- Open Access
Lipid phosphate phosphatase-3 regulates tumor growth via β-catenin and Cyclin-D1 signaling
© Chatterjee et al; licensee BioMed Central Ltd. 2011
- Received: 4 February 2011
- Accepted: 11 May 2011
- Published: 11 May 2011
The acquisition of proliferative and invasive phenotypes is considered a hallmark of neoplastic transformation; however, the underlying mechanisms are less well known. Lipid phosphate phosphatase-3 (LPP3) not only catalyzes the dephosphorylation of the bioactive lipid sphingosine-1-phosphate (S1P) to generate sphingosine but also may regulate embryonic development and angiogenesis via the Wnt pathway. The goal of this study was to determine the role of LPP3 in tumor cells.
We observed increased expression of LPP3 in glioblastoma primary tumors and in U87 and U118 glioblastoma cell lines. We demonstrate that LPP3-knockdown inhibited both U87 and U118 glioblastoma cell proliferation in culture and tumor growth in xenograft assays. Biochemical experiments provided evidence that LPP3-knockdown reduced β-catenin, CYCLIN-D1, and CD133 expression, with a concomitant increase in phosphorylated β-catenin. In a converse experiment, the forced expression of LPP3 in human colon tumor (SW480) cells potentiated tumor growth via increased β-catenin stability and CYCLIN-D1 synthesis. In contrast, elevated expression of LPP3 had no tumorigenic effects on primary cells.
These results demonstrate for the first time an unexpected role of LPP3 in regulating glioblastoma progression by amplifying β-catenin and CYCLIN-D1 activities.
- Vascular Endothelial Growth Factor
- U118 Cell
- SW480 Cell
- Athymic Nude Mouse
The lipid phosphate phosphatases (LPPs) have been shown to dephosphorylate sphingosine 1-phosphate (S1P) and its structural homologs to produce metabolic products such as sphingosine, ceramide, and lysophosphatidic acid (LPA) [1–3]. Three major isoforms, LPP1, LPP2, and LPP3 have been studied in various tissues and cell lines [3, 4], however, the role of LPPs in tumor progression is not well understood [1–4]. Although LPPs are localized to the endoplasmic reticulum (ER), cytoplasm and plasma membrane [5, 6], LPPs also associate with membrane microdomain rafts and caveolae [5–7]. Ovarian carcinoma cells secrete a high concentration of S1P and LPA [8, 9]. Accordingly, overexpressing sphingosine kinase 1 (Sphk1), one of the main enzymes that catalyzes the formation of S1P, results in increased tumorigenic activity in the Min mouse model of intestinal tumor progression . Additionally, elevated expression of LPPs has been postulated to reduce the level of S1P and its structural homologs, thereby down-regulating cell signaling. In support of this notion, in vitro experiments have provided evidence for the ability of LPPs to down-regulate the activity of extracellular signal-regulated kinase (Erk)1/2 and induce death of ovarian carcinoma cells [11, 12]. However, LPP3 has been shown to be highly expressed in sprouting endothelial cells and can act as a cell associated integrin ligand [13–15]. Accordingly, we have shown that an antibody against the extracellular domain of LPP3 inhibits cell-cell interactions and angiogenesis in vitro[14–16]. In a separate study, we have also shown that LPP3 is required for tumor adaptation and β-catenin signaling [17, 18]. These observations provided us with the initial impetus to carry out this study on the role of LPPs in the acquisition of neoplastic cellular behavior.
In addition to binding to E-cadherin, a key role for β-catenin has been established in Wnt (wingless) signaling. The activation of canonical Wnt signaling promotes the translocation of β- catenin to the nucleus, where it acts as a co-activator for the transcription factor T-cell factor/lymphocyte enhancer binding factor (TCF/LEF-1) complex. Increased Wnt/β-catenin signaling has been linked to cancer cell proliferation, stem cell self-renewal, and angiogenesis. In this regard, β-catenin signaling is often shown to up-regulate transcription of the CYCLIN-D1 protein. CYCLIN-D1 is a key regulator of cell cycle progression, the increased expression of which is a fundamental characteristic of tumor cell proliferation. Although Lpp3 is essential for vasculogenesis, aspects of embryonic development and Wnt signaling , the function of LPP3 during pathological processes, including tumor growth, remains unknown. A few reports have focused on the role of LPPs in angiogenesis [14, 16–19], but none has reported the expression patterns of LPP3 in human primary tumors or tumor cell lines. Importantly, the association of increased LPP3 expression with cell transformation and tumorigenesis has not been reported. To test whether LPP3 regulates tumor cell behavior, we knocked down LPP3 and investigated glioblastoma cell proliferation and tumor formation; conversely, we forced expression of LPP3 in LPP3-deficient human colon carcinoma (SW480) cells to address the hypothesis that LPP3 potentiates cell proliferation and tumor growth.
