ARTICLE

Dual Mechanisms for Lysophosphatidic Acid Stimulation of Human Ovarian Carcinoma Cells

Yu-Long Hu, Chris Albanese, Richard G. Pestell, Robert B. Jaffe

Affiliations of authors: Y.-L. Hu, R. B. Jaffe, Center for Reproductive Sciences, University of California, San Francisco; C. Albanese, R. G. Pestell, Division of Hormone Dependent Tumor Biology, Albert Einstein College of Medicine, Bronx, NY.

Correspondence to: Robert B. Jaffe, M.D., Center for Reproductive Sciences, HSW 1695, University of California, San Francisco, 505 Parnassus Ave., San Francisco, CA 94143–0556 (e-mail: jaffer{at}obgyn.ucsf.edu).


    ABSTRACT
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Background: Lysophosphatidic acid (LPA), at concentrations present in ascitic fluid, indirectly stimulates the growth of malignant ovarian tumors by increasing the expression of vascular endothelial growth factor (VEGF) in ovarian cancer cells. We investigated whether LPA could also directly promote ovarian tumor growth by increasing the level of cyclin D1, a key G1-phase checkpoint regulator, which thereby increases cell proliferation. Methods: Expression of cyclin D1 and LPA receptors (EDG4 and EDG7) was determined in six ovarian cancer cell lines (including OVCAR-3 cells) and immortalized ovarian surface epithelial cells (IOSE-29). Cyclin D1 promoter activity was measured in LPA-treated OVCAR-3 cells cotransfected with cyclin D1 promoter-driven luciferase constructs and cDNA expression plasmids for I{kappa}B{alpha}M (a nuclear factor {kappa}B [NF{kappa}B] super-repressor). Results: Four of six cancer cell lines, including OVCAR-3, overexpressed cyclin D1 protein relative to levels in IOSE-29 cells. LPA treatment increased cyclin D1 protein in a dose- and time-dependent manner in OVCAR-3 cells but not in IOSE-29 cells. LPA stimulated cyclin D1 promoter activity (3.0-fold, 95% confidence interval [CI] = 2.7-fold to 3.3-fold). Mutation of the NF{kappa}B-binding site in the cyclin D1 promoter to block NF{kappa}B binding and expression of I{kappa}B{alpha}M, which binds NF{kappa}B and inhibits its binding to the promoter, markedly diminished LPA stimulation of cyclin D1 promoter activity (activity stimulated only 1.4-fold, 95% CI = 1.1-fold to 1.7-fold, and 0.7-fold, 95% CI = 0.6-fold to 0.8-fold, respectively). EDG4 was overexpressed in all cancer cell lines studied relative to that in IOSE-29 cells, but EDG7 was overexpressed in only two lines. Conclusions: Dual mechanisms are probably involved in LPA stimulation of ovarian tumor growth in vivo. In addition to the previously characterized indirect mechanism that increases angiogenesis via VEGF, LPA may directly increase the level of cyclin D1 in ovarian cancer cells, increasing their proliferation.



    INTRODUCTION
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Ovarian carcinoma, predominantly derived from ovarian surface epithelium (OSE), is the most lethal gynecologic cancer in the developed world (1). It is characterized by widespread intraperitoneal dissemination and large volumes of ascitic fluid (2), which can stimulate ovarian cancer growth in vivo and in vitro. A mitogen in ascitic fluid has been identified as lysophosphatidic acid (LPA) (3,4). LPA, a bioactive phospholipid present in serum at concentrations of 2–20 µM (5), induces various cellular responses by interacting with specific cell-surface G protein-coupled receptors of the endothelial differentiation gene (EDG) subfamily (6,7). EDG2 (8), EDG4 (9), and EDG7 (10,11) are specific LPA receptors present on many cell types. LPA is found in high concentrations in ascitic fluid and plasma of ovarian cancer patients (12).

Angiogenesis is essential for tumor growth (13) and is induced by the binding of vascular endothelial growth factor (VEGF) to one of two VEGF receptors, Flt1 and KDR. Cancer patients have increased levels of serum VEGF, and elevated levels of VEGF mRNA have been observed in the majority of human cancer cells, including ovarian cancer cells (14). VEGF directly stimulates the growth of some malignancies (e.g., leukemia, lymphoma, and myeloma) that express KDR and/or Flt1 through an autocrine mechanism (15,16). We have previously demonstrated (17) that LPA stimulates VEGF expression in the ovarian cancer cell lines OVCAR-3, SKOV-3, and CAOV-3 through transcriptional activation but that LPA does not stimulate the expression of VEGF in nontumorigenic IOSE cells. Thus, LPA indirectly stimulates ovarian tumor growth, at least in part, by increasing angiogenesis via VEGF.

Some ovarian cancer cell lines (e.g., DOV-13, Hey-A8, and OCC-1), however, do not express VEGF (17,18) but are still sensitive to LPA (3,17), indicating that there is another mechanism involved in regulating their growth. LPA stimulates cell proliferation mediated by serum response element (SRE)-driven recruitment of immediate-early response genes associated with growth (6) and stimulates SRE-driven luciferase activity in ovarian cancer cells but not in IOSE cells (19). Overexpression of c-Fos, which has an SRE-binding site in its promoter region (20), increases the expression of cyclin D1 mRNA in fibroblasts (21). Cyclin D1, a member of a protein family that regulates cyclin-dependent protein kinase activity, can act as an oncogene and has been implicated in the development of several human neoplasms (22,23). It is a key regulator of the G1-phase checkpoint and promotes cell cycle progression from G1 phase to S phase. Cyclin D1 is overexpressed in various human cancers, including ovarian cancer (24,25). Antisense cyclin D1 cDNA expression abolishes growth of pancreatic, hepatocellular, and breast carcinoma cells in nude mice (2628), indicating a critical role for cyclin D1 in tumorigenesis. In our study, we investigated whether LPA directly promotes ovarian tumor growth by increasing the level of cyclin D1, which increases cell proliferation.


    METHODS
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Reagents and Cell Lines

We purchased 1-oleoyl-lysophosphatidic acid (i.e., LPA) and fatty acid-free bovine serum albumin from Sigma Chemical Corp. (St. Louis, MO). All restriction enzymes were from Promega (Madison, WI). All cell culture reagents were from the Cell Culture Facility, University of California, San Francisco. Ovarian cancer cell lines OVCAR-3, SKOV-3, and CAOV-3 were from the American Type Culture Collection (Manassas, VA). Ovarian cancer cell lines Hey-A8, OCC-1, and DOV-13 were provided by Gordon Mills and Robert Bast (University of Texas M. D. Anderson Cancer Center, Houston). IOSE-29 (simian virus 40 T antigen-immortalized normal ovarian surface epithelial cells) and normal OSE cells were provided by Nelly Auersperg (University of British Columbia, Vancouver, Canada). Human umbilical vein endothelial cells (HUVECs) were from Clonetics (San Diego, CA). Cyclin D1, c-Jun, c-Fos, and actin antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA). EDG4 and EDG7 antibodies were from Calbiochem (La Jolla, CA). pLuc-MCS (multiple cloning sites; control), pAP1-Luc, pNF{kappa}B-Luc (where NF{kappa}B is nuclear factor {kappa}B), and pSRE-Luc cis-reporting plasmids were from Stratagene (La Jolla, CA).

