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Molecular Cancer Research 5, 1201-1211, November 1, 2007. doi: 10.1158/1541-7786.MCR-06-0338
© 2007 American Association for Cancer Research
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Signaling and Regulation

Lysophosphatidic Acid Decreases the Nuclear Localization and Cellular Abundance of the p53 Tumor Suppressor in A549 Lung Carcinoma Cells

Mandi M. Murph1, Jennifer Hurst-Kennedy1, Victoria Newton1, David N. Brindley2 and Harish Radhakrishna1

1 School of Biology, Georgia Institute of Technology, Atlanta, Georgia and 2 Department of Biochemistry (Signal Transduction Research Group), University of Alberta, Edmonton, Alberta, Canada

Requests for reprints: Harish Radhakrishna, The Coca-Cola Company, One Coca-Cola Plaza, TEC-437, Atlanta, GA 30301. Phone: 404-676-4801; Fax: 404-515-5112. E-mail: hradha{at}earthlink.net


   Abstract
Top
 Notes
 Abstract
Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Lysophosphatidic acid (LPA) is a bioactive lipid that promotes cancer cell proliferation and motility through activation of cell surface G protein–coupled receptors. Here, we provide the first evidence that LPA reduces the cellular abundance of the tumor suppressor p53 in A549 lung carcinoma cells, which express endogenous LPA receptors. The LPA effect depends on increased proteasomal degradation of p53 and it results in a corresponding decrease in p53-mediated transcription. Inhibition of phosphatidylinositol 3-kinase protected cells from the LPA-induced reduction of p53, which implicates this signaling pathway in the mechanism of LPA-induced loss of p53. LPA partially protected A549 cells from actinomycin D induction of both apoptosis and increased p53 abundance. Expression of LPA1, LPA2, and LPA3 receptors in HepG2 hepatoma cells, which normally do not respond to LPA, also decreased p53 expression and p53-dependent transcription. In contrast, neither inactive LPA1 (R124A) nor another Gi-coupled receptor, the M2 muscarinic acetylcholine receptor, reduced p53-dependent transcription in HepG2 cells. These results identify p53 as a target of LPA action and provide a new dimension for understanding how LPA stimulates cancer cell division, protects against apoptosis, and thereby promotes tumor progression. (Mol Cancer Res 2007;5(11):1201–11)


   Introduction
Top
 Notes
 Abstract
 Introduction
Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Lysophosphatidic acid (LPA) is a bioactive lipid found in body fluids that influences a wide range of processes, including cell proliferation, migration, morphologic changes, survival, and neointimal formation (1-7). LPA is produced in blood by activated platelets to facilitate wound healing (8, 9) and it is also produced by a variety of cancer cells (10, 11). The diverse actions of LPA are primarily mediated by three seven-transmembrane, G protein–coupled receptors, LPA1/Edg2, LPA2/Edg4, and LPA3/Edg7, and possibly also by the metabolic receptor peroxisome proliferator-activated receptor-{gamma} (12-16). Recent studies also indicate that the orphan receptors GPR23 and GPR92 are high-affinity LPA receptors (15, 17). The LPA-binding G protein–coupled receptors can collectively activate Gi, Gs, Gq, and G12/13 (15, 18, 19). The resultant effects of LPA are mediated through the activation of downstream signaling pathways controlled by these G proteins (20).

LPA potently stimulates the growth, survival, and motility of a variety of cancer cells, some of which can themselves produce LPA (20). One mechanism for this is through the secretion of autotaxin, a secreted lysophospholipase D, which produces LPA from extracellular lysophosphatidylcholine (21, 22). Autotaxin is well known to be involved in promoting tumor development, metastasis, and angiogenesis and its effects can be explained by the extracellular production of LPA. The levels of LPA are high in ascites fluid and plasma of patients with ovarian tumors (23). LPA protects against apoptosis caused by chemotherapeutic agents (3). It promotes ovarian tumor development, possibly involving increased cyclin D expression (24). LPA also increases vascular endothelial growth factor production in some cancer cells, which stimulates angiogenesis (25). In a colon cancer cell line, LPA increases the synthesis of macrophage migration inhibitory factor, which promotes tumor growth (26). LPA levels are elevated in the blood of patients with multiple myeloma (27).

