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[Cancer Research 66, 6183-6191, June 15, 2006]
© 2006 American Association for Cancer Research

Cell, Tumor, and Stem Cell Biology

Osteopontin Is an Oncogenic Vav1– but not Wild-type Vav1–Responsive Gene: Implications for Fibroblast Transformation

Vered Schapira, Galit Lazer and Shulamit Katzav

The Hubert H. Humphrey Center for Experimental Medicine and Cancer Research, The Hebrew University-Hadassah Medical School, Jerusalem, Israel

Requests for reprints: Shulamit Katzav, The Hubert H. Humphrey Center for Experimental Medicine and Cancer Research, The Hebrew University-Hadassah Medical School, P.O. Box 12272, Jerusalem 91120, Israel. Phone: 972-2-6758350; Fax: 972-2-6414583; E-mail: katzav{at}md.huji.ac.il.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Mammalian wild-type Vav1 (wtVav1) encodes a specific GDP/GTP nucleotide exchange factor that is exclusively expressed in the hematopoietic system. Despite numerous studies, the mechanism underlying transformation of fibroblasts by oncogenic Vav1 (oncVav1) is not well defined. We identified osteopontin, a marker for tumor aggressiveness, as an oncVav1-inducible gene. Osteopontin is highly expressed in oncVav1-transformed NIH3T3 cells (NIH/oncVav1) but is barely detected in NIH3T3 expressing wtVav1 (NIH/wtVav1) even following epidermal growth factor stimulation, which normally induces osteopontin. Depleting oncVav1 in NIH/oncVav1 using small interfering RNA led to a considerable decrease in osteopontin, whereas reducing osteopontin expression did not affect oncVav1 expression, suggesting that oncVav1 operates upstream of osteopontin. Vav1-depleted NIH/oncVav1 cells, but not osteopontin-depleted NIH/oncVav1 cells, exhibited impaired extracellular signal-regulated kinase (ERK) and c-Jun NH2-terminal kinase phosphorylation. Inhibition of ERK phosphorylation in NIH/oncVav1 cells led to a decrease in osteopontin expression, implying that the elevated osteopontin expression in these cells is dependent on ERK phosphorylation. Vav1-depleted or osteopontin-depleted NIH/oncVav1 cells lost their tumorigenic properties as judged by the soft agar and invasion assays, although loss of osteopontin expression had a less dramatic effect. Suppression of Vav1 expression in NIH/oncVav1 cells led to reversion to "normal" morphology, whereas when only osteopontin expression was diminished cells retained their transformed morphology. This work strongly supports a role for oncVav1 as a master oncogene and provides clues to the molecular mechanism underlying oncVav1 transformation. (Cancer Res 2006; 66(12): 6183-91)


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Vav1 has been intensively studied by several investigators since it was detected as an oncogene [oncogenic Vav1 (OncVav1)] in 1989 (1). These studies have provided us with several insights into the role of the Vav proteins in mediating intracellular signals that modulate cell growth and differentiation (2). However, many critical questions remain regarding the ways in which Vav1 can also act as a transforming protein.

Vav1, the first Vav family member isolated, is a hematopoietic cell-specific signal transducer protein (1, 3). The other mammalian proteins in this family, Vav2 and Vav3, are more widely expressed (3). There is good evidence that the Vav proteins play a central role in triggering the cytoskeletal reorganization required for many physiologic processes by participating in several distinct pathways as a GDP/GTP nucleotide exchange factor (GEF) for Rho/Rac GTPases (4) and as scaffold proteins (3). Mammalian Vav proteins contain several characteristic structural motifs that enable their function as signal transducer proteins (2, 3). These include a dbl homology (DH) region that exhibits a GEF activity toward the Rho family GTPases, a pleckstrin homology domain that interacts with polyphosphoinositides, a Src homology (SH) 2 and two SH3 domains that mediate protein-protein interactions, a proline-rich motif that enables binding to SH3-containing proteins, and an acidic-rich region and a calponin homology (CH) region at the NH2 terminus of the protein that function as actin-binding domain in other proteins. An additional important characteristic of the Vav proteins is the close correlation between their activity as GEFs and their phosphorylation on tyrosine residues, which is regulated by activation of cell surface receptors (5).

OncVav1 was activated in vitro by replacement of 67–amino acid residues of the Vav1 NH2 terminus (part of its CH region) with 19–amino acid residues of pSV2neo sequences that was cotransfected with DNA of human esophageal carcinomas as a selectable marker in the search for novel oncogenes (1). When oncVav1-transformed NIH3T3 cells are injected i.v., they metastasize to mice lungs (6), suggesting that oncVav1 may contribute to the aggressiveness of transformed cells in vivo. In contrast, wild-type Vav1 (wtVav1) produces minimal transformation of NIH3T3 fibroblasts even when the protein is grossly overexpressed. Removal of the NH2-terminal sequences (65 residues), mimicking its original mode of activation, was sufficient to induce Vav1 transformation (7, 8). Mutant Vav1 proteins that have lost a larger portion of the NH2 terminus (CH domain plus the acidic-rich region up to 186–amino acid residues; {Delta}1-186) exhibit a more pronounced transforming activity than oncVav1, which is missing only 67 residues (9).