Expression of LPP3 Protein in a Subset of Primary Tumors
LPP3-Knockdown Reduces Glioblastoma Tumor Cell Proliferation and Migration
LPP3 Knockdown Reduced U87 and U118 Glioblastoma Tumor Growth
Elevated Expression of LPP3 Potentiates SW480 Growth in Xenograft Assay
Both the RGD and the Lipid Phosphatase Domains of LPP3 are required for SW480 Tumor Growth
In this report we showed that LPP3 was constitutively expressed in a subset of primary tumors and in U87 and U118 cells, whereas it was undetectable in a human adenocarcinoma E-cadherin deficient SW480 cell line. Because Wnt/β-catenin signaling is often constitutively active in many neoplastic cells, we posited that constitutive expression of LPP3 in glioblastoma cells may serve as a link in the acquisition of proliferative, invasive, and metastatic phenotypes. Glioblastoma is an aggressive type of brain tumor . The lethal behavior of glioblastoma is characterized by its ability to invade quickly, usually throughout the normal brain parenchyma, and to evade apoptosis, thereby rendering it incurable with traditional chemotherapy and radiation [21, 22]. In this regard, neural stem cells have been shown to differentiate into endothelial cells. In fact, glioblastoma cells might have the ability to transition into endothelial-like cells [23–25]. Although LPP3 function is restricted to endothelial cells, LPP3 could have neural function as well. Thus, herein we addressed the relationship between increased LPP3 expression and glioblastoma tumor growth in vitro and in vivo.
Increased proliferation and migration are key hallmarks of tumor cells. Beyond binding to VE- or E-cadherin, β-catenin transduces Wnt signaling to serve as a transcriptional co-activator that alters the expression of target genes involved in cell proliferation, survival, and differentiation. CYCLIN-D1 is one of the key targets of the Wnt/β-catenin/LEF-1 transcriptional machinery that induces the progression of cells through the G1 phase of the cell cycle. β-catenin and CYCLIN-D1 are major regulators of tumor cell proliferation and migration. LPP3 depletion reduced β-catenin and CYCLIN-D1 proteins, thereby impairing U87 and U118 glioblastoma cell proliferation and migration. Reduction of β-catenin species may have also been due to the increased phosphorylation of β-catenin in these cells. These results strongly suggest that LPP3 knockdown impedes cell proliferation and migration by reducing β-catenin and CYCLIN-D1 levels.
Xenograft experiments demonstrate that LPP3 knockdown decreased the levels of not only β-catenin and CYCLIN-D1, but also CD133 (a marker of stem-like cells), VEGF and IL-8. Expression of CD133 by U87 and U118 cells is consistent with recent reports [23–27]; notably, it has been proposed that CD133+ glioblastoma tumor cells give rise to tumor endothelium cells [23, 24]. Therefore, our results suggest a key role of LPP3 in the generation of CD133+ cells. VEGF and IL-8 are a pro-angiogenic growth factor and cytokine, respectively. Because VEGF and IL-8 are activated by β-catenin signaling , it was not surprising that U87 and U118 glioblastoma cells expressed high levels of these two angiogenic factors . Importantly, LPP3 knockdown significantly decreased levels of VEGF and IL-8, but not of TIMP-2, in these cells. Whether LPP3 knockdown affects differentiation of glioblastoma tumor cells into tumor endothelium will require detailed investigation. However, our data suggest that LPP3 knockdown also affects the level of CD133 protein and secretion of factors that are required for tumor neovascularization.
