNA transfection and generation of stable NIH 3T3-IGFR/RACK1 cell lines. Transfections were performed by the method of calcium phosphate coprecipitation as described previously (68). HEK293T cells were grown to 50% confluence and transfected with 10 µg of phEF-IGFR alone or cotransfected with 5 µg of phEF-IGFR and 5 µg of phEF-Neo, phEF-RACK1, phEF-HA-RACK1, or phEF-RACK1-HA for 12 to 16 h. Cells were allowed to recover for 24 h in fresh medium before serum starvation and treatment with IGF-I or phorbol 12-myristate 13-acetate (PMA). NIH 3T3-IGFR cells were plated at a density of 5 x 105/10-cm dish and were cotransfected with 10 µg of phEF-RACK1-HA or phEF-Neo and 1 µg of pBabePuro, a murine retrovirus long terminal repeat-based plasmid containing the puromycin resistance gene. Drug-resistant transfectants were selected in medium containing 5 µg of puromycin (Sigma)/ml beginning 24 h after transfection. Individual colonies were isolated and amplified as individual clones, and the remaining colonies were pooled and amplified as a mass culture.
Infections. Stock supernatants containing viruses encoding gag-IGFR (NM1), v-Ros (UR2), and v-Src (SR-A), as well as temperature-sensitive (ts) mutants of gag-IGFR (tsNM1), v-Ros (tsROS), and v-Src (tsSrc), were generated previously in this laboratory. Early passage primary CEF were infected with an aliquot of a given virus stock in the presence of 2 µg of Polybrene/ml. Infected CEF were maintained at 37°C prior to experiments. Cells that were infected with ts mutants were incubated at 41°C (the nonpermissive temperature) for 3 days before being shifted to 35°C (the permissive temperature) for the times indicated in the figure legends. Subcellular fractionation and protein analyses were subsequently performed. Oncogenic transformation, marked by distinct and characteristic morphologic changes of CEF, was typically evident 24 to 48 h after the shift to a permissive temperature.
Morpholino oligonucleotides. Morpholino oligonucleotides of the murine RACK1 sequence were generated by Gene Tools. The sequence of the antisense oligonucleotide is 5'-CACGAAGGGTCATTTGCTCGGTCAT-3' and that of the standard control oligonucleotide is 5'-CCTCTTACCTCAGTTACAATTTATA-3'. The manufacturer's protocol was used to deliver the morpholino oligonucleotides. Briefly, 3 x 105 trypsinized Swiss 3T3 cells were seeded into each well of a six-well culture plate 1 day prior to delivery. On the following day, the medium in each well was replaced with 1 ml of fresh medium. Antisense or control morpholino oligonucleotides were added to a final concentration of 10 µM and swirled for 10 s. Cells were then gently scraped off the surface of the well, pipetted up and down, and transferred to a new culture plate, and this process was repeated 4 h later. Cells were then maintained in culture for subsequent experiments.
Abs.
Anti-RACK1, anti-PKC
, anti-PKCµ, anti-FAK (for immunoblotting [IB]), anti-paxillin, anti-p130CAS, anti-Crk, anti-Shc, anti-phosphotyrosine-horseradish peroxidase (RC20-HRP), anti-mouse immunoglobulin G (IgG)-HRP, and anti-rabbit IgG-HRP Abs were purchased from Transduction Laboratories. Anti-p21Cip1, anti-p27Kip1, anti-Cdk4, anti-Cdk6, anti-Cdk2, anti-cyclin D1, anti-cyclin E, and anti-cyclin A Abs were purchased from Santa Cruz Biotechnology. Anti-phospho-Akt (Ser473), anti-phospho-p70S6 kinase (Ser411), and anti-phospho-ERK1/2 (Thr202/Tyr204) Abs were purchased from New England Biolabs. Anti-p85PI3K and anti-FAK (for immunoprecipitation) Abs were purchased from Upstate Biotechnology, Inc. Anti-ß1 integrin Ab was purchased from Gibco. Anti-IGFR (extracellular domain) Ab was purchased from Calbiochem. Anti-GAPDH (glyceraldehyde-3-phosphate dehydrogenase) Ab was purchased from Biodesign International. Anti-vinculin Ab, phalloidin-fluorescein isothiocyanate (FITC), and phalloidin-tetramethyl rhodamine isothiocyanate (TRITC) were purchased from Sigma. Anti-retinoblastoma (Rb) Ab was purchased from PharMingen. Anti-mouse IgG-FITC and anti-rabbit IgG-FITC Abs were purchased from Boehringer Mannheim. Abs were used according to the manufacturer's recommended dilutions unless otherwise noted. Anti-mouse IgG and anti-mouse IgM secondary Abs for immunoprecipitation were purchased from Accurate. Anti-IGFR (cytoplasmic domain) and anti-IRS-1 Abs were described previously (34, 69) and were used at 1:500 and 1:250 dilutions, respectively.