LPA signaling leads to the activation of both cell proliferative signaling pathways, such as the Ras/Raf/extracellular signal-regulated kinase pathway, and cell survival pathways, such as those involving phosphatidylinositol 3-kinase (PI3K) and Akt (28, 29). These effectors are primarily activated via pertussis toxin-sensitive, Gi signaling. Although much is known about the effects of LPA signaling on cancer cell growth and motility, relatively little is known about the effects of LPA on cell cycle regulators, such as the p53 tumor suppressor. p53 is a transcription factor that controls the expression of genes encoding proteins that regulate apoptosis and cell cycle progression (30-32). p53 is inactivated in ~50% of all cancers (33).

Given the strong mitogenic effects of LPA and the commonly observed loss of p53 function in cancer cells, we investigated the effects of LPA on the function of the p53 tumor suppressor. We found that signaling initiated by the LPA receptors (LPA1, LPA2, and LPA3) potently inhibited p53-dependent transcription, promoted the loss of p53 protein, and protected A549 lung tumor cells from actinomycin D–induced apoptosis. These results show that p53 is a target for the actions of LPA and they provide a new dimension for understanding how LPA promotes tumor growth.


   Results
Top
 Notes
 Abstract
 Introduction
 Results
Discussion
 Materials and Methods
 Acknowledgements
 References
 
LPA Decreases p53 Abundance in A549 Cells
We first determined how the bioactive lipid LPA affected the cellular distribution of p53 protein in A549 human lung carcinoma cells because these cells proliferate rapidly, express wild-type p53, and contain endogenous LPA receptors (34, 35). p53 was localized by using mouse anti-p53 antibody (DO-1) and indirect immunofluorescence; the nucleus was visualized by staining with Hoechst dye. In the majority of A549 cells (~70%), p53 was localized in the nucleus and, to a lesser extent, in a diffuse cytoplasmic pattern (Fig. 1A ). The nuclear p53 labeling in these cells was bright; thus, we will refer to these cells as p53NUC-bright cells. In contrast, p53 staining was greatly reduced or completely absent from the nucleus in ~30% of control A549 cells (Fig. 1A, arrows and Fig. 2B , SFM); no detectable difference was observed in either the fluorescence intensity or distribution of p53 in the cytoplasm of these "nuclear p53-diminished" cells (we will henceforth refer to these cells as p53NUC-diminished cells). We quantified the fluorescence intensity of p53 staining in the nucleus of these two populations of cells by using MetaMorph image analysis software and normalized it to the fluorescence intensity of DNA labeled with Hoechst dye. In cells exhibiting bright nuclear p53 staining, the nuclear fluorescence intensity of p53 was 2.2-fold greater than that observed in cells exhibiting reduced p53 staining (p53NUC-bright cells: fluorescence intensity = 0.83 ± 0.06, n = 15 cells; p53NUC-diminished cells: fluorescence intensity = 0.38 ± 0.03, n = 15 cells).