The mechanisms underlying transformation by Vav1 have been the subject of intensive research. Because Vav1 functions as a GEF and the morphology of the oncVav1-transformed cells is reminiscent of cells transformed by the dbl oncogene or active versions of Rho family GTPase (10), it seemed conceivable that oncVav1-induced transformation is the result of deregulated GEF activity. Indeed, the mutant Vav1 protein ({Delta}1-186) shows constitutive, phosphorylation-independent exchange activity, correlated with its transforming activity (9). Nuclear magnetic resonance studies showed that Tyr174 within the acidic-rich region of Vav1 makes several contacts with side chains of amino acids within the DH domain (11). Following phosphorylation of Tyr174, the tyrosine is released from its binding pocket, which allows access of the DH domain to its substrate. Yet, the oncVav1 mutant ({Delta}1-66) that retains the inhibitory residue Y174, the entire acidic domain, and part of the CH domain (5) exhibits an exchange activity that is phosphorylation dependent (9).

The possibility that GEF-independent mechanisms might also be involved in Vav1 transformation has recently emerged. For instance, the activity of Vav1 in GEF-independent pathways was shown to be critical for its activity in T cells, including Ca2+ mobilization (12, 13) and stimulation of the transcription factor nuclear factor of activated T cells (14). In addition, association of the SH2 (15) and SH3 (16) regions of oncVav1 ({Delta}1-66) with other proteins was shown to be critical for transformation. Additionally, the NH2-terminal tail could mediate protein-protein interactions that inhibit the biological activity of Vav1 in the resting state and are relieved on activation. Indeed, we have recently described the binding of Ly-GDI, another potential regulator of Rho GTPases, to the NH2 terminus of Vav1 (13). The significance of this interaction to the function of Vav1 in T cells has been shown (13). However, because Ly-GDI is expressed only in the hematopoietic system, other proteins may affect Vav transformation in NIH3T3 fibroblasts via association with the CH region. Thus, the mechanisms of transformation by oncVav1 ({Delta}1-66) remain enigmatic.

Here, we investigated whether oncVav1 ({Delta}1-66) up-regulates specific genes involved in cancer. We now report that osteopontin, a marker of tumor malignancy (17, 18), is overexpressed in cells transformed by oncVav1 ({Delta}1-66) but repressed in cells expressing wtVav1. This study explored the role of osteopontin in oncVav1-induced transformation and investigated the effects of oncVav1 and osteopontin on various signaling pathways.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell Culture
NIH3T3, NIH/oncVav1 (1), and NIH/wtVav1 (7) cells were maintained in DMEM supplemented with 10% calf serum (Life Technologies, Grand Island, NY). For treatment with epidermal growth factor (EGF), cells were grown to subconfluence and then starved in DMEM containing 0.5 mg/mL insulin and transferrin for 24 hours. Cells were then treated with DMEM containing 100 ng/mL murine EGF for the indicated time intervals.

RNA Purification and Semiquantitative Real-time PCR
Total RNA was purified using EZ-RNA kit according to the manufacturer's instructions (Biological Industries, Beit-Haemek, Israel). For reverse transcription, 2 µg total RNA and 0.5 pmol/µL random primers (Promega, Madison, WI) were added to each Moloney murine leukemia virus reverse transcriptase reaction (Promega). The primers used were as follows: 5'-ATTTCCAGCTGTCCATTG-3' (forward) and 5'-CTGAAACACCTCCATCTTG-3' (reverse) for Vav1 detection, 5'-GATCGATAGTCAAGCAAGTT-3' (forward) and 5'-GGTATAGTGATATAGACTGT-3' (reverse) for osteopontin detection, and 5'-CCAAGGTCATCCATGACAAC-3' (forward) and 5'-GGCCATGAGGTCCACCACC-3' for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) detection.

Quantitative Real-time PCR
Real-time PCR (RT-PCR) was done using assays on demand that contain Taqman PCR master mix (Applied Biosystems, Foster City, CA) and the required primers. The following assays on demand were used: hOPN, #Hs00167093_m1; mOPN, #Mn00436767_m1; hVav1, #Hs00232108_m1; and h18sRNA, #Hs99999901_s1. Nontemplate control (contamination sample's reagents) and samples with 18S RNA for normalization were also used. The analysis was done using the ABI Prism 7000 RT-PCR technology (Applied Biosystems). The assays were done thrice in triplicate.

Transfection
The calcium phosphate precipitate method of transfection was used for all transient transfections with Vav1 small interfering RNA (siRNA) plasmids. LipofectAMINE 2000 was used for introduction of the osteopontin siRNA duplexes (100 pmol siRNAs) following the manufacturer's instructions (Invitrogen, Carlsbad, CA). To monitor transfection efficiency, green fluorescent protein (GFP) plasmid was cotransfected.

Antibodies and Immunofluorescence Reagents and Immunofluorescence Microscopy
The following antibodies were used in our studies: anti-Vav1 polyclonal antibodies were raised in rabbits against a specific peptide of Vav1 (residues 528-541; 7); anti-Vav1 monoclonal antibody (mAb); anti-phosphotyrosine mAb (4G10), anti-Rac, and anti–extracellular signal-regulated kinase (ERK) antibodies (Upstate Biotechnology, Lake Placid, NY); anti-phosphorylated phospholipase (PLC) {gamma} (Tyr783), anti-PLC{gamma} antibodies, anti-phosphorylated ERK mAbs, anti-phosphorylated c-Jun NH2-terminal kinase (JNK) mAbs, and anti-phosphorylated JNK (Cell Signaling Technology, Beverly, MA); anti-osteopontin (R&D Systems, Minneapolis, MN); and anti-actin (Santa Cruz Biotechnology, Santa Cruz, CA); donkey anti-goat and goat anti-rabbit Cy3-conjugated antibodies (Jackson ImmunoResearch, West Grove, PA) were used in the immunofluorescence analysis. Immunofluorescence and confocal microscopy analysis were done as described previously (13).