Stimulation of cells. Cells grown to confluence or after transfection (HEK293T) were incubated in serum-free DMEM. After 24 h, the cells were incubated in serum-free DMEM supplemented with 100 ng of IGF-I (Intergen) per ml, 0.2 µM PMA (Sigma), or 5% BCS for 2, 5, 10, 20, 30, or 50 min as indicated in the figure legends. Cells were then immediately placed on ice. Unstimulated control cells were mock treated with serum-free DMEM for 10 min.
Preparation of cell lysates and immunoprecipitates.
Cells were placed on ice and were washed twice with cold Tris-Glu buffer (25 mM Tris-HCl, pH 7.4; 150 mM NaCl; 5 mM KCl; 1 mM sodium phosphate; 0.1% glucose). For protein analysis, cells were lysed with radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris-HCl, pH 7.4; 150 mM NaCl; 1% deoxycholate; 1% Triton X-100; 5 mM EDTA; 1% aprotinin; 1 mM sodium orthovanadate; 0.4 mM phenylarsine oxide [PAO]) for immunoprecipitation or with Western lysis buffer (10 mM Tris-HCl, pH 7.4; 1% sodium dodecyl sulfate [SDS], 1 mM phenylmethylsulfonylfluoride [PMSF]; 1% aprotinin; 1 mM sodium orthovanadate; 0.4 mM PAO; 25 mM NaF) for direct Western analysis. For coimmunoprecipitation, cells were permeabilized for a minimum of 30 min with one of three solutions, namely, digitonin buffer (1% digitonin [Boehringer Mannheim]; 150 mM NaCl; 20 mM Tris-HCl, pH 7.4; 1 mM PMSF; 1% aprotinin; 1 mM sodium orthovanadate; 0.4 mM PAO; 25 mM NaF; 5 mM EDTA), NP-40 buffer (1 or 0.5% NP-40; 20 mM Tris-HCl, pH 7.4; 150 mM NaCl; 5 mM EDTA; 1 mM PMSF; 1% aprotinin; 1 mM sodium orthovanadate), or CHAPS {3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate} buffer (10 mM CHAPS 
; 50 mM Tris-HCl, pH 8.0; 150 mM NaCl; 2 mM EDTA; 1 mM PMSF; 1% aprotinin; 1 mM sodium orthovanadate; 15 mM NaF; 10 mM sodium pyrophosphate) as indicated in the figure legends. Cell lysates were clarified by centrifugation at 12,000 rpm for 10 min at 4°C. Protein concentration of the supernatants were determined by Bradford assay, and equivalent amounts of protein from cell lysates were used for immunoprecipitation or analyzed directly by SDS-polyacrylamide gel electrophoresis (PAGE) followed by Western blotting. For immunoprecipitations, the indicated Ab was added to lysates and allowed to rock at 4°C for 2 to 4 h (kinase assays) or overnight (others). Then, 1 µg of anti-mouse IgG Ab (for monoclonal primary Abs) or anti-mouse IgM Ab (for anti-RACK1 Ab) and 25 µl of protein A-Sepharose beads (Repligen) were added to the lysates, which were allowed to rock for another 2 h at 4°C. Immunoprecipitates were washed twice by gentle vortexing in fresh cold RIPA buffer (for protein analysis), 0.2% digitonin buffer (for digitonin-permeabilized lysates), or 0.5% NP-40 buffer (for NP-40-permeabilized lysates). Washed immunoprecipitates and total cell lysates were denatured by boiling after the addition of one-sixth of the volume of 6x reducing SDS-PAGE sample buffer (360 mM Tris, pH 6.6; 12% SDS; 600 mM dithiothreitol [DTT]; 60% glycerol; 0.6% bromophenol blue) and analyzed by SDS-PAGE and immunoblotting.
In vitro kinase assays. (i) Integrin-associated kinase assay.