Cell Culture and LPA Stimulation

OVCAR-3, Hey-A8, OCC-1, and DOV-13 cells were cultured in RPMI-1640 medium with 10% fetal calf serum (FCS). CAOV-3 and SKOV-3 cells were cultured in Dulbecco’s modified Eagle medium with 10% FCS. IOSE-29 and normal OSE cells were cultured in medium 199/MCDB 105 medium with 10% FCS. For LPA stimulation, cells were plated on six-well plates and cultured in complete growth medium. When cells reached confluence, they were washed twice with prewarmed phosphate-buffered saline (PBS) and cultured in serum-free medium overnight. LPA (0.2–20 µM) was added to the cultures, and cultures were incubated at 37 °C, as indicated. After incubation, cells were harvested and used to isolate total cellular RNA for northern blot analysis.

RNA Extraction and Northern Blot Analysis

Total cellular RNA was isolated from ovarian epithelial cancer cell lines and normal OSE cells by use of a High Pure RNA isolation kit (Roche, Indianapolis, IN) according to the manufacturer’s instructions. Equal amounts of total RNA (10 µg per lane) were separated by electrophoresis on denaturing 1.2% agarose gels containing 2.2 M formaldehyde and transferred to nylon membranes. RNAs were cross-linked to the membranes with UV irradiation and hybridized in Expresshyb hybridization buffer (BD Biosciences Clontech, Franklin Lakes, NJ) at 68 °C for 2 hours with 32P-labeled cDNA probes for Flt1, KDR, cyclin D1, or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) synthesized with a random primer rediPrime DNA labeling kit (Amersham Pharmacia Biotech, Piscataway, NJ), according to the manufacturer’s instructions. The blots were washed for three 20-minute periods at room temperature in 2x standard saline citrate (SSC) (1x SSC = 150 mM sodium chloride and 15 mM sodium citrate) containing 0.05% sodium dodecyl sulfate (SDS) and then washed for two 30-minute periods at 50 °C in 0.1x SSC containing 0.1% SDS. Washed membranes were then exposed to Kodak X-Omat AR film (Eastman Kodak, Rochester, NY) with two intensifying screens for 1–2 days at –70 °C.

Western Blot Analysis

After LPA exposure, ovarian cancer cells and IOSE-29 cells were washed with ice-cold PBS and lysed in 50 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM EDTA, 1% Nonidet P-40, 1 mM phenylmethylsulfonyl fluoride, aprotinin at 10 µg/mL, leupeptin at 10 µg/mL, and pepstatin at 10 µg/mL. Lysates were clarified by centrifugation at 20 800g for 20 minutes at 4 °C. Supernatants were collected, and 50 µg of total protein was subjected to SDS–polyacrylamide gel electrophoresis in 10% gels. Proteins were transferred to a polyvinylidene difluoride membrane and probed with antibodies directed against cyclin D1, EDG4, EDG7, c-Jun, c-Fos, and actin. Blots were then washed, and bands were visualized by incubation with horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology) and enhanced chemiluminescence reagents (Amersham Pharmacia Biotech).

Preparation of Cyclin D1 Promoter-Luciferase (CD1-Luc) Fusion Constructs

A 1882-base-pair (bp) PvuII fragment of human cyclin D1 (CD1) genomic clone, which contains the entire promoter region, was subcloned into the vector pA3-Luc to form the construct –1745 CD1-Luc (29). The constructs –66 CD1-Luc, –66 CD1/ATFm-Luc, –66 CD1/NF{kappa}Bm-Luc, and –22 CD1-Luc were created by polymerase chain reaction with specific primers, as described previously (2931). The ATF/CRE site in the –66 CD1/ATFm-Luc construct was mutated from 5'-TAACGTCAC ACGGAC-3' to 5'-TcgCGTCcCcCGGAC-3' (where lowercase letters are mutated bases), and the NF{kappa}B site in the –66 CD1/NF{kappa}Bm-Luc construct was mutated from 5'-AGGGGAGTTTT-3' to 5'-AccccAGTTTT-3'. The 3' end of the cyclin D1 promoter in all CD1-luciferase reporter constructs is +138 bp relative to the transcription start site. Various CD1-Luc constructs (e.g., –1745 CD1-Luc, –66 CD1-Luc, and –22 CD1-Luc) that contain different lengths of the cyclin D1 promoter region were used to identify possible LPA response elements in the cyclin D1 promoter. Constructs –66 CD1/NF{kappa}Bm-Luc and –66 CD1/ATFm-Luc were used to test whether NF{kappa}B and ATF transcription factors are involved in LPA stimulation of cyclin D1 transcription.

Cell Transfection and Luciferase Activity Measurements

For cell transfection, OVCAR-3 cells were plated in 12-well cluster plates (1.5 mL of medium per well) in triplicate. When OVCAR-3 cells were approximately 70% confluent, they were transfected for 2 hours with 1 µg of a CD1-Luc fusion construct, 7.5 µL of Superfect transfection reagent (Qiagen, Valencia, CA), and 0.02 µg of pRL-CMV, an internal control plasmid containing the cytomegalovirus (CMV) promoter linked to a constitutively active Renilla luciferase reporter gene. OVCAR-3 and IOSE-29 cells were 70% confluent for cotransfection, and incubations were for 2 hours. For cotransfection experiments with OVCAR-3 cells, 0.2 µg of CMV-I{kappa}B{alpha} mutant (I{kappa}B{alpha}M) plasmid or pCMX control expression plasmid, 1 µg of a CD1-Luc fusion construct, and 0.02 µg of pRL-CMV plasmid were cotransfected. For cotransfection experiments with IOSE-29 cells, 0.2 µg of a cis-reporting plasmid or 1 µg of a CD1-Luc fusion construct; 1 µg of pCDEF3 control expression vector containing the human polypeptide elongation factor 1{alpha} (EF1{alpha}) promoter (another control for EDG4 and EDG7), EDG4/EF3, or EDG7/EF3 expression constructs; and 0.02 µg of pRL-CMV plasmid were cotransfected. After transfection, the medium was replaced with fresh growth medium, cells were incubated for 24 hours, and the medium was replaced with serum-free medium. After starvation in serum-free medium for 8 hours, cells were incubated in the presence or absence of 20 µM LPA for another 24 hours. Cells were harvested with passive lysis buffer (Promega), and luciferase activity was determined with a Dual-Luciferase Reporter Assay kit (Promega), according to the manufacturer’s protocol. Thus, 56 hours after transfection, cells were harvested, and luciferase activity was determined.