In the converse experiment, we determined the impact of elevated LPP3 expression in LPP3-deficient SW480 colon tumor cells and performed a xenograft assay using athymic nude mice. In this regard, tumorigenic SW480 cells harbor two mutant alleles of Adenomatous Polyposis Coli (APCmut/mut) gene, but normal β-catenin. Accordingly, we have demonstrated that the forced expression of hLPP3 and mLpp3 in SW480 cells potentiated tumor growth and metastasis in the xenograft assay, whereas hLPP1 and hLPP2 had no effect. Importantly, the forced expression of LPP3 decreased the basal phosphorylation status of β-catenin, thereby enhancing the stability of β-catenin species. Altogether, these data suggest that the increased proliferation and tumor growth are LPP3-specific and mediated by increased β-catenin and CYCLIN-D1 functional activity.
To define the minimal functional domains of LPP3 involved in the potentiation of tumor growth, we generated retroviral constructs carrying functional mutations. Stable clones of SW480 cells expressing these constructs were implanted in nude mice. hLPP3 induced robust tumor growth, whereas hLPP3-RAD or the hLPP3-PI supported SW480 tumor growth at a diminished rate. However, the tumor volume was significantly reduced (as in control cells) in mice receiving SW480 cells expressing the hLPP3-RAD + PI construct. These data suggest that the LPP3 protein has at least two functional domains, the lipid phosphatase and the LPP3-RGD cell adhesion domains. The forced expression of LPP3 in NIH-3T3 fibroblast cells did not transform these cells (data not shown), suggesting that the LPP3 is not tumorigenic, but in combination with genetic instability such as the mutation of APC, elevated expression of LPP3 potentiates tumor progression by stimulating β-catenin and CYCLIN-D1 transcriptional activity.
One long-held view is that LPP3 down-regulates cell signaling by virtue of its ability to dephosphorylate S1P and its structural homologs. In contrast, our data show the ability of LPP3 to potentiate tumor growth by amplifying β-catenin and CYCLIN-D1 activities. Therefore, these unexpected set of results implicate LPP3 as a potential target for inhibiting the growth of glioblastoma and perhaps other tumors that express high levels of LPP3.
Cells and Reagents
Human glioblastoma U87 and U118 cell lines were purchased from the American Type Culture Collection (Manassas, VA). E-cadherin-deficient (ECD) human colon adenocarcinoma SW480 (APCmut/mut) cells were a kind gift of Dr. Cara Gottardi (Northwestern University, Chicago, IL). Preparation of rabbit anti-LPP3 polyclonal antibodies (pAbs) has been described previously [14, 15]. Mouse anti-VCIP/LPP3 (39-1000) monoclonal (mAb) and anti-LPP2 pAb were purchased from Invitrogen (Carlsbad, CA) and Exalpha Biologicals, Inc (Shirley, MA), respectively. Anti-CYCLIN-D1 (2978), rabbit anti-phospho-β-catenin (Ser33, Ser37, and Thr41) and rabbit anti-CD133 polyclonal antibodies were purchased from Cell Signaling Technology, Inc. (Denver, MA). Anti-β-catenin (SC-7963), anti-GAPDH (SC-51906) and anti-GRB2 pAbs were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Secondary antibodies were purchased from Promega (Madison, WI).
Glioblastoma tissue arrays were purchased from U.S. Biomax Inc. (Rockville, MD). Paraffin-embedded tissues were collected from normal pancreas and 63 pancreatic endocrine tumor (PET) affected patients resected at the University Hospital of Verona, Italy. Informed consent for the analysis of normal and neoplastic tissues was obtained from patients in accordance with the approval of the Verona University and Hospital Trust's Ethical Committee. Four-micrometer-thick sections were prepared. Antigen retrieval was performed in 10 mM citrate buffer (pH 6.0) and heated in a microwave at 360 W for 20 min. Endogenous peroxidase and non-specific sites were blocked by incubation, respectively, with 3% H2O2 and 10% goat serum at room temperature. Sections were then incubated with a rabbit IgG or a rabbit anti-LPP3-RGD antibody (5 μg/ml) for 1 h and 30 min at room temperature, washed three times with PBST (PBS/0.2% Tween-20), and incubated for 30 min with anti-rabbit conjugated peroxidase (DakoEnVision+®, Peroxidase, Mouse and Rabbit Ready-to-use). After 3 washes with PBST, diaminobenzidine (DAB) (Vector Biolaboratories, SK-4100) was used as a peroxidase substrate for 10 minutes at room temperature.