Anti-ß1 integrin immunoprecipitates were washed once in CHAPS buffer, twice in HBS (20 mM HEPES, pH 7.4; 150 mM NaCl), and twice in kinase buffer (50 mM HEPES, pH 7.0; 10 mM MnCl2; 10 mM MgCl2; 2 mM NaF; 1 mM sodium orthovanadate). Immunoprecipitates were resuspended in 30 µl of kinase buffer, and reactions were initiated by addition of 1 µl (10 µCi) of [
-32P]ATP and incubated at 30°C for 20 min. Reactions were terminated by the addition of 10 µl of 6x SDS-PAGE sample buffer and boiling for 5 min. The supernatants were resolved by SDS-10% PAGE and electroblotted to nitrocellulose filters according to standard protocols. Phosphorylated proteins were detected by exposure of membranes to Kodak XAR-5 film at -70°C for appropriate times.
(ii) IRS-1-associated PI3K assay.
Anti-IRS-1 immunoprecipitates were assayed for in vitro PI3K activity by using [-32P]ATP and phosphatidylinositol (Avanti Polar Lipids) as substrates, as described previously (34). Phosphorylated products were separated on Silica Gel 60 TLC plates (Merck) and detected by autoradiography.
(iii) Cdk assay.
Cells were lysed in a lysis buffer containing 50 mM HEPES (pH 7.4), 50 mM NaCl, 0.5% Triton X-100, 1 mM DTT, 20 mM NaF, 10 mM NaPPi, 400 µM PAO, 1 mM sodium vanadate, 1% aprotinin, and 5 µg of leupeptin/ml. Then 200 µg of total cellular protein was immunoprecipitated with anti-cyclinE Ab for 16 h at 4°C, followed by the addition of protein A-Sepharose beads and further incubation for 1 h. Beads were collected by centrifugation, washed twice in lysis buffer without protease and phosphatase inhibitors, and washed once in kinase reaction buffer (50 mM HEPES, pH 7.4; 10 mM MgCl2). Immunoprecipitates were resuspended in 40 µl of kinase reaction buffer supplemented with 50 µM ATP, 1 µg of histone H1, and 0.7 µCi of [-32P]ATP and then incubated for 20 min at 30°C. The reaction was terminated by boiling after the addition of the appropriate volume of 6x SDS loading buffer, and the supernatant was analyzed as described above for integrin-associated kinase assay.
SDS-PAGE and immunoblotting. Protein samples were fractionated on SDS-polyacrylamide gels and electroblotted onto nitrocellulose membranes according to standard protocols. The membrane was blocked by incubating in TBS-Tween (10 mM Tris-HCl, pH 7.4; 50 mM NaCl; 0.1% Tween 20) containing 5% (wt/vol) nonfat dry milk or 3% (wt/vol) BSA (for RC20-HRP) and sequentially incubated with the indicated primary Ab and a HRP-conjugated secondary anti-mouse or anti-rabbit Ab (depending on the primary Ab). Membranes were then washed with TBS-Tween and developed by the enhanced chemiluminescence method according to the manufacturer's instructions (Amersham). Where indicated, the membrane was stripped of Ab by three washes with glycine buffer (0.2 M glycine, pH 2.0) and three washes with TBS-Tween and then immunoblotted with the indicated Ab.
Subcellular fractionation. (i) S100/P100 fractionation. Cells were placed on ice and washed twice with Tris-Glu buffer. Cells were then collected in Tris-Glu buffer and centrifuged at 1,500 rpm for 5 min at 4°C. Cells were then resuspended in a hypotonic buffer (20 mM Tris-HCl, pH 7.4; 10 mM KCl; 1 mM EDTA; 1 mM DTT; 1% aprotinin; 1 mM PMSF) and kept on ice for 10 min. The cells were Dounce homogenized with a type B pestle (20 to 40 strokes) until >90% of the cells were broken and the release of nuclei was confirmed by examination under a microscope. The cell suspension was adjusted with NaCl to 125 mM. The cell lysate was then centrifuged at 1,500 rpm for 10 min to pellet the nuclei and cell debris (P1 fraction), and the supernatant was collected and further centrifuged at 100,000 x g in polycarbonate tubes for 30 min at 4°C. The supernatant (S100 fraction), containing cytosolic components, was removed and adjusted to 1% Triton X-100. The pellet (P100 fraction), containing cellular membrane components, was solubilized in RIPA buffer containing 0.1% SDS and then centrifuged at 12,000 rpm for 10 min at 4°C to generate the soluble P100 fraction (supernatant). The S100 and soluble P100 fractions were analyzed by immunoprecipitation, SDS-PAGE, and immunoblotting.