Data Analysis

Each experiment was performed in duplicate or triplicate. All experiments were repeated at least three times on different occasions. The results are expressed as the mean and 95% confidence interval (CI).


    RESULTS
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
VEGF Receptors in HUVECs, Normal OSE Cells, IOSE-29 Cells, and Ovarian Cancer Cell Lines

Because VEGF plays an important role in the autocrine growth stimulation of some neoplasms that express VEGF receptors KDR and/or Flt1 (15,16), we first determined whether the increased VEGF level induced by LPA directly stimulated the proliferation of ovarian cancer cells. Treatment with VEGF had a minimal effect on ovarian cancer proliferation in vitro (data not shown). Total RNAs from four ovarian cancer cell lines, IOSE-29 cells, primary OSE cells, and HUVECs were analyzed by northern blotting for Flt1 and KDR mRNAs. HUVECs expressed both Flt1 and KDR mRNAs; OSE cells expressed KDR mRNA at a size and level similar to that in HUVECs. Flt1 and KDR mRNAs were not detected in IOSE-29 cells and the four human ovarian cancer cell lines tested, whereas GAPDH mRNA was detected in all cell lines (Fig. 1Go).



View larger version (54K):
[in this window]
[in a new window]
 
Fig. 1. Analysis of mRNA encoding vascular endothelial growth factor (VEGF) receptors, Flt1 and KDR, from human umbilical vein endothelial cells (HUVECs), normal ovarian surface epithelial cells (OSE), IOSE-29 cells (simian virus 40 T antigen-immortalized normal ovarian surface epithelial cells), and four ovarian cancer cell lines (OVCAR-3, CAOV-3, SKOV-3, and DOV-13). HUVECs were used as positive controls for VEGF receptor detection. Northern blot analyses were performed with total RNA. The filter was hybridized with 32P-labeled KDR, Flt1, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA probes. GAPDH was the loading control. Blots shown are representative of three experiments performed in duplicate, all with similar results.

 
Cyclin D1 Overexpression in Ovarian Cancer Cell Lines

We used western blot analysis with cyclin D1 antibody to investigate whether cyclin D1 protein was overexpressed in six ovarian cancer cell lines relative to the expression in IOSE-29 cells, which have a cyclin D1 level similar to that of normal OSE cells (data not shown). Cyclin D1 protein (36 kd) was overexpressed in Hey-A8, OCC-1, DOV-13, and OVCAR-3 cells but not in SKOV-3 or CAOV-3 cells (Fig. 2Go). Hey-A8, OCC-1, and DOV-13 cells expressed cyclin D1 at a high level and grew more rapidly than the other cancer cell lines tested in vitro (data not shown).



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 2. Analysis of human cyclin D1 protein expression in IOSE-29 cells (simian virus 40 T antigen-immortalized normal ovarian surface epithelial cells) and six ovarian cancer cell lines (Hey-A8, OCC-1, DOV-13, SKOV-3, CAOV-3, and OVCAR-3). Western blot analyses were performed with 50 µg of total cell protein from each cell line. The blot was probed sequentially with anti-cyclin D1 and anti-actin antibodies. Blots presented are representative of at least three experiments performed in duplicate, all with similar results.

 
Induction of Cyclin D1 Protein by LPA in OVCAR-3 and Other Ovarian Cancer Cell Lines

Because cyclin D1 was overexpressed in ovarian cancer cells, it might be involved in LPA-stimulated ovarian cancer growth. To investigate this possibility, we examined whether LPA affected the expression of cyclin D1 protein in vitro by western blot analysis. We used human OVCAR-3 cells in this study because after intraperitoneal injection, female athymic mice develop intraperitoneal carcinomatosis and massive ascites, similar to those seen in stage III ovarian cancer (17). When OVCAR-3 cells were treated overnight (i.e., 18 hours) with LPA at 0.2 µM, 2 µM, and 20 µM (all physiologic concentrations), cyclin D1 levels increased in an LPA dose-dependent manner compared with untreated controls (Fig. 3, AGo). The level of cyclin D1 was slightly increased by 0.2 µM LPA, increased substantially with 2 µM LPA, and increased further by 20 µM LPA, all compared with untreated control cells. To determine the time course of this effect, OVCAR-3 cells were incubated with or without 20 µM LPA for 2–24 hours. The level of cyclin D1 increased after a 2-hour incubation, reached its maximum level after 6 hours, and continued to be elevated after 18 and 24 hours, all compared with untreated control cultures (Fig. 3, BGo).



View larger version (43K):
[in this window]
[in a new window]
 
Fig. 3. Lysophosphatidic acid (LPA) induction of cyclin D1 protein expression in human OVCAR-3 cells and IOSE-29 cells (simian virus 40 T antigen-immortalized normal ovarian surface epithelial cells) in a time- and concentration-dependent manner. Western blot analyses were performed with 50 µg of total protein extracted from untreated and LPA-treated ovarian cell cultures in each lane. The blots also were probed with an anti-actin antibody as a loading control. Blots shown are representative of three experiments performed in duplicate, all with similar results. In each panel, a representative blot was scanned, and the results were expressed as the fold increase compared with untreated controls, after normalizing to actin. A) OVCAR-3 cells were incubated with LPA at 0, 0.2, 2, and 20 µM for 18 hours before total protein was extracted. After treatment with LPA as indicated, cyclin D1 protein increased by 1.4-fold (0.2 µM), 2.1-fold (2 µM), and 2.4-fold (20 µM). Other experiments had similar results. B) OVCAR-3 cells were incubated with 20 µM LPA for 0, 2, 4, 6, 18, and 24 hours before total protein was extracted. Treatment with LPA increased cyclin D1 protein by 1.2-fold (2 hours), 1.4-fold (4 hours), 1.9-fold (6 hours), 1.8-fold (18 hours), and 1.5-fold (24 hours). C) IOSE-29 cells were incubated for the times indicated with 20 µM LPA before total protein was extracted. In this experiment zero time points were the control. LPA had essentially no effect on cyclin D1 protein in IOSE-29 cells.

 
Because most ovarian cancers are thought to originate from normal OSE cells, we assessed whether cyclin D1 expression was also induced by LPA in OSE cells. Normal OSE constitutes a small component of the ovary, and normal OSE cells survive only five or six passages in culture. Consequently, it is difficult to obtain fresh OSE cells in sufficient quantities for multiple experiments. Therefore, we used IOSE-29 cells instead of normal OSE cells to investigate whether LPA increased cyclin D1 expression. After IOSE-29 cells were treated with 20 µM LPA for 2–24 hours, we found that levels of cyclin D1 were essentially the same as in 0-hour control cells (Fig. 3, CGo).