Recombinant cDNA constructs, cell culture, retroviral transduction, and stable cell line generation
pLNCX2 and the amphotropic packaging cell line, HEK293 (human kidney fibroblasts), were purchased from BD Biosciences (San Jose, CA). Generation of recombinant cDNA constructs in pLNCX2 retroviruses have been described previously [14, 15, 18]. Control and LPP shRNA retroviral constructs were purchased from Origene (Rockville, MD). All transfection experiments have been previously described [14, 15, 18]. Stable clones were selected by western blotting, and LPP expression levels were determined by immunoblotting as described previously [14, 15, 18].
Cell extraction and immunoprecipitation
Cell extracts were prepared using 20 mM Tris (pH 7.5), 150 mM NaCl, 1% Triton X-100, 0.25% NP-40, and 1 mM EDTA buffer. All biochemical experiments, including western blot (WB) analyses, were performed as described previously [14, 15, 18, 28]
Cell migration assay
Migration assay was carried out as described previously . In brief, modified transwell Boyden chambers (8 μM) were used for chemotactic cell migration. Human U87 and U118 cell were plated overnight in complete media, next day cells at 60% density were infected with control- or LPP3-shRNA retroviral particles for 12 hours. Media containing viral particles were removed and replenished with media with containing insulin, transferrin and selenium (hereafter called Migration Assay Media, MAM) without serum. After 4 hours, cells detached with 2 mM EDTA, pH 7.4, pelleted, washed twice with PBS (pH 7.5), and resuspended in MAM. The upper chamber was filled with 400 μl of MAM containing ~1 × 104 cells, and the lower chamber was filled with 500 μl of MAM, no chemo-attractant was used. Cells were incubated for 6 h at 37°C in a CO2 incubator. After 6 hours, inserts were retrieved and cells in the upper chamber removed with q-tips, while cells migrated to the lower face of the membrane were fixed and stained with 0.5% crystal violet. Excess dye was washed with running tap water, membranes were then examined using a phase-contrast microscope. At least seven 200 × magnification fields were selected for scoring. Cell migration assay was repeated 3 times with each trial being performed in triplicate.
Enzyme-linked Immunosorbent Assay (ELISA)
ELISAs were performed as described previously [18, 28]. To monitor the secretion of VEGF, IL-8, and TIMP-2, U87 and U118 cell lines were left in medium containing 5 μg/ml puromycin for 24 hours. Then, the cell culture media was collected, centrifuged and subjected to ELISA according to the manufacturer's recommendations (Research Diagnostics and R & D Systems). The ELISA detection limit for each factor ranged from 6 pg/ml to 1600 pg/ml, and the intra- and inter-assay variations were 4.1-6.2% and 6.5-10%, respectively.
All experiments described in this study were carried out according to the University of Illinois (UIC) Animal Care Committee (ACC) and NIH guidelines. Eight-week old female (~25 g) athymic nude mice from Jackson Laboratory (Bar Harbor, ME) were used for this study. SW480 cells expressing vector (pLNCX2) alone and various LPP constructs were trypsinized, resuspended and washed with sterile PBS, pH 7.4. After resuspension in PBS, pH 7.4, cells were passed through a 100 μM cell strainer and enumerated using a hemocytometer. Cells (2 × 104 in 150 μl) were subcutaneously (s.c.) injected into each of 12 animals (n = 12) per group. Tumor growth was monitored for 21 or 28 days. One mouse from each group was selected randomly, anesthetized, photographed, and sacrificed and the tissues were recovered for further analysis.
Analysis of variance (ANOVA) was used for statistical analyses and values of P < 0.05 were considered statistically significant. Data represent means ± s.e.m. All analyses were performed using GraphPad Prizm 5.0 software (La Jolla, CA).
The authors thank John Kim for technical assistance. We acknowledge the contribution of Prof. Aldo Scarpa and Dr. Stefania Beghelli for providing the tissue samples and Dr. Marzia Vezzalini (Dept of Pathology and Diagnostics, University of Verona, Verona, Italy) for her skillful technical assistance. Studies were supported by National Institutes of Health grants (R01HL079356; HL079356-03S1) and by the University of Illinois at Chicago (UIC) Center for Clinical and Translational Science (CCTS) Award Number UL1RR029879, from the National Center for Research Resources, to K.K.W. E.E.K. was supported by National Institutes of Health T32GM070388 and T32HL072742 Training grants.
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