(ii) CSK fractionation. Cells were placed on ice, washed twice with Tris-Glu buffer, and gently incubated in 0.5 ml of CSK buffer (1% Triton X-100; 10 mM PIPES [piperazine-N,N'-bis(2-ethanesulfonic acid)], pH 6.8; 100 mM KCl; 2.5 mM MgCl2; 1 mM CaCl2; 300 mM sucrose; 1 mM PMSF; 1% aprotinin; 1 mM sodium orthovanadate) for 3 to 5 min. The buffer was collected with minimal perturbation of the cell monolayer. Another 0.5 ml of CSK buffer was added to the cells, which were incubated and collected as described above, and combined together to form the Triton-soluble fraction which was then adjusted to 1% SDS. The remaining cellular material was extracted from the tissue culture dish with 1 ml of Western lysis buffer to generate the Triton-insoluble fraction. Both fractions were boiled with SDS-PAGE sample buffer for 5 min, and equivalent aliquots were analyzed by SDS-PAGE and immunoblotting.
Immunofluorescence cell staining. Cells were washed twice with Tris-Glu buffer, trypsinized, and counted by hemacytometer. Typically, 5 x 104 cells were plated on 22-by-22-mm glass coverslips (Fisher) placed in 35-mm dishes. Coverslips were routinely coated with poly-L-lysine (Sigma) for 30 min and allowed to dry prior to cell plating. At 40 h after being plated, cells were washed twice in 1x Hanks balanced salt solution (Gibco-BRL) and fixed at room temperature with 2% formaldehyde-0.05% glutaraldehyde-0.05% Triton X-100 for 20 min. After fixation, cells were washed twice with phosphate-buffered saline (PBS), incubated for 30 min in blocking solution (3% BSA and 0.05% Triton X-100 in PBS), and then incubated for 1 h with anti-paxillin, anti-vinculin, or anti-IGFR Ab. After three rinses with wash buffer (0.1% NP-40 in PBS), cells were incubated in the dark with FITC-conjugated anti-mouse or anti-rabbit secondary Abs or phalloidin for 1 h. Cells were rinsed three times with wash buffer and mounted onto microscope slides with the ProLong Antifade Kit (Molecular Probes) containing an antifade agent. Labeled cells were examined with an Olympus IX70 fluorescence microscope. Swiss 3T3 cells were fixed with 4% paraformaldehyde in PBS at room temperature for 20 min. After three washes with PBS, cells were permeabilized with 0.4% Triton X-100 for 3 min at room temperature. Anti-IGFR (1:200), anti-paxillin (1:50), anti-vinculin (1:50), anti-mouse IgG-FITC (1:200), and anti-rabbit IgG-FITC (1:200) Abs and phalloidin-FITC (1:200) and phalloidin-TRITC (1:200) were diluted in blocking buffer and centrifuged at maximum speed for 3 min prior to using the supernatant to label the cells.
Receptor internalization assay.
Exponentially growing cells were trypsinized, counted, and seeded at a density of 106/10-cm dish. At 48 h after being plated, cells were incubated in serum-free DMEM for 20 h before incubation in serum-free DMEM supplemented with 100 ng of IGF-I/ml for 0, 2, 5, 10, 20, 30, or 50 min. Cells were placed on ice, washed twice with cold PBS containing 5 mM EDTA (PBS-EDTA), and collected by pipetting up and down in 4 ml of PBS-EDTA. Cells were incubated on ice with anti-IGFR Ab diluted 1:100 in PBS-EDTA for 45 min. After 1 wash with cold PBS, cells were incubated on ice with anti-mouse IgG(H+L)-FITC secondary Ab diluted 1:50 in PBS for 30 min. After two washes with cold PBS, cells were fixed at room temperature with 8% formaldehyde in PBS for 30 min. After two washes with PBS, cells were resuspended in 1 ml of PBS and stored in the dark prior to flow cytometric analysis at the Mount Sinai cytometry facility.
Cell cycle analysis. Synchronization of cells in G1 was accomplished by incubation of confluent cultures in DMEM with 0.5% BCS for 24 h. Cells were then incubated in DMEM with 2% BCS and 100 ng of IGF-I/ml for 48 h. Cells were washed twice in Tris-Glu buffer, trypsinized, and counted before and after treatment with IGF-I. Approximately 106 cells were removed and washed with cold PBS by gentle vortexing and centrifugation at 1,000 rpm (300 to 350 x g) for 5 min. Cells were then fixed at 4°C with 70% ethanol for 2 h. After two washes with PBS, cells were gently resuspended in 1 ml of DNA staining solution (PBS, pH 7.4; 1 µg of RNase A/ml; 20 µg of propidium iodide/ml). Samples were stored in the dark at 4°C prior to flow cytometry analysis at the Mount Sinai cytometry facility.