To determine whether treatment with LPA increased the level of cyclin D1 in other ovarian cancer cells, we incubated three more ovarian cancer cell lines, OCC-1, DOV-13, and CAOV-3, with 20 µM LPA for 24 hours and determined the level of cyclin D1 protein by western blot analysis. Although basal levels of cyclin D1 did vary among the three cancer cell lines, all cell lines treated with LPA contained increased levels of cyclin D1 protein compared with corresponding untreated controls (Fig. 4Go).



View larger version (32K):
[in this window]
[in a new window]
 
Fig. 4. Lysophosphatidic acid (LPA) and human cyclin D1 protein expression in the ovarian cancer cell lines OCC-1, DOV-13, and CAOV-3. Western blot analyses were performed with 50 µg of total protein. The blot was probed with anti-cyclin D1 and anti-actin antibodies. OCC-1, DOV-13, and CAOV-3 cells were incubated in the presence (+) or absence (–) of 20 µM LPA for 24 hours before total protein was extracted. Blots shown are representative of three experiments performed in duplicate, all with similar results.

 
LPA Induction of Cyclin D1 mRNA in OVCAR-3 Cells

We used northern blot analysis to determine whether LPA induced higher levels of cyclin D1 protein and mRNA in OVCAR-3 cells. Cells were treated with various concentrations of LPA for various times. The increased levels of LPA-induced cyclin D1 mRNA corresponded with the increased levels of LPA-induced cyclin D1 protein in a time- and dose-dependent fashion (Fig. 5Go). In the dose–response experiment, the level of cyclin D1 mRNA (4.5 kilobases [kb]) increased progressively from 0.2 to 20 µM LPA. In a time course experiment in which OVCAR-3 cells were incubated with 20 µM LPA, the level of cyclin D1 mRNA increased progressively for 2–6 hours and then declined after 18 and 24 hours; levels at 18 and 24 hours, however, were still higher than the basal level.



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5. Northern blot analysis of cyclin D1 mRNA expression in OVCAR-3 cells in the presence or absence of lysophosphatidic acid (LPA). Blots presented are representative of at least three experiments performed in duplicate, all with similar results. A) OVCAR-3 cells were incubated with LPA at 0, 0.2, 2, and 20 µM for 18 hours before total mRNA was extracted. B) OVCAR-3 cells were treated with 20 µM LPA for 0, 2, 4, 6, 18, and 24 hours before total mRNA was extracted.

 
AP1 and NF{kappa}B in LPA Stimulation of Cyclin D1 Promoter Transcription

The cyclin D1 promoter region contains binding sites for transcription factors SP1, AP1, NF{kappa}B, and ATF/CREB, all of which regulate cyclin D1 transcription (2931). Because treatment with LPA increased the level of cyclin D1 mRNA (4.5 kb) in OVCAR-3 cells in a time- and dose-dependent manner, we investigated whether LPA stimulated cyclin D1 promoter activity by using transient transfection of cyclin D1 promoter-luciferase reporter constructs into OVCAR-3 cells (Fig. 6Go). Luciferase activity of construct –1745 CD1-Luc, which contains the entire cyclin D1 promoter, was increased 3.0-fold (95% CI = 2.7-fold to 3.3-fold) after LPA treatment compared with untreated controls. Luciferase activity of a promoter construct with a deletion between positions –1745 and –66 was increased 2.5-fold (95% CI = 2.1-fold to 2.9-fold) after LPA treatment compared with untreated controls. However, the luciferase activity of construct –22 CD1-Luc, a promoter construct with a minimal promoter fragment (from positions –22 to +138), was not altered by LPA treatment. These results indicate that the LPA response element is located principally between positions –66 and –22 in the cyclin D1 promoter. Luciferase activity of the construct with an ATF/CRE site mutation (at position –57) was stimulated less by LPA (2.0-fold, 95% CI = 1.6-fold to 2.4-fold), and the luciferase activity of the construct with an NF{kappa}B site mutation (at position –33) was stimulated even less (1.4-fold, 95% CI = 1.1-fold to 1.7-fold), both compared with the untreated control for each plasmid. Luciferase activity of the construct with a promoter deletion between positions –1745 and –66 and a mutation of the ATF/CRE or NF{kappa}B site was low with or without LPA treatment, indicating that these transcription factors may have important roles in the basal transcription of cyclin D1 (Fig. 6, AGo).



View larger version (30K):
[in this window]
[in a new window]
 
Fig. 6. Mechanism of lysophosphatidic acid (LPA) stimulation of cyclin D1 expression. OVCAR-3 cells were cotransfected with 1 µg of a CD1-luciferase reporter construct (–1745 CD1-Luc, –66 CD1-Luc, –66 CD1/ATFm-Luc, –66 CD1/NF{kappa}Bm-Luc, and –22 CD1-Luc) and 20 ng of pRL-CMV vector (an internal control plasmid for transfection efficiency). A) LPA stimulation of cyclin D1 promoter activity. Twenty-four hours after transient transfection, the cells were treated with LPA (+LPA) or without LPA (–LPA) at 20 µM for 24 hours, and luciferase activity was measured. Data are the mean and half the 95% confidence interval (error bars) of values from three experiments performed in triplicate (n = 9 points). B) I{kappa}B{alpha} mutant (I{kappa}B{alpha}M, an NF{kappa}B super-repressor) and LPA stimulation of cyclin D1 promoter transcription. OVCAR-3 cells were cotransfected with CMV-I{kappa}B{alpha}M, where CMV is cytomegalovirus) or pCMX control expression plasmids (each at 0.2 µg), the CD1-luciferase fusion construct indicated, and the pRL-CMV vector at the same time. Twenty-four hours after transient transfection, the cells were treated with or without 20 µM LPA for 24 hours, and luciferase activity was measured. Data are the mean and half the 95% confidence interval (error bars) of values from three experiments performed in triplicate (n = 9 points). C) Analysis of LPA induction of protein expression of c-Fos and c-Jun, two major components of the AP1 transcription factor, in OVCAR-3 cells by western blot analysis. OVCAR-3 cells were incubated for the indicated times with 20 µM LPA before total protein was extracted. The blot was probed with anti-c-Fos, anti-c-Jun, and anti-actin antibodies. Actin was the loading control. Blots presented are representative of at least three experiments performed in duplicate, all with similar results.