Immunoprecipitation and Immunoblotting
Cells lysis, immunoprecipitation, and immunoblotting were done as described previously (13).

RNA Interference in NIH/oncVav1 Cells
OncVav1. Two siRNA targeting sequences were selected corresponding to the open reading frame of human Vav1 gene using an online design program (Promega). These sequences included a synthetic double-stranded oligonucleotide (5'-TCGAGTGCCTATGCAGCGAGTTCgagtactgGAACTCGCTGCATAGGCATTTTT-3') that corresponds to a 19-nucleotide sequence from wtVav1 (nucleotide 979-998), which are separated by a 9-nucleotide linker (lowercase letters) from the reverse complement of the same 19-nucleotide sequence, and an additional synthetic double-stranded oligonucleotide (5'-TCGAGCGAGACAACGAGACACTGgagtactgCAGTGTCTCGTTGTCTCGCTTTTT-3') that corresponds to wtVav1 (nucleotides 1,217-1,236) and is composed of the same characteristics mentioned above. Both sequences used for siRNA plasmid construction were blasted to avoid silencing other unrelated genes. These oligonucleotides were subcloned into the Vector pSuppressor, which generates biologically active siRNAs from the U6 promoter (Imgenex, San Diego, CA), thus yielding si1Vav and si2Vav, respectively. We used a circular control plasmid that contains a scrambled sequence without significant homology to rat, mouse, or human gene sequences [scrambled vector (scVector)]. The sequences of siRNA insert were verified by DNA sequencing.

Osteopontin. siRNA duplexes were designed according to Teramoto et al. (19): OPNi-840 (sense GUUUCACAGCCACAAGGACdTdT and antisense GUCCUUGUGGCUGUGAAACdTdT) and the negative control, the scrambled OPNi (sense CAGUACAACGCAUCUGGCAdTdT and antisense UGCCAGAUGCGUUGUACUGdTdT). These siRNA duplexes are called OPNi and scOPN throughout, respectively.

Cell Invasion Assay (Matrigel Invasion Assay)
We used the Matrigel invasion chambers (BD Biosciences, San Jose, CA) to assess migration (20). Briefly, a single-cell suspension of 200,000 cells per well was added to the upper chamber of a prehydrated membrane filter of 8-mm pore size detailed above. The lower chamber was filled with conditioned medium of semiconfluent NIH3T3 cells. Cell invasiveness was determined at the lower surface of the filter after 24 hours following the recommended procedure by BD Biosciences. Noninvading cells on the upper side of the filter and Matrigel were wiped off and migrating cells on the reverse side of the filter were stained with DiffQuick, which consists of a methanol fixative, buffered eosin, and phosphate-buffered azure B (Dade Behring, Deerfield, IL). Experiments were reproduced thrice in triplicate and counting was done on five different fields of each filter.

Soft Agar Assay
Cells were transiently transfected as indicated (see Results), trypsinized, suspended in DMEM containing 0.4% agar and 10% calf serum, and plated onto a bottom layer containing 0.8% agar. The cells were plated at a density of 1 x 105 per well in a six-well plate, and colonies were counted 14 days later. Each experiment was done thrice in triplicate.

ERK Inhibition
NIH/oncVav1 and NIH/wtVav1 cells were grown to 80% confluence and then starved in DMEM containing 5 mg/mL insulin and transferrin for 24 hours. The cells were then incubated for 2 hours with 10 µmol/L UO126.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Osteopontin expression is elevated in NIH3T3 cells transformed by oncVav1. To explore the transforming potential of oncVav1, we investigated differences in gene expression in oncVav1 and wtVav1 cells by probing a Cancer Gene Array with cDNAs prepared from three populations of NIH3T3 cells: untransfected, stably transfected with oncVav1 (NIH/oncVav1), and stably transfected with wtVav1 (NIH/wtVav1). Using this approach, several genes exhibited different patterns of RNA expression in these cell populations (data not shown). Strikingly, osteopontin RNA was highly expressed in NIH/oncVav1 cells compared with NIH3T3 and NIH/wtVav1 cells. Osteopontin is a secreted adhesive glycoprotein with diverse functions and has been implicated in cancer development, progression, and metastasis (17, 18). In fact, osteopontin was recently used as a marker for the aggressiveness of tumors (17, 18).

Semiquantitative RT-PCR confirmed that osteopontin is highly expressed in NIH/oncVav1 cells compared with NIH3T3 and NIH/wtVav1 cells (Fig. 1A ). Osteopontin was not detectable in NIH/wtVav1 cells even after 30 cycles of PCR (Fig. 1A). The housekeeping gene, GAPDH, showed no significant differences in its RNA expression in the three lines used (Fig. 1A). RT-PCR results were compatible with the semiquantitative PCR results, showing osteopontin RNA expression 14 times higher in NIH/oncVav1 cells than in NIH/wtVav1 cells (data not shown). Similar results were obtained following transient transfection of NIH3T3, Jurkat T cells, and HCT (a human colon tumor cell line) with plasmids containing wtVav1 or oncVav1 (data not shown), attesting to the generality of our observation.