Monolayer cell growth. Exponentially growing cells were trypsinized, counted by hemacytometer, and plated at cell densities of 5 x 103 or 1 x 105 cells/35-mm dish as indicated in the figure legends. At 24 to 36 h after being plated, the cells were incubated in fresh growth medium containing 5 or 2% BCS, with or without IGF-I (100 ng/ml). Cells were counted on day 0 (i.e., the first day of medium change) and every other day thereafter. Medium with or without IGF-I was changed every other day.
Colony assay. Subconfluent cells in exponential growth were trypsinized, counted, and evenly resuspended at a density of 105 cells/3 ml of DMEM containing 0.3% Bacto Agar, antibiotics, and 7.5% BCS, with or without IGF-I (100 ng/ml). Cell suspensions were layered in 6-cm dishes over a 0.7% agar base containing the above components. Cells were grown at 37°C with addition of 0.5 ml of DMEM containing 7.5% BCS, plus 100 ng of IGF-I/ml where appropriate, every 3 to 4 days. After 4 weeks, colonies of >0.2 mm were counted from 10 randomly selected fields at x40 magnification.
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by IGF-IR (32) is consistent with this notion.
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were associated with both RACK1 and IGF-IR. PMA treatment also led to an association of PKCµ with RACK1 and IGF-IR in these cells (Fig. 1B). In Rat2-IGFR fibroblasts, treatment with IGF-I and PMA each enhanced the association of RACK1 with PKCµ (Fig. 1B) but not with PKC or PKCß (data not shown). We suspect that the experimental conditions used were not sufficient to demonstrate the PMA-induced in vivo association of PKCß and RACK1, which was reported previously (47). These results indicate that IGF-I and PMA are each capable of stimulating the association of certain PKC isozymes ( and µ) with RACK1 and IGF-IR. Although these data together are not conclusive, they do suggest that activation of IGF-IR or PKC may lead to the formation of a protein complex comprised of IGF-IR, PKC, and RACK1 in mammalian fibroblast and epithelial cells. Subcellular redistribution of RACK1 in oncogene-transformed mammalian and avian cells. We have previously characterized the activation of signaling pathways and biological behavior of mammalian and avian cells transformed by PTK oncogenes including those of the IR family. To begin to understand the potential role of RACK1 in cell transformation, we analyzed the subcellular distribution of RACK1 via P100 and S100 fractionation of oncogene-transformed cells and their respective untransformed controls. Subcellular fractionation of RACK1 in cell lines stably transformed by various oncogenes revealed a significant redistribution of RACK1 from the cytosolic to the membrane fraction in 3Y1 fibroblasts transformed by v-Src and RIE cells transformed by NM1 (encoding an oncogenic IGF-IR [35]) or T6 (encoding an oncogenic IR [43]) (Fig. 2A), but not in NIH 3T3 fibroblasts that were transformed by an activated H-Ras mutant (data not shown), compared to their respective controls. These results suggest that redistribution of RACK1 to the cellular membrane may be induced by PTK oncogenes. To further explore this idea, we infected CEF with NM1, UR2 (encoding the v-Ros PTK receptor-like oncogene [39]), or SR-A (encoding v-Src) virus and analyzed the subcellular distribution of RACK1 after the cells were morphologically transformed (Fig. 2B). Consistent with the results obtained in mammalian cells, there was a redistribution of RACK1 to the membrane fraction in NM1- and SR-A-infected cells compared to control CEF. Intriguingly, membrane localization of RACK1 was not observed in UR2-infected cells, suggesting that some but not all PTK receptor oncogenes can cause localization of RACK1 to the membrane (Fig. 2B). Furthermore, infection of CEF with the virus encoding ts kinase mutants of v-Src (tsSrc), v-Ros (tsROS), or NM1 (tsNM1) demonstrated that the oncogene-induced localization of RACK1 to the cellular membrane did not occur 2 h after the infected cells were shifted from a nonpermissive to a permissive temperature (Fig. 2C). At this time the PTKs of these oncogenes were found to be activated (data not shown). Translocation of RACK1 to the membrane occurred in CEF infected with tsSrc and tsNM1 but not tsROS after a 48-h shift to the permissive temperature (Fig. 2C) when full cellular morphologic transformation was clearly evident (data not shown), a finding which is in agreement with the results obtained with the constitutively activated versions of these oncogenes (Fig. 2B). Therefore, these results suggest that localization of RACK1 to the membrane does not occur as an immediate response to the kinase activity of these oncogenes. Instead, this redistribution may occur secondary to the transformation induced by certain oncogenic PTKs. However, the biological significance of the altered localization of RACK1 to the cellular membrane in such transformed cells is currently unclear.