Figure 1
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  		FIGURE 1. LPA stimulation reduces the nuclear localization and cellular abundance of p53 in A549 cells. A. Localization of endogenous p53 in A549 lung carcinoma cells. Endogenous p53 was localized in A549 cells using mouse anti-p53 antibody and fluorescently labeled anti-mouse secondary antibodies as described in Materials and Methods. DNA was labeled with Hoechst dye. Arrows, cells that displayed reduced nuclear staining of p53, which are referred in the text as p53NUC-diminished cells. B. LPA dose dependence on loss of nuclear p53 localization. A549 cells grown on glass coverslips were serum starved for 16 h before treatment with the indicated concentrations of LPA for 6 h. Total p53 was localized by indirect immunofluorescence microscopy using mouse anti-p53 (DO-1) antibody followed by fluorescently labeled secondary antibody. The fluorescence intensity of nuclear p53 staining was normalized to DNA labeled with Hoechst dye and quantified by MetaMorph. Points, mean of the fluorescence intensity of nuclear p53 in LPA-treated cells relative to that observed in untreated cells and are from a representative experiment that was repeated at least thrice with similar results; bars, SE. C. Cellular abundance of p53. A549 cells were treated without LPA (Untreated) or with 10 µmol/L LPA for 6 h before immunoblotting of whole-cell extracts with mouse anti-p53 antibodies or mouse anti-paxillin antibodies. The protein band intensities were quantified by MetaMorph image analysis following background subtraction and normalized to the untreated samples as described in Materials and Methods. The relative band intensities are indicated below each band. D. Cellular abundance of p53 in serum-replete versus serum-starved cells. A549 cells were grown in the presence of serum or serum starved for 12 h before Western blotting. A parallel sample of serum-replete cells was incubated with 10 µmol/L LPA for 6 h before Western blotting of whole-cell extracts with mouse anti-p53 antibodies or mouse anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antibodies. The protein band intensities were quantified by MetaMorph image analysis following background subtraction and normalized to the untreated serum-starved samples as described in Materials and Methods. The relative band intensities are indicated below each band.

 

Figure 2
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  		FIGURE 2. The LPA-induced reduction of nuclear p53 is inhibited by lactacystin. A. Population effect of LPA on nuclear p53 localization. Serum-starved A549 cells were incubated in the presence or absence of 10 µmol/L LPA for 6 h before indirect immunofluorescence localization of p53. Note the marked reduction in the nuclear p53 staining intensity in LPA-treated cells. B. Quantification of p53NUC-bright cells. Serum-starved A549 cells were stimulated for 6 h with SFM, 10 µmol/L LPA, 1 µmol/L OMPT, 2 µg/mL actinomycin D (ActD), 20 µmol/L lactacystin, or lactacystin and LPA before the localization of p53 by indirect immunofluorescence of triplicate samples. One hundred cells per sample were randomly selected and scored as having either bright or diminished nuclear p53 staining. Columns, mean percentage of p53NUC-bright cells; bars, SE. **, P < 0.01, compared with control SFM cells.

 
To investigate the LPA dose dependence of the loss of nuclear p53, A549 cells were incubated with various concentrations of LPA for 6 h and the nuclear p53 staining intensity was quantified and normalized to the fluorescence intensity of DNA labeled with Hoechst dye. We observed a progressive reduction in nuclear p53 labeling as the concentration of LPA was increased through the physiologic range. Reduction in nuclear p53 staining was first apparent at 0.01 µmol/L and was the greatest at 10 µmol/L LPA (~50% of control). Western blotting of whole-cell extracts also showed that treatment with 10 µmol/L LPA for 6 h in serum-free medium (SFM) reduced the total cellular abundance of p53 by ~50% (Fig. 1C and D). In contrast, LPA treatment did not affect the cellular abundance of the focal adhesion protein paxillin, which was used as a loading control. LPA is present in serum at concentrations ranging from 0.1 to 10 µmol/L (8). We next compared the abundance of p53 in serum-deprived and serum-replete cells and asked whether LPA would decrease p53 in serum-replete cells (Fig. 1D). Although the abundance of p53 was increased in serum-deprived cells (~40% greater), relative to serum-replete cells, LPA still reduced p53 abundance in serum-replete cells. Taken together, these results indicated that LPA reduced the nuclear localization and total cellular abundance of the p53 tumor suppressor in A549 cells.

In addition to a reduction in the nuclear localization of p53 within a given cell, we observed that LPA also decreased the proportion of p53NUC-bright cells (Fig. 2A). We quantified the proportion of p53NUC-bright cells in control and LPA-treated samples and found that treatment of A549 cells with a physiologic concentration of 10 µmol/L LPA for 6 h decreased the proportion of p53NUC-bright cells by ~35% relative to control cells (Fig. 2B, 10 µmol/L LPA). Treatment of A549 cells with the LPA3-selective agonist (2S)-1-oleoyl-2-O-methyl-glycero-3-phosphothionate (OMPT; ref. 36) decreased the proportion of p53NUC-bright cells, even more than LPA, by ~60%, relative to control cells (Fig. 2B, 1 µmol/L OMPT). Genotoxic drugs, such as the transcription inhibitor actinomycin D, enhance the nuclear accumulation of p53, which ultimately induces apoptosis in cells (37-39). Treatment of A549 cells with 2 µg/mL actinomycin D indeed increased the proportion of p53NUC-bright cells relative to untreated cells (Fig. 2B).