Figure 1
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  		Figure 1. Expression of osteopontin mRNA in noninduced NIH3T3, NIH/oncVav1, and NIH/wtVav1 cells (A) and following stimulation with EGF (B and C). A, RNA was prepared from the indicated established cell lines and used as templates in a semiquantitative RT-PCR assay done with osteopontin (OPN)–specific primers for 27 and 30 cycles. GAPDH was amplified on the same samples for 35 cycles. B and C, NIH3T3, NIH/oncVav1, and NIH/wtVav1 cells were either left untreated (0 hour) or treated with EGF for various time intervals (1, 6, and 24 hours). At the indicated times, cells were lysed and RNA was prepared and analyzed for osteopontin (B) and Vav1 (C) RNA expression using semiquantitative RT-PCR. Thirty and 33 cycles were done for osteopontin and Vav1 (B and C) and 21 cycles were done for detection of GAPDH (B and C). Representative of three experiments.

 
EGF stimulation of NIH3T3 cells induces osteopontin (21). As expected, osteopontin mRNA was elevated in NIH3T3 cells following 1 hour of EGF treatment and remained elevated after 24 hours of treatment (Fig. 1B). In contrast, osteopontin mRNA expression was very high in unstimulated starved NIH/oncVav1 cells and did not increase following EGF stimulation (Fig. 1B). Osteopontin mRNA was not detectable in untreated NIH/wtVav1 cells, appearing only after 24 hours of stimulation with EGF (Fig. 1B). EGF had no effect on the expression of oncVav1 and wtVav1 (Fig. 1C). These results were supported by Western blot analysis of osteopontin protein expression (data not shown). Thus, oncVav1 induces a dramatic increase in osteopontin expression regardless of EGF stimulation.

Knocked down Vav1 expression in NIH/oncVav1: effects on osteopontin expression and signal transduction cascades. To determine whether the oncogenicity of oncVav1 is solely dependent on osteopontin expression and/or on its participation in various signal transduction cascades, we suppressed oncVav1 and osteopontin in NIH/oncVav1 cells by using siRNA and analyzed the biological consequences of such treatments. RT-PCR analysis showed strikingly diminished expression of oncVav1 in NIH/oncVav1 cells treated with si1Vav or si2Vav plasmids (78% and 58% reduction below control levels, respectively; Fig. 2A ). Treatment of NIH/oncVav1 cells with Vav1 siRNA plasmids also significantly reduced osteopontin mRNA expression (60-80%; Fig. 2A). Treatment of NIH/oncVav1 cells with OPNi, which was reported previously to be an effective siRNA duplex for down-regulating osteopontin (19), led to a 70% reduction in osteopontin expression in these cells (Fig. 2A). No change in oncVav1 mRNA levels was noted in osteopontin-depleted NIH/oncVav1 cells (Fig. 2A). Western blot analysis shows that oncVav1 protein expression was considerably reduced in NIH/oncVav1 cells treated with either si1Vav or si2Vav plasmids, whereas no change was seen in the level of other proteins, such as actin or Rac (Fig. 2B). Osteopontin protein levels in NIH/oncVav1 cells were reduced following treatment with OPNi, si1Vav, or si2Vav but were unchanged in scOPN-treated cells (Fig. 2C). These results further establish the link between oncVav1 and osteopontin mRNA and protein expression and validate the use of Vav1 siRNA as a tool to drastically reduce the expression of osteopontin and oncVav1. Moreover, these results clearly indicate that oncVav1 operates upstream of osteopontin.


Figure 2
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  		Figure 2. siRNA-mediated knockdown of oncVav1 (A) and osteopontin RNA (A) and protein (B and C) expression in NIH/oncVav1 cells. NIH/oncVav1 cells were transiently transfected with either an empty vector, two siRNA plasmids directed against Vav1-specific sequences (si1Vav and si2Vav), or a scrambled (scOPN) or an effective OPNi siRNA duplex. Microarray analysis (A) and Western blotting (B and C) were done after 48 hours. A, RNA prepared from the transfected cell lysates was tested by RT-PCR for expression of oncVav1 and osteopontin and normalized by the expression of 18S RNA in each sample. Experiments were done in triplicate and reproduced thrice. Bars, SE. B, NIH/oncVav1 cells transiently transfected with a scVector (lane 1), an empty vector (Vector; lane 2), and si1Vav and si2Vav plasmids (si1Vav and si2Vav; lanes 3 and 4) were lysed 48 hours following transfection. Cell lysates were separated on SDS-PAGE and immunoblotted with anti-Vav1, anti-actin, and anti-Rac antibodies. Similar results were obtained in three independent experiments. C, a similar experiment to (B) was done using scrambled (scOPN) and effective OPNi siRNA duplexes. Cell lysates were separated on SDS-PAGE and immunoblotted with anti-osteopontin and anti-actin antibodies. Similar results were obtained in three independent experiments.

 
To assess how reduced oncVav1 and osteopontin expression affect signal transduction cascades in NIH/oncVav1 cells, we analyzed the level of oncVav1 tyrosine phosphorylation following EGF treatment of cells transfected with unrelated vector, si1Vav, or si2Vav (Fig. 3A ). oncVav1 was tyrosine phosphorylated in NIH/oncVav1 cells treated with EGF (Fig. 3A). As expected, treatment with si1Vav or si2Vav dramatically reduced oncVav1 expression; the small amount of oncVav1 remaining seems to be tyrosine phosphorylated (Fig. 3A, lanes 5-8).