 
To confirm the role of NF{kappa}B in LPA stimulation of cyclin D1 promoter transcription, we cotransfected OVCAR-3 cells with pCMV-I{kappa}B{alpha}M [NF{kappa}B super-repressor expression plasmid containing I{kappa}B{alpha} cDNA with mutations at Ser-32 and Ser-36 of the amino terminus (32)] or a pCMX control vector with a CD1-luciferase construct in the presence or absence of 20 µM LPA. Expression of the NF{kappa}B super-repressor (I{kappa}B{alpha}M) cDNA abolished the LPA activation of CD1-luciferase activity of constructs –1745 CD1-Luc and –66 CD1-Luc (LPA-stimulated activity was reduced to 0.7-fold, 95% CI = 0.6-fold to 0.8-fold). Cotransfection of the control pCMX expression vector did not interfere with LPA stimulation of luciferase activities of CD1-luciferase constructs. Specifically, LPA still stimulated the –1745 CD1-Luc and –66 CD1-Luc by 3.1-fold (95% CI = 2.7-fold to 3.5-fold) and 1.7-fold (95% CI = 1.4-fold to 2.0-fold), respectively (Fig. 6, BGo).

c-Jun and c-Fos are major components of transcription factor AP1. Both Jun–Jun and Jun–Fos dimers recognize AP1 site located at position –954 in the cyclin D1 promoter, whereas Jun–ATF and Fos–ATF complexes recognize ATF/CRE sites (33) at position –57 or the AP1 site at position –954 in the cyclin D1 promoter (29,30). Because mutation of the ATF/CRE site in the cyclin D1 promoter region and a promoter deletion (from positions –1745 to –66) containing the AP1 site partially diminished LPA activation of the cyclin D1 promoter (Fig. 6, A and BGo), we used western blot analysis to determine whether LPA induced the expression of c-Fos and c-Jun, which bind to these sites, in OVCAR-3 cells. When OVCAR-3 cells were incubated with 20 µM LPA, the level of c-Fos protein (62 kd) increased markedly and consistently for 2–6 hours and then declined by 18 and 24 hours but not to unstimulated control levels. LPA stimulation of c-Jun protein (39 kd) occurred after 2 hours, reached a maximum level after 4 hours, then declined gradually for 6–18 hours, and returned to unstimulated control values after 24 hours. LPA had no effect on the level of actin (internal control) (Fig. 6, CGo).

EDG4 and EDG7 Expression in IOSE-29 Cells and Ovarian Cancer Cell Lines

We next analyzed the expression of the LPA receptor proteins EDG4 and EDG7 in IOSE-29 cells and six ovarian cancer cell lines by western blot analysis. Although EDG7 protein (40 kd) was detected in IOSE-29 cells and the six ovarian cancer cell lines, the highest levels of EDG7 were detected in SKOV-3 and CAOV-3 cells (Fig. 7Go). Levels of EDG7 were similar in normal OSE cells and IOSE-29 cells (data not shown). EDG4 (50 kd) was detected by western blot analysis in all six cancer cell lines but not in IOSE-29 cells (Fig. 7Go). The levels of EDG4 expression detected are consistent with those in our previous study (17), in which we detected EDG4 mRNA in all ovarian cancer cell lines studied but not in normal OSE or IOSE-29 cells.



View larger version (36K):
[in this window]
[in a new window]
 
Fig. 7. Western blot analysis of the lysophosphatidic acid receptors EDG4 and EDG7 in IOSE-29 cells and six ovarian cancer cell lines (Hey-A8, OCC-1, DOV-13, SKOV-3, CAOV-3, and OVCAR-3). The blots also were probed with anti-actin antibody as a loading control. Blots presented are representative of at least three experiments performed in duplicate, all with similar results.

 
Effect of LPA Receptors EDG4 and EDG7 on LPA Stimulation of Cyclin D1 Promoter-, AP1-, NF{kappa}B-, and SRE-Driven Luciferase Transcription in IOSE-29 Cells

We next determined whether IOSE-29 cells acquired LPA responsiveness after the forced expression of EDG4 or EDG7 cDNAs, by assessing the LPA stimulation of cyclin D1 promoter-, AP1-, NF{kappa}B-, and SRE-driven luciferase transcription activity (Fig. 8Go). When EDG4 or EDG7 cDNAs were cotransfected with the AP1-, NF{kappa}B-, or SRE-luciferase plasmid, both cDNAs mediated increased AP1-, NF{kappa}B-, or SRE-driven luciferase transcription activity induced by 20 µM LPA. The increase mediated by EDG4 was larger than that mediated by EDG7. Luciferase activity in IOSE-29 cells cotransfected with pCDEF3 control expression vector (containing the EF1{alpha} promoter) and an AP1-, NF{kappa}B-, or SRE-luciferase plasmid was minimally responsive to LPA; however, the luciferase activity in IOSE-29 cells cotransfected with EDG4/EF3 or EDG7/EF3 expression plasmids and a control pLuc-MCS (containing no transcription factor-binding sites) was not affected by LPA (Fig. 8, AGo). Forced expression of EDG4 or EDG7 cDNAs in IOSE-29 cells increased the LPA stimulatory effect on cyclin D1 promoter activity by 2.6-fold (95% CI = 2.4-fold to 2.8-fold) or 1.8-fold (95% CI = 1.6-fold to 2.0-fold), respectively, from construct –1745 CD1-Luc but not from the shorter construct –22 CD1-Luc construct, both compared with control vector pCDEF3 (Fig. 8, BGo). Thus, primarily EDG4, and to a lesser extent EDG7, are required for LPA stimulation of cyclin D1 expression in ovarian cancer cells.



View larger version (27K):
[in this window]
[in a new window]
 
Fig. 8. Lysophosphatidic acid (LPA) receptors EDG4 and EDG7 and LPA stimulation of serum response element (SRE)-, NF{kappa}B-, and AP1-driven luciferase reporter gene transcription (A) and of cyclin D1 promoter activity (B) in IOSE-29 cells (simian virus 40 T antigen-immortalized normal ovarian surface epithelial cells). SRE-, NF{kappa}B-, AP1-, and MCS-Luc-driven cis-reporting plasmids (A) or CD1-luciferase fusion constructs (B) were cotransfected with pCDEF3 vector containing the human polypeptide elongation factor (EF) 1{alpha} promoter, EDG4/EF3 expression constructs, or EDG7/EF3 expression constructs into IOSE-29 cells. After transfection, the medium was replaced by fresh growth medium, and cells were incubated for 24 hours. After starvation in serum-free medium for 8 hours, cells were incubated in the presence or absence of 20 µM LPA for an additional 24 hours. Cells were harvested with passive lysis buffer; 56 hours after transfection, cells were harvested and luciferase activity was determined. Data are the ratio of values from LPA-treated groups to values from non–LPA-treated controls (mean ± half the 95% confidence interval [error bars]) from three experiments performed in triplicate (n = 9 points).

 

    DISCUSSION
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Cyclin D1 belongs to a family of three closely related D-type cyclins (cyclin D1, cyclin D2, and cyclin D3) that are expressed in all proliferating cell types. In this article, we have shown that cyclin D1 protein is overexpressed in four of the six ovarian cancer cell lines tested. LPA, which can be produced by ovarian cancer cells (34), increased the levels of cyclin D1 protein and mRNA through transcriptional activation in OVCAR-3 cells but not in IOSE-29 cells. LPA also increased cyclin D1 protein in OCC-1, DOV-13, and CAOV-3 cells, indicating that ovarian cancer cells may commonly respond to LPA by this mechanism. LPA appeared to stimulate ovarian tumor growth directly by inducing the expression of cyclin D1, which contributed, at least in part, to increased cell proliferation. LPA appeared to act mainly through EDG4-mediated pathways, because the pattern of LPA-induced expression of cyclin D1 in ovarian cancer cell lines and in IOSE-29 cells was strongly associated with that of EDG4 but not with that of EDG7.