The rapid degradation of p53 in nontransformed cells is facilitated by ubiquitin- and proteosome-mediated degradation, which can occur both in the nucleus and in the cytoplasm (40-42). We next tested the effects of inhibiting proteasomal degradation on the proportion of p53NUC-bright cells by using the proteasomal inhibitor lactacystin (20 µmol/L). Lactacystin has been shown to inhibit nuclear proteasomal degradation (43, 44). Treatment of cells with lactacystin alone also increased the proportion of p53NUC-bright cells relative to untreated cells (Fig. 2B). More importantly, incubation of A549 cells with both lactacystin (20 µmol/L) and LPA (10 µmol/L) prevented the LPA-induced decrease in p53NUC-bright cells (Fig. 2B). These results indicate that the LPA-induced reduction of p53 in A549 cells is likely mediated by ubiquitin- and proteasomal-induced degradation.

Activation of LPA Receptors Inhibits p53-Stimulated Transcription
A major function of p53 in cells is to stimulate the transcription of proteins involved in DNA repair, apoptosis, or cell cycle arrest (45). To investigate whether LPA stimulation resulted in the reduction of an endogenous p53-regulated gene product, we tested the effects of 10 µmol/L LPA on the cellular distribution of the cyclin D inhibitor p21CIP1. Activated p53 stimulates the transcription of the cyclin inhibitor p21CIP1, which in turn promotes G1 cell cycle arrest (46). In untreated cells, both p53 and p21CIP1 were predominantly localized to the nucleus (Fig. 3A ). Approximately 50% of all cells in the population showed p21CIP1 labeling in the nucleus (Fig. 3B). Treatment with LPA (10 µmol/L) reduced both p53 and p21CIP1 labeling and decreased the proportion of cells showing nuclear p21CIP1 staining to 25% of the total population (Fig. 3B). In contrast, treatment of A549 cells with a genotoxic agent, doxorubicin (1 µg/mL), enhanced the nuclear staining of both p53 and p21CIP1 (Fig. 3A). Doxorubicin significantly increased the proportion of cells showing p21CIP1 staining in the nucleus to >85% of the population (Fig. 3B).


Figure 3
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  		FIGURE 3. LPA stimulation decreases the abundance of p21CIP1. A. Localization of p21CIP1. A549 cells were serum starved for 24 h and then stimulated for 6 h with SFM, 10 µmol/L LPA, or 1 µg/mL doxorubicin (Dox). The cells were then fixed and double labeled for endogenous p53 and p21 by indirect immunofluorescence. The majority of LPA-stimulated cells showed a reduction in the nuclear labeling of p53 and p21 relative to untreated cells. B. The proportion of cells that showed nuclear labeling of p21 following LPA (10 µmol/L) or doxorubicin (1 µg/mL) stimulation was quantified by evaluation of 100 cells/replicate of triplicate samples. Columns, mean percentage of cells with nuclear p21 labeling; bars, SE. **, P < 0.01, compared with control SFM cells.

 
Because the expression of p21CIP1 is transcriptionally regulated by p53, we hypothesized that the LPA-induced reduction in cellular p21CIP1 was caused by a reduction in the transcriptional activity of p53. To directly test this hypothesis, we used a p53-stimulated luciferase reporter gene assay to determine the effects of LPA stimulation on p53-dependent transcription (Fig. 4 ). A549 cells were transiently cotransfected with a plasmid encoding a firefly luciferase reporter gene, whose expression is driven by a basal promoter and an upstream p53 response element, and with the plasmid pRL-TK, which constitutively expresses Renilla luciferase. The latter plasmid serves to control for variations in transfection efficiency. Stimulation of native A549 cells with 10 µmol/L LPA caused a significant reduction in p53-dependent transcription of luciferase to ~40% of that observed in untreated cells (Fig. 4A, Control). This indicated that the observed reduction in p53 abundance correlated with a decrease in p53-dependent transcription, which likely explains the concurrent loss of p21CIP1.