Figure 3
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  		Figure 3. Signaling pathways following siRNA-mediated knockdown of oncVav1 in NIH/oncVav1 cells stimulated with EGF. A, NIH3T3 cells (lanes 1 and 2) and NIH/oncVav1 cells (lanes 3 and 4) were transiently transfected with an empty vector. NIH/oncVav1 cells were also transiently transfected with si1Vav (lanes 5 and 6) and si2Vav (lanes 7 and 8) plasmids. Cells were either nonstimulated (–; lanes 1, 3, 5, and 7) or stimulated with EGF (+; lanes 2, 4, 6, and 8). Cell lysates were immunoprecipitated with anti-Vav antibodies, and proteins were resolved on SDS-PAGE and then immunoblotted with either anti-phosphotyrosine (top) or anti-Vav antibodies (bottom). B to D, cell lysates were prepared as in (A), except that NIH/oncVav1 cells were also transfected with a scVector. Proteins were separated on SDS-PAGE and immunoblotted with either anti-phosphorylated ERK mAb (B, pERK, top) or anti-ERK mAbs (B, bottom), anti-phosphorylated JNK mAb (C, pJNK, top) or anti-JNK mAbs (B, bottom), or anti-PLC{gamma} mAb (D, pPLC{gamma}, top) or anti-PLC{gamma} mAbs (D, bottom).

 
Several reports clearly link wtVav1 to the activation of ERK, a known Ras-dependent kinase in various hematopoietic cell lineages, via a pathway involving the Ras GTPase, B-Raf, mitogen-activated protein kinase (MAPK) kinase (MEK) 1, and MEK2 kinases (22–24). Additionally, the MEK inhibitor PD90859 inhibited Vav1-induced activation of ERK (23). However, the activation of Ras and ERK appeared normal following stimulation of the T-cell receptor (TCR) in Vav1-null Jurkat T cells (25). The link between oncVav1 and ERK activation has thus far not been resolved. We analyzed ERK phosphorylation in NIH/oncVav1 cells treated with Vav1siRNA plasmids (Fig. 3B). ERK phosphorylation is increased following stimulation of NIH3T3 cells with EGF and in EGF-treated NIH/oncVav1 cells transfected with a scVector or an unrelated vector (Fig. 3B). This increase is eliminated in cells transfected with either si1Vav or si2Vav despite the substantial amount of ERK protein present in these samples (Fig. 3B). Our results clearly show that oncVav1 is required for ERK phosphorylation in NIH/oncVav1 cells following EGF treatment.

Mammalian wtVav1 has been shown to activate JNK in various cells, such as fibroblasts (26), mast cells (27), and T cells (28). We analyzed JNK phosphorylation in NIH/oncVav1 cells with or without knockdown of oncVav1. JNK phosphorylation was inhibited in NIH/oncVav1 cells following transfection with oncVav1 siRNA, although total JNK protein levels remained high (Fig. 3C).

Vav1 controls Ca2+ flux by participating in the regulation of PLC{gamma} activation in the hematopoietic system (12, 13). However, it is not known if oncVav1 plays a role in PLC{gamma} phosphorylation in fibroblasts. We found that treatment with either si1Vav1 or si2Vav1 had no effect on PLC{gamma}1 phosphorylation (Fig. 3D). Thus, oncVav1 is critical for ERK and JNK phosphorylation but not for PLC{gamma} phosphorylation in NIH/oncVav1 cells (Fig. 3).

Osteopontin gene expression in NIH/oncVav1 is affected by phosphorylation of ERK. We next examined the possibility that the up-regulation of osteopontin in NIH/oncVav1 cells might stem from the function of oncVav1 as a signal transducer protein. ERK phosphorylation is elevated to a greater extent in EGF-stimulated NIH/oncVav1 cells than in NIH/wtVav1 cells (Fig. 4A ). Moreover, a basal level of phosphorylated ERK is noted in unstimulated NIH/oncVav1 cells but not in NIH/wtVav1 cells (Fig. 4A). To examine whether increased phosphorylation of ERK in NIH/oncVav1 cells is responsible for the observed elevation in osteopontin expression, we treated NIH/wtVav1 and NIH/oncVav1 cells with a MEK inhibitor (UO126) and analyzed the levels of ERK phosphorylation and osteopontin protein expression (Fig. 4A and B). Treatment with UO126 inhibited ERK phosphorylation in both NIH/wtVav1 and NIH/oncVav1 cells (Fig. 4A). Concomitantly, osteopontin expression was markedly reduced in NIH/oncVav1 cells (Fig. 4A). The level of osteopontin in NIH/wtVav1 cells remained barely detectable (Fig. 4A). Analysis of osteopontin gene expression by RT-PCR confirms that UO126 inhibits osteopontin expression in NIH/oncVav1 (data not shown).


Figure 4
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  		Figure 4. Osteopontin gene expression in NIH/oncVav1 is affected by phosphorylation of ERK (A and B), but its depletion does not affect ERK phosphorylation (C). A, NIH/wtVav1 cells (lanes 1-4) and NIH/oncVav1 cells (lanes 5-8) were either nonstimulated (–; lanes 1, 3, 5, and 7) or stimulated with EGF for 1 hour (+; lanes 2, 4, 6, and 8). Cells were either treated for two hours with UO126 before EGF stimulation (+; lanes 3, 4, 7, and 8) or remained nontreated (–; lanes 1, 2, 5, and 6). Cell lysates were resolved on SDS-PAGE and then immunoblotted with either anti-pERK (top), anti-ERK (middle), or anti-osteopontin antibodies (bottom). Representative of one of two experiments. B, densitometry was done on blots from two separate experiments. Results are relative expression of either pERK/ERK or osteopontin/ERK. Bars, SE. C, NIH3T3 cells (lanes 1 and 2) and NIH/oncVav1 cells (lanes 3-8) were transiently transfected with an empty vector (lanes 1-4), scOPN (lanes 5 and 6), and OPNi siRNA duplexes (lanes 7 and 8). Cells were either nonstimulated (–; lanes 1, 3, 5, and 7) or stimulated with EGF (+; lanes 2, 4, 6, and 8). Cell lysates were resolved on SDS-PAGE and then immunoblotted with either anti-phosphorylated ERK mAb (top) or anti-ERK mAbs (bottom).