Other pathways can be involved in the development of various human cancers, such as the retinoblastoma (Rb)/cyclin D1/p16 pathway (35). A previous study (36) showed that cyclin D1 was overexpressed in four of six ovarian cancer cell lines and that ovarian cancer cells that coexpress endogenous Rb and p16 were insensitive to overexpression of functional p16 protein. Thus, LPA-induced cyclin D1 in ovarian cancer cell lines with endogenous p16 protein, such as CAOV-3 and OVCAR-3, might circumvent the need to disrupt p16 expression. However, other mechanisms may also be involved, because some cancer cell lines (e.g., Hey-A8, DOV-13, OCC-1, and SKOV-3) express Rb but not p16 protein (37). The lower level of cyclin D1 that we observed in CAOV-3 cells may be caused by their lack of Rb, which would ultimately lead to disassembly of the complex containing cyclin D1 and cyclin-dependent kinase 4/6 and the increased turnover of cyclin D1 (38).

We demonstrated that LPA specifically stimulates cyclin D1 promoter activity approximately threefold and that the LPA response element is located in the promoter region of the cyclin D1 gene, principally between positions –66 and –22, and contains ATF/CRE and NF{kappa}B sites. LPA stimulation of cyclin D1 promoter activity was diminished more by mutation of the NF{kappa}B site than by mutation of the ATF/CRE site. I{kappa}B{alpha}M binds to NF{kappa}B and blocks translocation of NF{kappa}B into the nucleus, which prevents the induction of specific NF{kappa}B target genes (39). Because I{kappa}B{alpha}M abolished LPA-enhanced cyclin D1 promoter activity, we speculate that I{kappa}B{alpha}M inhibited both basal and LPA-inducible cyclin D1 promoter transcription, just as mutation of the NF{kappa}B site in the –66 CD1-Luc plasmid also markedly reduced both basal and LPA-inducible luciferase activity (Fig. 6, AGo). LPA activates NF{kappa}B activity by inducing degradation of the I{kappa}B{alpha} inhibitor (40). When NF{kappa}B signaling is blocked by the forced expression of I{kappa}B{alpha}M, angiogenesis is inhibited and the tumorigenicity of ovarian cancer cells is reduced (41). In another study (31), cyclin D1 protein and mRNA were reduced in myoblast cells expressing I{kappa}B{alpha}M.

Our data demonstrate that mutation of the ATF/CRE site or deletion of the AP1 site (positions –1745 to –66) in the cyclin D1 promoter reduced the ability of LPA to activate the cyclin D1 promoter. Forced expression of c-Jun and c-Fos increased cyclin D1 expression through both AP1 and ATF/CRE sites (29,30,42,43). In fibroblasts derived from c-Fos-/- fosB-/- mouse embryos that have a defect in proliferation associated with selective reduction of the cyclin D1 level, induction of c-Fos expression restored both cyclin D1 expression and DNA synthesis, indicating pivotal roles for c-Fos and cyclin D1 in cell proliferation (43). c-Jun-/- fibroblasts have reduced levels of cyclins D1 and D3 (44). In this article, we demonstrated that LPA dramatically induces c-Fos and c-Jun proteins in ovarian cancer cells. Maximal stimulatory effects of LPA on the expression of c-Fos observed after 2–4 hours of incubation and on that of c-Jun observed after 4 hours of incubation occurred much earlier than those on cyclin D1 protein observed after 6–18 hours of incubation. Therefore, LPA probably stimulates the expression of cyclin D1, in part, by activating c-Jun and c-Fos, which bind to the AP1 and ATF/CRE sites and then increase the transcription of cyclin D1. Because c-Jun is also required for cyclin D3 expression (44) and because the cyclin D3 promoter region contains AP1 sites (45), LPA likely also induces cyclin D3 expression in ovarian cancer cells to further increase cell proliferation. Because the region of the cyclin D1 promoter from positions –1745 to –66 contains one AP1 site, four SP1 sites, and two STAT (signal transducer and activator of transcription) sites (29,42,46), we do not exclude the possibility that SP1 and STATs are also involved in LPA-stimulated cyclin D1 expression in ovarian cancer cells. Demonstrating this involvement will require further experiments.

Some ovarian cancer cell lines, particularly SKOV3 and CAOV3, expressed EDG7 protein at higher levels than IOSE-29 cells. However, EDG4 protein (in this article) and mRNA (17) were detected at high levels in all ovarian cancer cell lines studied but not in normal OSE or IOSE-29 cells. Forced expression of EDG4 or EDG7 cDNAs caused IOSE-29 cells to become sensitive to LPA stimulation of cyclin D1 promoter-, SRE-, AP1-, and NF{kappa}B-driven gene transcription, with the effect of EDG4 being larger than that of EDG7. Ovarian cancer cells (such as OVCAR-3) are insensitive to apoptosis induced by serum starvation, in contrast to normal and immortalized OSE cells, which are sensitive to such apoptosis. We found that when IOSE-29 cells were transiently transfected with EDG4 or EDG7 cDNAs, the survival rate of the transfected cells in serum-free medium containing 20 µM LPA was substantially greater than that of nontransfected IOSE cells under similar conditions (data not shown). Thus, the expression of EDG4 or EDG7 may suppress apoptosis in ovarian epithelial cells. Recently, cyclin D1 expression was shown to be increased by platelet-derived growth factor, epidermal growth factor, and basic fibroblast growth factor through a phosphatidylinositol 3-kinase (PI3K) pathway (47,48). This PI3K pathway can be activated by LPA, resulting in increased cell proliferation (6). PI3K gene amplification occurs in more than 40% of ovarian cancer cell lines and primary ovarian tumors (49). Thus, LPA also may induce cyclin D1 expression in ovarian cancer cells through the PI3K signaling pathway.

We previously showed (17) that LPA stimulates VEGF expression through transcriptional activation in ovarian cancer cells but not in IOSE-29 cells. In this article, we did not detect the VEGF receptors Flt1 and KDR in the four ovarian cancer cell lines studied, although we detected KDR in normal OSE cells. Because VEGF has little effect on ovarian cancer cell growth in vitro and LPA increases cyclin D1 expression in ovarian cancer cells but not in IOSE-29 cells, dual mechanisms are probably involved in LPA-stimulated ovarian tumor growth in vivo—an indirect endothelial cell-dependent mechanism that involves increasing angiogenesis via VEGF and a direct endothelial cell-independent mechanism that involves increasing cell proliferation via cyclin D1. Thus, blockade of the two distinct LPA mechanisms by LPA antagonist(s) and/or antisense LPA receptor RNA (particularly that for EDG4) may be a useful approach for inhibiting ovarian tumor growth.