Figure 4
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  		FIGURE 4. Overexpression of LPA1, LPA2, or LPA3 suppresses p53-dependent transcription in A549 cells. A. A549 cells were transiently transfected with plasmids encoding either empty vector, FLAG-LPA1, FLAG-LPA2, or FLAG-LPA3 along with p53 luciferase reporter and pRL-TK (Renilla luciferase) constructs. Twenty-four hours after transfection, cells were incubated overnight with 10 µmol/L LPA before measuring both firefly and Renilla luciferase as described in Materials and Methods. The data are percent of control and the graphs are from a representative experiment, which was repeated at least thrice with similar results. Columns, mean of triplicate samples; bars, SE. **, P < 0.01, comparison of cells treated with or without LPA to untreated, control cells. B. Extracellular phospholipase B (PLB) prevents the inhibition of p53-dependent transcription by overexpressed LPA1 receptors. A549 cells were transiently transfected with plasmids encoding vector alone or LPA1 for 24 h before treatment overnight with 3 units/mL phospholipase B. Firefly and Renilla luciferase activities were measured. Columns, average of three independent experiments and shown as percentage of control; bars, SE.

 
Previous studies have shown that LPA1, LPA2, and LPA3 can stimulate common effector pathways (18). Because A549 cells predominantly express LPA1 (35), we investigated the effects of overexpressing LPA1, LPA2, and LPA3 on p53-dependent transcription in A549 cells to determine if other LPA receptors shared the ability to attenuate p53 (Fig. 4A). A549 cells were cotransfected with a plasmid encoding LPA1, LPA2, and LPA3 as well as the p53 transcriptional reporter gene plasmids. Remarkably, overexpression of LPA1, LPA2, or LPA3 all led to a severe reduction in p53-dependent transcription to <20% of that observed in the corresponding control, untransfected cells (Fig. 4A). Interestingly, the inhibition of p53-dependent transcription on overexpression of LPA receptors did not require the addition of exogenous LPA. Many tumor cells produce extracellular LPA either through the action of the secreted lysophospholipase D, autotaxin, on lysophosphatidylcholine or through the action of the recently described intracellular acylglycerol kinase (21, 22, 47). We hypothesized that these cells produced LPA themselves, which in turn would activate the increased numbers of transfected receptors. We tested this by adding phospholipase B to the incubation medium. Phospholipase B degrades both lysophosphatidylcholine and LPA and therefore decreases LPA receptor activation (48). Addition of exogenous phospholipase B, indeed, prevented the inhibition of p53-dependent transcription by overexpressed LPA1 receptors in A549 cells (Fig. 4B). This result supports the hypothesis that A549 cells produce the LPA needed to activate the endogenous or overexpressed LPA receptors. However, an alternative possibility is that overexpressed LPA receptors inhibit p53-mediated transcription in a ligand-independent manner. Support for this possibility comes from the observation that overexpressed LPA1 receptors protect Schwann cells from apoptosis induced by serum deprivation in a ligand-independent manner (28).