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RACK1 overexpression increases cell spreading, focal adhesions, and actin stress fibers. Initial observations that RACK1-overexpressing cells assumed a less-refractile appearance under light microscopy compared to control cells (Fig. 5A, right) suggested to us that RACK1 overexpression might alter the cytoskeleton and/or cellular adhesion properties of NIH 3T3-IGFR cells. To further investigate the effect of RACK1 overexpression on the actin cytoskeleton and focal adhesions, we used immunofluorescence microscopy to examine RACK1-overexpressing and control cells that were stained with phalloidin or Abs specific for paxillin and vinculin, proteins known to be present in focal adhesions (Fig. 4A). Phalloidin staining revealed that the total number of actin stress fibers in RACK1-overexpressing cells was generally increased with a distribution over a greater area per cell relative to control cells (Fig. 4A, top panels). Indeed, anti-IGFR staining showed that individual RACK1-overexpressing cells generally occupied a greater surface area (Fig. 4A, bottom panels), suggesting that overexpression of RACK1 results in increased spreading of NIH 3T3-IGFR cells. Anti-paxillin staining demonstrated that the total number of focal adhesions, particularly in areas away from the cell periphery, was increased in RACK1-overexpressing cells compared to control cells, in which focal adhesions were restricted primarily to the ends of cell projections, e.g., filopodia (Fig. 4A, second row). Staining with anti-vinculin Ab demonstrated a similar increase in focal adhesions in RACK1-overexpressing cells relative to control cells (Fig. 4A, third row). Together, these findings indicate that overexpression of RACK1 in NIH 3T3-IGFR cells results in an increase in cell spreading, which is consistent with an increase in focal adhesions and actin stress fibers within each cell.
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Overexpression of RACK1 inhibits IGF-I-dependent cell growth and transformation. Overexpression of RACK1 in NIH 3T3 cells was previously reported to inhibit the growth of those cells in normal growth conditions (13). We found that, consistent with this observation, RACK1 overexpression suppressed the exponential monolayer growth of NIH 3T3-IGFR cells (data not shown). Because RACK1 overexpression in NIH 3T3-IGFR cells caused an increase in cell spreading (Fig. 4A) and reduced light refractility of cells in confluent culture (Fig. 5A, right), we investigated whether RACK1 overexpression affected the ability of these cells to proliferate at high cell densities. Growth analysis of cells seeded initially at 75% confluence showed that both the initial growth rates and the final saturation densities of RACK1-overexpressing cell lines were 40 to 80% lower than those of the empty vector-transfected control cultures (Fig. 5A, left). These results were consistent among independent clones of RACK1-overexpressing cells and suggest that overexpression of RACK1 suppresses cell growth in monolayer cultures at both low and high cell densities.
The association of RACK1 with IGF-IR after its activation by IGF-I (Fig. 1A) suggests that RACK1 plays a direct role in the biological functions mediated by IGF-IR. We chose to examine the effect of RACK1 overexpression in NIH 3T3 cells that stably expressed human IGF-IR at levels two- to threefold above the endogenous IGF-IR level because it would allow us to analyze IGF-I-inducible cell proliferation in both anchorage-dependent (i.e., monolayer) and anchorage-independent (i.e., soft agar) conditions since NIH 3T3 cells expressing elevated levels of IGF-IR can be induced by IGF-I to form colonies (29, 32, 35). To determine the effect of RACK1 overexpression on IGF-I-dependent monolayer growth, equivalent numbers of RACK1-overexpressing and control cells were seeded in medium containing a low concentration of serum with or without supplemental IGF-I, and cell numbers were subsequently determined at regular intervals (Fig. 5B). In the presence of only a low concentration of serum, the growth rate of each RACK1-overexpressing cell line was inhibited relative to control NIH 3T3-IGFR cells at all of the time points examined. The addition of exogenous IGF-I significantly stimulated the growth rate of the control cells but not of the RACK1-overexpressing cells. A comparison of the IGF-I-dependent increase in cell density after 6 days showed that the growth of NIH 3T3-IGFR control cells was stimulated by 100% compared to 30 to 60% for the RACK1-overexpressing cells. These results indicate that overexpression of RACK1 inhibits IGF-I-inducible cell growth in anchorage-dependent conditions.