    NOTES
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Supported in part by Public Health Service grants P01CA64602 (to R. B. Jaffe) and R01CA70897, R01CA86072, and R01CA75503 (to R. G. Pestell) from the National Cancer Institute, National Institutes of Health, Department of Health and Human Services, and by grants from the Susan G. Komen Breast Cancer Foundation and the Department of Defense (to R. G. Pestell). Y.-L Hu is a John Kerner Scholar in Ovarian Cancer Research. R. G. Pestell is a recipient of the Weil Caulier Irma T. Hirschl Career Scientist award and is the Diane Belfer Faculty Scholar in Cancer Research.

We thank Gordon B. Mills and Robert C. Bast for Hey-A8, OCC1, and DOV-13 ovarian cancer cell lines; Nelly Auersperg for OSE and IOSE-29 cells; and Meng Kian Tee for helpful advice and discussion.


    REFERENCES
 Top
 Notes
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

1 Greenlee RT, Hill-Harmon MB, Murray T, Thun M. Cancer statistics, 2001. CA Cancer J Clin 2001;51:15–36.[Abstract/Free Full Text]

2 Auersperg N, Edelson MI, Mok SC, Johnson SW, Hamilton TC. The biology of ovarian cancer. Semin Oncol 1998;25:281–304.[Medline]

3 Xu Y, Fang XJ, Casey G, Mills GB. Lysophospholipids activate ovarian and breast cancer cells. Biochem J 1995;309:933–40.[Medline]

4 Xu Y, Gaudette DC, Boynton JD, Frankel A, Fang XJ, Sharma A, et al. Characterization of an ovarian cancer activating factor in ascites from ovarian cancer patients. Clin Cancer Res 1995;1:1223–32.[Abstract]

5 Eichholtz T, Jalink K, Fahrenfort I, Moolenaar WH. The bioactive phospholipid lysophosphatidic acid is released from activated platelets. Biochem J 1993;291:677–80.[Medline]

6 Goetzl EJ, An S. Diversity of cellular receptors and functions for the lysophospholipid growth factors lysophosphatidic acid and sphingosine 1-phosphate. FASEB J 1998;12:1589–98.[Abstract/Free Full Text]

7 Moolenaar WH, Kranenburg O, Postma F, Zondag GC. Lysophosphatidic acid: G-protein signaling and cellular responses. Curr Opin Cell Biol 1997;9:168–73.[CrossRef][Medline]

8 An S, Dickens MA, Bleu T, Hallmark OG, Goetzl EJ. Molecular cloning of human Edg2 protein and its identification as a functional cellular receptor for lysophosphatidic acid. Biochem Biophys Res Commun 1997;231:619–22.[CrossRef][Medline]

9 An S, Bleu T, Hallmark OG, Goetzl EJ. Characterization of a novel subtype of human G protein-coupled receptor for lysophosphatidic acid. J Biol Chem 1998;273:7906–10.[Abstract/Free Full Text]

10 Bandoh K, Aoki J, Hosono H, Kobayashi S, Kobayashi T, Murakami-Murofushi K, et al. Molecular cloning and characterization of a novel human G-protein-coupled receptor, Edg7, for lysophosphatidic acid. J Biol Chem 1999;274:27776–85.[Abstract/Free Full Text]

11 Im DS, Heise CE, Harding MA, George SR, O’Dowd BF, Theodorescu D, et al. Molecular cloning and characterization of a lysophosphatidic acid receptor, Edg-7, expressed in prostate. Mol Pharmacol 2000;57:753–9.[Abstract/Free Full Text]

12 Xu Y, Shen Z, Wiper DW, Wu M, Morton RE, Elson P, et al. Lysophosphatidic acid as a potential biomarker for ovarian and other gynecologic cancers. JAMA 1998;280:719–23.[Abstract/Free Full Text]

13 Folkman J, Watson K, Ingber D, Hanahan D. Induction of angiogenesis during the transition from hyperplasia to neoplasia. Nature 1989;339:58–61.[CrossRef][Medline]

14 Ferrara N, Davis-Smyth T. The biology of vascular endothelial growth factor. Endocr Rev 1997;18:4–25.[Abstract/Free Full Text]

15 Bellamy WT, Richter L, Frutiger Y, Grogan TM. Expression of vascular endothelial growth factor and its receptors in hematopoietic malignancies. Cancer Res 1999;59:728–33.[Abstract/Free Full Text]

16 Dias S, Hattori K, Zhu Z, Heissig B, Choy M, Lane W, et al. Autocrine stimulation of VEGFR-2 activates human leukemic cell growth and migration. J Clin Invest 2000;106:511–21.[Abstract/Free Full Text]

17 Hu YL, Tee MK, Goetzl EJ, Auersperg N, Mills GB, Ferrara N, et al. Lysophosphatidic acid induction of vascular endothelial growth factor expression in human ovarian cancer cells. J Natl Cancer Inst 2001;93:762–8.[Abstract/Free Full Text]

18 Yoneda J, Kuniyasu H, Crispens MA, Price JE, Bucana CD, Fidler IJ. Expression of angiogenesis-related genes and progression of human ovarian carcinomas in nude mice. J Natl Cancer Inst 1998;90:447–54.[Abstract/Free Full Text]

19 Goetzl EJ, Dolezalova H, Kong Y, Hu YL, Jaffe RB, Kalli KR, et al. Distinctive expression and functions of the type 4 endothelial differentiation gene-encoded G protein-coupled receptor for lysophosphatidic acid in ovarian cancer. Cancer Res 1999;59:5370–5.[Abstract/Free Full Text]

20 Hill CS, Wynne J, Treisman R. The Rho family GTPases RhoA, Rac1, and CDC42Hs regulate transcriptional activation by SRF. Cell 1995;81:1159–70.[Medline]

21 Miao GG, Curran T. Cell transformation by c-fos requires an extended period of expression and is independent of the cell cycle. Mol Cell Biol 1994;14:4295–310.[Abstract]

22 Wang TC, Cardiff RD, Zukerberg L, Lees E, Arnold A, Schmidt EV. Mammary hyperplasia and carcinoma in MMTV-cyclin D1 transgenic mice. Nature 1994;369:669–71.[CrossRef][Medline]

23 Bodrug SE, Warner BJ, Bath ML, Lindeman GJ, Harris AW, Adams JM. Cyclin D1 transgene impedes lymphocyte maturation and collaborates in lymphomagenesis with the myc gene. EMBO J 1994;13:2124–30.[Abstract]