To investigate whether overexpression of LPA receptors inhibited the activity of p53 in other cell types, we used HepG2 human hepatoma cells. These cells do not express endogenous Edg family LPA receptors (29, 49) and liver cells secrete abundant lysophosphatidylcholine (29, 49), which can be converted to LPA by autotaxin (22). We first examined the effects of overexpressing LPA receptors or green fluorescent protein, as a control, on the cellular distribution of endogenous p53 (Fig. 5A ). Indirect immunofluorescence was used to localize the stabilized form of p53 (phosphorylated on Ser15). The p53 protein was localized in the nucleus and in the cytoplasm of untransfected cells and in cells transfected with green fluorescent protein alone (Fig. 5A, GFP). In cells, overexpressing LPA1, LPA2, or LPA3, p53 was absent from the nucleus and greatly diminished in the cytoplasm (Fig. 5A). Next, we examined the effects of LPA receptor overexpression on the transcriptional activity of p53 in HepG2 cells. As expected, LPA stimulation of native HepG2 cells did not affect p53-dependent transcription significantly (Fig. 5B, Control), which is consistent with previous reports showing that HepG2 cells are unresponsive to LPA (49-51). Overexpression of any of the three Edg family-encoded LPA receptors, LPA1, LPA2, or LPA3, strongly reduced p53-dependent transcription in HepG2 cells to approximately 20% to 35% of that observed in the corresponding control, untransfected cells. Again, this effect was observed independently of whether LPA was added to the incubations (Fig. 5B). This could be due either to LPA-independent signaling by the overexpressed receptors or to endogenous production of LPA and stimulation of LPA receptors. Addition of phospholipase B reversed the inhibition of p53-dependent transcription of luciferase in HepG2 that expressed LPA1 (results not shown) as it did in A549 cells. We also compared the effects of overexpressing another Gi-coupled receptor, the M2 muscarinic acetylcholine receptor (mAChR), or the inactive LPA1 R124A mutant receptor on p53-mediated transcriptional activation (Fig. 5C). Arg124 is conserved in all Edg family G protein–coupled receptors and is critical for interaction with LPA or sphingosine-1-phosphate (52, 53). Unlike LPA1, overexpression of M2 mAChRs did not alter p53-dependent transcription and stimulation of these cells with the muscarinic agonist carbachol (10 mmol/L) actually enhanced p53-mediated transcription. Most importantly, we observed that overexpression of LPA1 R124A also did not inhibit p53-mediated transcription, whereas wild-type LPA1 inhibited luciferase expression by 50%. Regardless of whether overexpressed LPA receptors inhibit p53 through a ligand-independent mechanism or through an autocrine mechanism, our results indicate that signaling through functional LPA receptors promotes the reduction in the cellular abundance of p53, which in turn results in decreased p53-dependent transcription.


Figure 5
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  		FIGURE 5. Overexpression of LPA1, LPA2, or LPA3 suppresses p53 transcriptional activity in HepG2 cells. A. HepG2 cells grown on coverslips in complete medium were transiently transfected with pEGFP (control) or FLAG-tagged LPA receptors (as indicated in the figure) before processing for indirect immunofluorescence. Primary antibody staining to determine the localization of LPA receptors was done using mouse anti-FLAG antibodies and Ser15-phosphorylated p53 was detected using rabbit anti-phosphorylated p53 (Ser15) antibodies in the presence of 10% saponin. Fluorescently labeled secondary Cy antibodies were used to visualize primary antibodies. B. Effects on p53-dependent transcription. HepG2 cells were seeded in 24-well dishes and then transiently transfected in SFM with 0.2 µg plasmids encoding either empty vector, FLAG-LPA1, FLAG-LPA2, or FLAG-LPA3 along with 0.2 µg p53 luciferase reporter and 8 ng pRL-TK (Renilla luciferase) construct. Twenty-four hours after transfection, cells were incubated overnight with 10 µmol/L LPA before measuring both firefly and Renilla luciferase as described in Materials and Methods. The data are presented here as percent of control and the graphs are from a representative experiment, which was repeated at least thrice with similar results. Columns, mean of triplicate samples; bars, SE. **, P < 0.01, comparison of cells treated with or without LPA to untreated, control cells. C. Neither inactive LPA1 R124A nor M2 mAChRs inhibit p53-mediated transcription. A549 cells were transfected with plasmids encoding either LPA1, LPA1 R124A, or M2 mAChRs along with the p53 luciferase reporter construct and the Renilla control plasmid. The cells were stimulated with either 10 µmol/L LPA or 1 mmol/L carbachol, respectively, and processed for determination of luciferase activities. The data are presented here as percent of control and the graphs are from a representative experiment, which was repeated at least thrice with similar results. Columns, mean of triplicate samples; bars, SE. **, P < 0.01, comparison of cells treated with or without LPA to untreated, control cells.