 
The finding that UO126 blocked the expression of osteopontin in NIH/oncVav1 cells indicates that expression of this gene is coupled to the ERK pathway. To further substantiate these results, we examined whether signaling events, such as ERK phosphorylation, are compromised by osteopontin depletion (Fig. 4C). Following EGF stimulation, ERK was phosphorylated in NIH/oncVav1 cells transfected with either an empty vector and in cells treated with scOPN or OPNi (Fig. 4C). Thus, a reduction in osteopontin expression did not alter ERK phosphorylation in contrast to the loss of ERK phosphorylation seen following reduction of oncVav1 expression (Fig. 3B). Osteopontin depletion had no effect on JNK and PLC{gamma} phosphorylation following EGF stimulation with the same paradigm shown in Fig. 4C (data not shown).

Taken together, the reduction in osteopontin expression due to inhibition of ERK phosphorylation (Fig. 4A and B) and the fact that depletion of osteopontin does not affect ERK phosphorylation (Fig. 4C) clearly indicate that expression of osteopontin can be coupled to the ERK pathway. Because osteopontin expression in NIH/oncVav1 cells is not markedly induced following EGF stimulation (Figs. 2 and 4), it is conceivable that the basal level of activation of ERK in these cells, but not in NIH/wtVav1 cells, suffices for osteopontin overexpression (mRNA and protein levels) in cultured NIH/oncVav1 cells.

Tumorigenic properties of NIH/oncVav1 cells depleted of oncVav1 or osteopontin. NIH/oncVav1 cells exhibit transformed morphology: the cells are round with many cytoplasmic extensions. To determine whether this morphology is dependent on oncVav1 expression, we transfected NIH3T3 and NIH/oncVav1 cells with a GFP expression plasmid and either an empty expression plasmid (vector) or the plasmid carrying si1Vav1 and analyzed expression of oncVav1 and GFP and the morphology of the transfected cells (Fig. 5A ). oncVav1 accumulated in the cytoplasm of NIH/oncVav1 cells but was not found in NIH3T3 cells (Fig. 5). Following transfection with si1Vav1 plasmid, no expression of oncVav1 was noted in NIH/oncVav1 cells expressing GFP. Notably, these cells also underwent a dramatic change in cell shape, reverting to a nontransformed phenotype similar to NIH3T3 cells (Fig. 5A). Transfection with si2Vav1 plasmid had similar effects (data not shown). Treatment of NIH/oncVav1 cells with OPNi had no effect on the morphology of NIH/oncVav1 cells (Fig. 5B).


Figure 5
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  		Figure 5. Morphology and expression of oncVav1 (A and B) and osteopontin (B) in NIH/oncVav1 cells transfected with specific Vav1 plasmids and osteopontin siRNA duplexes. A, NIH3T3 and NIH/oncVav1 cells were cotransfected with an empty vector or Vav1-specific siRNA plasmids together with the GFP. Cells were permeabilized and stained with anti-Vav antibodies followed by Cy5-conjugated anti-rabbit antibodies (red). Cells were then analyzed by confocal microscopy. Green, GFP; Merge, an overlay of Vav and GFP staining; DIC, differential interference contrast image. B, a similar experiment to (A), except that NIH/oncVav1 cells were treated with scrambled (scOPN) and OPNi siRNA duplexes. Cells were stained as detailed above and analyzed for oncVav1 or osteopontin expression. More than one cell is presented in each panel.

 
NIH/oncVav1 cells are transformed cells, which have been shown to be metastatic (6). We tested whether NIH/oncVav1 cells depleted of oncVav1 or osteopontin retain their transforming properties as manifested by growth in agar (Fig. 6A and B ) and Matrigel invasion (Fig. 6C and D). The growth in agar assay has been used many years as a justifiable assay to test the transforming properties of cells. NIH/oncVav1 cells transfected with si1Vav1 exhibit greatly reduced ability to grow in agar compared with NIH/oncVav1 cells transfected with the scVector (Fig. 6A and B). OPNi-treated NIH/oncVav1 cells also exhibited a considerable reduction in growth compared with the scOPN-treated cells (Fig. 6A and B). The size of the colonies of NIH/oncVav1 treated with si1Vav1 was smaller than the ones observed in cells treated OPNi (Fig. 6B).


Figure 6
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  		Figure 6. Tumorigenic properties of NIH/oncVav1 cells transfected with Vav1 and osteopontin-specific siRNA plasmids: Growth in soft agar (A and B) and invasiveness (C and D). A, NIH/oncVav1 cells treated as indicated were trypsinized 48 hours following transfection, suspended in DMEM containing 0.4% agar and 10% calf serum, and plated onto a bottom layer containing 0.8% agar. Cells were plated at a density of 1 x 105 per well in a six-well plate, and the number of colonies was counted 14 days later. Representative of three experiments in triplicates. Bars, SE. B, four representative colonies grown in agar for each treatment described in (A). C, NIH3T3, NIH/oncVav1, and NIH/oncVav1 cells transfected with Vav1 and osteopontin-specific siRNA plasmids or duplexes were assayed by the in vitro Matrigel assay that enables the estimation of invasiveness of cells through ECM. Representative of five experiments in duplicates. D, a representative staining of the invading cells described in (A).