To determine the effect of RACK1 overexpression on IGF-I-induced anchorage-independent growth, we analyzed the colony-forming ability in soft agar of control and RACK1-overexpressing NIH 3T3-IGFR cells. As expected, the number of colonies in control cultures that were supplemented with IGF-I was increased by 160% (Fig. 5C). In contrast, the colony growth of RACK1-overexpressing cells was increased by only 6 to 28% (Fig. 5C), indicating that overexpression of RACK1 significantly suppresses the IGF-I-mediated anchorage-independent growth of NIH 3T3-IGFR cells. Independent stable clones that overexpressed RACK1 with either an amino-terminal HA epitope (HA-RACK1) or a carboxyl-terminal histidine tag (RACK1-His) also displayed a reduction of IGF-I-induced colony formation (data not shown), providing additional evidence for a role of RACK1 in cell transformation. Together, these data demonstrate that overexpression of RACK1 inhibits IGF-I-dependent growth under both anchorage-independent and anchorage-dependent conditions, thus implicating RACK1 in IGF-IR-mediated biological functions.
Activation of IGF-IR and its major signaling pathways are not affected. To explore potential mechanisms that may account for the alteration of IGF-I-dependent cell proliferation and transformation in NIH 3T3-IGFR cells, we next examined the major IGF-IR-mediated signal transduction pathways. Control and RACK1-overexpressing cells were serum starved, treated with IGF-I for up to 50 min, and analyzed for the activation and internalization of IGF-IR, the tyrosine phosphorylation and activation of the major IGF-IR substrates Shc and IRS-1, and the activation of components of the PI3K and MAPK pathways (Fig. 6). Overall, our results indicate that there were no significant differences between RACK1-overexpressing and control cells in the early activation kinetics of any of the signaling molecules examined. The IGF-I-stimulated internalization of IGF-IR was detectable as early as 2 min and peaked after 20 min of IGF-I treatment (Fig. 6A). Tyrosine phosphorylation of IGF-IR reached a maximum within 2 min of IGF-I treatment and decreased dramatically after 10 min (Fig. 6B), indicating that overexpression of RACK1 does not affect receptor autophosphorylation, activation, and internalization induced by IGF-I. Tyrosine phosphorylation of IRS-1, association of IRS-1 with p85PI3K and PI3K activity, and phosphorylation of Akt/PKB, a downstream effector of PI3K which is activated by the lipid products of PI3K, were detectable within 2 min, persisting above unstimulated level for at least 50 min (Fig. 6B). The activating phosphorylation of p70S6K, another downstream effector of PI3K, was detectable after 20 min and persisted for at least 50 min (Fig. 6B). The overall increased association of IRS-1 with p85PI3K and PI3K activity is consistent with the slightly elevated IRS-1 level in this RACK1-overexpressing cell line (data not shown). The IGF-I-stimulated tyrosine phosphorylation of Shc was detectable within 2 min and returned to basal levels within 50 min (Fig. 6B). The phosphorylation of ERK1 and ERK2 on activating tyrosine and threonine residues was markedly increased at 2 min, reached a maximum at 5 min, and remained above basal levels after 50 min of IGF-I treatment (Fig. 6B). Together, these data suggest that overexpression of RACK1 does not affect the IGF-I-dependent activation and internalization of IGF-IR and the IGF-I-mediated activation of downstream signal transduction pathways involving Shc, IRS-1, PI3K, and MAPK.
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RACK1 overexpression inhibits cell cycle progression by altering cell cycle regulators. To further explore the mechanism by which overexpression of RACK1 inhibits cell proliferation, we analyzed cell cycle progression in control and RACK1-overexpressing cells by measuring cellular DNA content by flow cytometry analysis after the synchronization of confluent cultures in G0 by serum withdrawal and the subsequent release into G1 and S by treatment with IGF-I (Fig. 9A). We determined the minimum concentration of serum necessary for IGF-I-dependent cell cycle progression in control NIH 3T3-IGFR cells to be 2% BCS (data not shown). After treatment with IGF-I and BCS for 48 h, a significant proportion of control cells (at least 30%) had entered S phase compared with cells that overexpressed RACK1, which were predominantly (>95%) in G0 or G1 (Fig. 9A). This suggests that overexpression of RACK1 in NIH 3T3-IGFR cells leads to inhibition of cell cycle progression in G0, in G1, or at the G1/S transition point and is consistent with the delay in G1 progression observed in an unsynchronized population of NIH 3T3 cells that overexpressed RACK1 (13). Interference of IGF-I-induced cell cycle progression in NIH 3T3-IGFR cells is likely to underlie the suppression of IGF-I-mediated anchorage-independent and anchorage-dependent growth observed in these cells.