24 Pestell RG, Albanese C, Reutens AT, Segall JE, Lee RJ, Arnold A. The cyclins and cyclin-dependent kinase inhibitors in hormonal regulation of proliferation and differentiation. Endocr Rev 1999;20:501–34.[Abstract/Free Full Text]

25 Worsley SD, Ponder BA, Davies BR. Overexpression of cyclin D1 in epithelial ovarian cancers. Gynecol Oncol 1997;64:189–95.[CrossRef][Medline]

26 Kornmann M, Arber N, Korc M. Inhibition of basal and mitogen-stimulated pancreatic cancer cell growth by cyclin D1 antisense is associated with loss of tumorigenicity and potentiation of cytotoxicity to cisplatinum. J Clin Invest 1998;101:344–52.[Abstract/Free Full Text]

27 Uto H, Ido A, Moriuchi A, Onaga Y, Nagata K, Onaga M, et al. Transduction of antisense cyclin D1 using two-step gene transfer inhibits the growth of rat hepatoma cells. Cancer Res 2001;61:4779–83.[Abstract/Free Full Text]

28 Lee RJ, Albanese C, Fu M, D’Amico M, Lin B, Watanabe G, et al. Cyclin D1 is required for transformation by activated Neu and is induced through an E2F-dependent signaling pathway. Mol Cell Biol 2000;20:672–83.[Abstract/Free Full Text]

29 Albanese C, Johnson J, Watanabe G, Eklund N, Vu D, Arnold A, et al. Transforming p21ras mutants and c-Ets-2 activate the cyclin D1 promoter through distinguishable regions. J Biol Chem 1995;270:23589–97.[Abstract/Free Full Text]

30 Watanabe G, Howe A, Lee RJ, Albanese C, Shu IW, Karnezis AN, et al. Induction of cyclin D1 by simian virus 40 small tumor antigen. Proc Natl Acad Sci U S A 1996;93:12861–6.[Abstract/Free Full Text]

31 Guttridge DC, Albanese C, Reuther JY, Pestell RG, Baldwin AS Jr. NF-kappa B controls cell growth and differentiation through transcriptional regulation of cyclin D1. Mol Cell Biol 1999;19:5785–99.[Abstract/Free Full Text]

32 Van Antwerp DJ, Martin SJ, Kafri T, Green DR, Verma IM. Suppression of TNF-alpha-induced apoptosis by NF-kappa B. Science 1996;274:787–9.[Abstract/Free Full Text]

33 Hai T, Curran T. Cross-family dimerization of transcription factors Fos/Jun and ATF/CREB alters DNA binding specificity. Proc Natl Acad Sci U S A 1991;88:3720–4.[Abstract]

34 Shen Z, Belinson J, Morton RE, Xu Y, Xu Y. Phorbol 12-myristate 13-acetate stimulates lysophosphatidic acid secretion from ovarian and cervical cancer cells but not from breast or leukemia cells. Gynecol Oncol 1998;71:364–8.[CrossRef][Medline]

35 Bartek J, Bartkova J, Lukas J. The retinoblastoma protein pathway in cell cycle control and cancer. Exp Cell Res 1997;237:1–6.[CrossRef][Medline]

36 Todd MC, Sclafani RA, Langan TA. Ovarian cancer cells that coexpress endogenous Rb and p16 are insensitive to overexpression of functional p16 protein. Oncogene 2000;19:258–64.[CrossRef][Medline]

37 Fang X, Jin X, Xu HJ, Liu L, Peng HQ, Hogg D, et al. Expression of p16 induces transcriptional downregulation of the RB gene. Oncogene 1998;16:1–8.[CrossRef][Medline]

38 Sherr CJ, Roberts JM. CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev 1999;13:1501–12.[Free Full Text]

39 Yamamoto Y, Gaynor RB. Therapeutic potential of inhibition of the NF-kappa B pathway in the treatment of inflammation and cancer. J Clin Invest 2001;107:135–42.[Free Full Text]

40 Shahrestanifar M, Fan X, Manning DR. Lysophosphatidic acid activates NF-kappa B in fibroblasts. A requirement for multiple inputs. J Biol Chem 1999;274:3828–33.[Abstract/Free Full Text]

41 Huang S, Robinson JB, Deguzman A, Bucana CD, Fidler IJ. Blockade of nuclear factor-kappa B signaling inhibits angiogenesis and tumorigenicity in human ovarian cancer cells by suppressing expression of vascular endothelial growth factor and interleukin-8. Cancer Res 2000;60:5334–9.[Abstract/Free Full Text]

42 Watanabe G, Lee RJ, Albanese C, Rainey WE, Batlle D, Pestell RG. Angiotensin II activation of cyclin D1-dependent kinase activity. J Biol Chem 1996;271:22570–7.[Abstract/Free Full Text]

43 Brown JR, Nigh E, Lee RJ, Ye H, Thompson MA, Saudou F, et al. Fos family members induce cell cycle entry by activating cyclin D1. Mol Cell Biol 1998;18:5609–19.[Abstract/Free Full Text]

44 Wisdom R, Johnson RS, Moore C. c-Jun regulates cell cycle progression and apoptosis by distinct mechanisms. EMBO J 1999;18:188–97.[Abstract/Free Full Text]

45 Wang Z, Sicinski P, Weinberg RA, Zhang Y, Ravid K. Characterization of the mouse cyclin D3 gene: exon/intron organization and promoter activity. Genomics 1996;35:156–63.[CrossRef][Medline]

46 Matsumura I, Kitamura T, Wakao H, Tanaka H, Hashimoto K, Albanese C, et al. Transcriptional regulation of the cyclin D1 promoter by STAT5: its involvement in cytokine-dependent growth of hematopoietic cells. EMBO J 1999;18:1367–77.[Abstract/Free Full Text]

47 Page K, Li J, Wang Y, Kartha S, Pestell RG, Hershenson MB. Regulation of cyclin D1 expression and DNA synthesis by phosphatidylinositol 3-kinase in airway smooth muscle cells. Am J Respir Cell Mol Biol 2000;23:436–43.[Abstract/Free Full Text]

48 Takuwa N, Fukui Y, Takuwa Y. Cyclin D1 expression mediated by phosphatidylinositol 3-kinase through mTOR-p70 (S6K)-independent signaling in growth factor-stimulated NIH 3T3 fibroblasts. Mol Cell Biol 1999;19:1346–58.[Abstract/Free Full Text]

49 Shayesteh L, Lu Y, Kuo WL, Baldocchi R, Godfrey T, Collins C, et al. PIK3CA is implicated as an oncogene in ovarian cancer. Nat Genet 1999;21:99–102.[CrossRef][Medline]

Manuscript received July 19, 2002; revised March 7, 2003; accepted March 19, 2003.


This article has been cited by other articles in HighWire Press-hosted journals:


             
Copyright © 2003 Oxford University Press (unless otherwise stated)
Oxford University Press Privacy Policy and Legal Statement