 
The in vitro Matrigel invasion assay is commonly considered to be a measurement of the first step of tumor cell invasion (20). NIH/oncVav1 cells transfected with si1Vav1 and si2Vav1 plasmids have greatly reduced ability to invade Matrigel (82% for si1Vav1 and 70% for si2Vav1), with levels of invasiveness similar to the nontransformed parent NIH3T3 cells (Fig. 6C and D). OPNi-treated NIH/oncVav1 cells exhibited a 45% reduction in invasion compared with scOPN-treated cells (Fig. 6C and D), suggesting that osteopontin is only one part of the mechanism by which oncVav1 controls tumorigenesis.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We have shown here that oncVav1, but not wtVav1, up-regulates the osteopontin gene and explored the contribution of osteopontin to oncVav1-mediated transformation. Osteopontin is considered a marker of the malignant phenotype (17, 18). Elevated levels of osteopontin in a wide variety of malignant cells and in the surrounding stroma of numerous human cancers suggest that it may play a key role in tumorigenesis (17, 18). Currently, increased osteopontin expression is associated with tumor invasion, progression, or metastasis in cancers of the breast, stomach, lung, prostate, liver, and colon (17, 18). Osteopontin is not expressed in the normal matrix of most tissues, other than mineralized tissues, but it is synthesized by cells in response to various stimuli and thus is considered to be a cytokine (29).

Increased expression of osteopontin has also been found in cells transformed by other oncogenes. Elevated osteopontin expression was noted in NIH3T3 fibroblasts transformed by ras (30), and these ras-transformed fibroblasts require osteopontin for growth (31). The v-src oncogene stimulates osteopontin expression in murine cells (32) and osteopontin synthesis is suppressed in cells that do not express c-src (33), suggesting that the wild-type gene may also play a role in normal expression of osteopontin. It was recently reported that osteopontin expression is also induced in a model of murine leukemia caused by Bcr-Abl (34). These studies show that a variety of oncogenes can induce osteopontin, but none of them directly compared osteopontin expression in the presence of the corresponding proto-oncogenes. We show here not only that oncVav1 stimulates the expression of osteopontin but also that wtVav1 depresses its expression. This result raises the possibility that osteopontin might contribute to the transforming capability of oncVav1.

Our demonstration that osteopontin is up-regulated in NIH/oncVav1 cells is the first report of an effect of oncVav1 on downstream genes. Recently, several groups reported elevated expression of cyclin D1 due to the presence of a different oncVav1 mutant ({Delta}1-186). Abe et al. (9) reported that the increase in Vav1 transforming activity correlated with the activation of the c-Jun, Elk-1, and nuclear factor-{kappa}B (NF-{kappa}B) transcription factors as well as increased transcription from the cyclin D1 promoter. They also showed that wtVav1 has little, if any, effect on any of these activities. However, this study also indicated that the oncVav1 used in our study (oncVav1{Delta}1-66) does not induce cyclin D1 (9). Fernandez-Zapico et al. reported a different effect of wtVav1 on cyclin D1 expression, showing that wtVav1 up-regulates cyclin D1 in pancreatic tumors (35). This discrepancy might result from differences in the experimental systems. Our observation that oncVav1 and wtVav1 have opposite effects on osteopontin expression is unique.

Several mechanisms might explain the oncVav1-mediated up-regulation of osteopontin. One possibility is that this induction is direct, via an effect of oncVav1 on the osteopontin promoter as was shown for ras-mediated osteopontin overexpression (36). However, this option was ruled out because we could not detect any induction by oncVav1 of the osteopontin promoter plasmids (data not shown). The other possibility is that the osteopontin up-regulation in NIH/oncVav1 cells stems from changes in signaling pathways, such as ERK, in these cells. Indeed, treatment of NIH/oncVav1 cells with the specific MEK1/2 inhibitor UO126 resulted in a marked decrease in osteopontin expression along with down-regulation of ERK phosphorylation (Fig. 4). Osteopontin expression in NIH/wtVav1 cells was barely detectable even before the treatment with UO126. These data suggest that sustained ERK phosphorylation, such as that observed in NIH/oncVav1 cells, is required to induce osteopontin (Fig. 4). These results are further supported by the fact that ablation of osteopontin in NIH/oncVav1 cells does not change the level of EGF-induced ERK activation (Fig. 4C). Thus, in our experimental system, ERK phosphorylation takes place upstream of osteopontin expression. The MAPK signaling cascade has also been implicated in osteopontin regulation in various cell systems, such as HL-60 cells (37), JB6 cells (38), and human osteoblastic cells (39), and in response to injury of rat arterial muscle cells (40). The association between ERK phosphorylation and osteopontin expression was shown by the use of both MEK inhibitors (34, 37, 38, 40) and a dominant-negative ERK plasmid (39). Interestingly, the dependence of osteopontin up-regulation on ERK activation was shown in cells transformed by another oncogene, Bcr-Abl, via a cascade, including Ras, phosphatidylinositol 3-kinase, {alpha}-protein kinase C, Raf-1, and MEK (34). Together, these data suggest that activation of the MAPK cascade is a crucial component in the regulation of osteopontin expression regardless of the stimulus. It is plausible that additional signaling cascades activated by oncVav1 in NIH3T3 cells also contribute to osteopontin up-regulation.