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The effect of RACK1 overexpression on the regulation of IGF-I-dependent cell cycle regulation was further examined in these cells. IGF-I treatment of cells for 12 h led to a significant reduction in the level of p27Kip1 concomitant with a dramatic increase in cyclin E-associated kinase activity in control cells (Fig. 9D). However, the extent of these changes in p27Kip1 and cyclin E-associated kinase activity was significantly attenuated in RACK1-overexpressing cells (Fig. 9D). Not surprisingly, the p27Kip1 level in control cells was increased to a level similar to that of RACK1-overexpressing cells upon serum starvation (Fig. 9D). Collectively, these data indicate that overexpression of RACK1 leads to the upregulation of CKIs with a corresponding suppression of Cdk2 activity and Rb phosphorylation, which may at least partially account for the inhibition of IGF-I-stimulated monolayer and anchorage-independent growth observed in these cells.
Blockade of endogenous RACK1 expression by antisense oligonucleotides inhibits cell spreading and cell proliferation. To more precisely establish the function of endogenous RACK1 and to confirm our results from an overexpression system, which may produce nonphysiologic effects, we used antisense oligonucleotides designed to block RACK1 translation, in an attempt to suppress the level of endogenous RACK1 protein and to examine the effect on cell morphology and monolayer proliferation. After unsuccessful attempts with phosphorothioate oligonucleotides and antisense expression vectors, we chose to use morpholino oligonucleotides, which are extremely stable and have recently been used successfully to block the expression and function of endogenous proteins (56). Standard control or RACK1 antisense morpholino oligonucleotides were introduced into Swiss 3T3 cells according to the method recommended by the manufacturer as described in Materials and Methods. An effect on cell morphology and cell proliferation was not immediate and only became evident at least 4 days after introduction of oligonucleotides, which may be explained by the high stability of RACK1 protein (t1/2 = 40 h [data not shown]) and/or suboptimal intracellular concentrations of antisense morpholino oligonucleotide. Beginning 4 days after the introduction of oligonucleotides, Swiss 3T3 cells with antisense oligonucleotides were observed to adopt a more refractile appearance under light microscopy compared to control cells (Fig. 10A, upper left panels) and parental cells (data not shown). Analysis of cell morphology by fluorescence staining with phalloidin demonstrated that, while control cells assumed a spread phenotype similar to that of parental cells, antisense oligonucleotide-treated cells generally displayed a more contracted and less spread morphology (Fig. 10A). Costaining with phalloidin and antivinculin to assess the actin cytoskeleton and focal adhesions, respectively, demonstrated fewer stress fibers in antisense oligonucleotide-treated cells compared to control cells, which were also found to possess a greater number of focal adhesions that were distributed throughout the cell (Fig. 10B). After introduction of the oligonucleotides, the monolayer growth of antisense and control cells was analyzed as described for RACK1-overexpressing cells. Proliferation of antisense cells was reduced by 34% after 2 days and by 50% after 4 days of growth relative to that of control cells (Fig. 10C). Suppression of cell spreading and monolayer growth in antisense cells was correlated with a reduction of the RACK1 protein level by 50 to 60% (Fig. 10D), suggesting that RACK1 plays an important role in cell spreading, formation of focal adhesions, and cell proliferation. The diminished ability of cells to spread when RACK1 expression was reduced by antisense oligonucleotide is consistent with the enhancement of cell spreading observed when RACK1 is overexpressed and thus strongly points to a role for endogenous RACK1 in mediating cell spreading.
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, clone 1; 
, clone 2) and RACK1-overexpressing (
, P; 
, clone 1; 
, clone 4; 
, clone 20) NIH 3T3-IGFR cells were plated at a density of 105 cells/35-mm dish in medium containing 5% BCS and counted in duplicate every other day. Photomicrographs of control (C) and RACK1-overexpressing (clones 1 and 4) cells were taken on day 1 (at subconfluent cell density, x40 magnification) and day 6 (at confluent cell density, x100 magnification). (B) Control (
, clone 9; 