Numerous studies have implicated wtVav1 in ERK (22–24), JNK (26–28) and PLC{gamma} (12, 13) phosphorylation, but most of these studies were done in hematopoietic cells, where wtVav1 is physiologically active. Little was known thus far regarding the involvement of oncVav1 in these pathways in fibroblasts. Our studies clearly show a role for oncVav1 in ERK and JNK phosphorylation but not in PLC{gamma} phosphorylation (Fig. 3). oncVav1 probably influences ERK phosphorylation in a different manner than wtVav1 in T cells. In T cells, wtVav1 transduces TCR signals to Ras by controlling the membrane recruitment of RasGRP1 and Sos1 and Sos2 through the transmembrane adapter protein LAT (22). However, as several of the hematopoietic signaling proteins, including LAT, are absent in fibroblasts, oncVav1 might function in these cells through its association with GRB2 (41) or through additional unknown mechanisms. The lack of involvement of oncVav1 in PLC{gamma} phosphorylation in fibroblasts (Fig. 3) is in agreement with our previous study in T cells (13), pointing to a cardinal difference in signaling pathways between oncVav1 and wtVav1.

Depleting oncVav1 in NIH/oncVav1 cells drastically reduces their ability to grow in soft agar and migrate through Matrigel (Fig. 6), a model for tumor invasiveness. The fact that osteopontin ablation also reduces the growth in soft agar and migration of NIH/oncVav1 cells, although to a lesser extent (Fig. 6), attests to the importance of osteopontin expression for tumorigenesis and cell migration, a critical step in tumor invasion and metastasis. Reorganization of the actin cytoskeleton is the primary mechanism of cell motility and is essential for most types of cell migration. Actin reorganization is regulated by Rho family small GTPases, such as Rho, Rac, and Cdc42 (42, 43). These proteins alternate between an inactive GDP-bound state and an active GTP-bound state. When the protein is in the active GTP-bound conformation, it interacts with effector proteins that propagate further signaling events that lead ultimately to the desired biological responses. As a known GEF for the Rho/Rac GTPases, wtVav1 participates in their regulation, mainly manifested by cytoskeleton reorganization (44). For example, Vav1 is involved in the transmission of migratory signals in T cells (45). Ablation of oncVav1 by Vav1-specific siRNA reverted the morphology of the transformed NIH/oncVav1 cells, whereas ablation of osteopontin expression had a minimal effect on cell morphology (Fig. 5), suggesting that osteopontin does not contribute to cytoskeleton reorganization but to another process involved in motility. Similarly, it was shown that the regulation of endothelial cell motility in the presence of osteopontin is Rac independent (46).

Osteopontin and Vav1 might be linked through integrin signaling. wtVav1 plays a critical role as a downstream signal transducer protein following integrin engagement (47, 48). ß3 Tyrosine phosphorylation and {alpha}vß3-mediated adhesion are required for Vav1 association and Rho activation in leukocytes (47). Additionally, Vav proteins were shown to play an essential role coupling ß2 to Rho GTPases and regulating multiple integrin-induced events important in leukocyte adhesion and phagocytosis (48). Osteopontin contains a RGD tripeptide sequence that can be bound by {alpha}vß3, {alpha}vß1, and {alpha}vß5 integrins on the cell surface (49). The binding of integrins to osteopontin results in distinct functional consequences, including cell adhesion and cell migration (50). Osteopontin also interacts with cellular receptor(s) other than integrins, such as CD44, through domains that do not contain the RGD sequence (51, 52). CD44 has been implicated in a wide variety of cellular processes, including cell-cell and cell-matrix interactions, cell migration, metastasis, lymphocyte homing, and T-cell activation (53). In NIH3T3 cells transformed by H-RasV12, osteopontin mainly interacts with CD44 and not with integrins (19). This interaction contributes to the malignancy of these cells (19). It is conceivable that osteopontin functions in a similar manner in NIH/oncVav1 cells, enhancing the effect of oncVav1 transforming properties via a feed-forward mechanism.

Alternatively, the contribution of osteopontin to the metastatic process could be a result of its ability to regulate the activity of several extracellular matrix (ECM)–degrading proteins (54, 55). Such enzymes play a key role in events, which enable cancer cells to invade into the surrounding environment (54, 55). Philip et al. have shown that osteopontin up-regulates pro-matrix metalloproteinase-2 expression in a NF-{kappa}B-dependent fashion during ECM invasion (54). Osteopontin also increases cell invasiveness in human mammary carcinoma through stimulation of urokinase plasminogen activator (55). The cell-matrix adhesive and cytokine-signaling properties of osteopontin provide mechanisms for mediating these invasive processes (54, 55).

In summary, our studies reveal that osteopontin plays a role in the molecular mechanism that underlies transformation by oncVav1 but that additional pathways are involved that do not include osteopontin. More studies are necessary to identify additional genes that are differentially regulated by oncVav1 and wtVav1. This is particularly relevant in light of recent reports that Vav1 is an important gene in mammalian neoplasms (35, 56).


   Acknowledgments
 
Grant support: Israel Science Foundation (599/04) and the Israel Cancer Association in memory of Profs. Dafna and Dov Israeli.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Dr. Susan Lewis for editing the article, Y. Markson for help with the figures, Dr. M. Tarshish for assistance with the confocal microscopy, Dr. R. Bar-Shavit for the HCT cell line, and Dr. N. Hijaya for the osteopontin promoter plasmids.


   Footnotes
 
Note: V. Schapira and G. Lazer contributed equally to this work.

Received 10/17/05. Revised 2/15/06. Accepted 4/ 5/06.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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