 
The LPA-Dependent Reduction of p53 Abundance Is Dependent on PI3K
To investigate the LPA signaling pathways that facilitate the reduction of p53, we quantified the effects of different pharmacologic inhibitors of known LPA signaling pathways on the LPA-induced reduction in the proportion of p53NUC-bright cells. Treatment of these cells with LPA alone (10 µmol/L) decreased the proportion of p53NUC-bright cells to ~35% of the total population (Fig. 6, Control ). Preincubation of these cells with the LPA1/LPA3-selective antagonist VPC32183(S) by itself increased the proportion of p53NUC-bright cells. As expected, the antagonist completely prevented the LPA-induced reduction of nuclear p53 staining. Previous studies have shown that LPA receptors promote cell survival of cultured Schwann cells by activation of the PI3K pathway (28). Consistent with this, incubation with the PI3K inhibitor LY294002 also prevented the LPA-induced reduction of nuclear p53 labeling in A549 cells. Interestingly, LY294002 treatment alone also increased the proportion of p53NUC-bright cells, similar to the effects of VPC32183(S). In contrast, the mitogen-activated protein kinase/extracellular signal-regulated kinase kinase inhibitor PD98059 did not protect cells from the LPA-induced reduction of nuclear p53 staining. These results suggest that the LPA-dependent degradation of p53 is dependent on LPA receptor activation and on PI3K.


Figure 6
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  		FIGURE 6. LPA-dependent reduction of cellular p53 is dependent on LPA binding and PI3K. A549 cells were grown on glass coverslips and serum starved for 24 h. The cells were incubated with SFM, 10 µmol/L LPA, 50 µmol/L LY294002, 10 µmol/L VPC32183(S), 50 µmol/L PD980559, or a combination of LPA with the aforementioned reagents for 6 h before fixation and indirect immunofluorescence localization of p53. Quantification of cells exhibiting bright p53 nuclear staining was conducted as described previously in Fig. 1B. **, P < 0.01, compared with control, LPA-treated cells. The data are from a representative experiment that was repeated thrice with similar results.

 
LPA Partially Protects A549 Cells from Genotoxic Stress-Induced Cell Death
Several genotoxic drugs, such as the transcription inhibitor actinomycin D, enhance the nuclear accumulation of p53, which ultimately induces apoptosis in cells (37-39). To investigate the physiologic significance of LPA-induced reduction of p53, we examined the effects of LPA on actinomycin D–induced cell death in A549 cells. LPA protects both ovarian cancer cells and normal intestinal epithelial cells from apoptosis induced by chemotherapeutic agents (54, 55). A549 cells were incubated with or without 10 µmol/L LPA in the presence or absence of 2 µg/mL actinomycin D 24 h before determining cell viability (Fig. 7A ). Actinomycin D reduced the percentage of viable cells by 52% compared with control cells. In contrast, coincubation of actinomycin D and LPA only resulted in a 25% reduction in viable cells. Thus, LPA partially protected cells from actinomycin D–induced cell death. We also examined the effects of these treatments on the abundance of total p53 and "stabilized" p53 (i.e., p53 that is phosphorylated at Ser15; Fig. 7B). Relative to untreated cells, LPA significantly reduced the abundance of both total and phosphorylated p53 to 0.47 and 0.33, respectively. Treatment with 2 µg/mL actinomycin D increased the abundance of total and phosphorylated p53 to 1.64 and 2.66, respectively. Addition of both LPA and actinomycin D increased the abundance of total and phosphorylated p53 to 1.27 and 1.89, respectively, which was less than the increase observed with actinomycin D alone. This indicates that the ability of LPA to partially protect A549 cells from actinomycin D–induced cell death correlates with the ability of LPA to suppress the increase in total and phosphorylated p53 abundance observed with actinomycin D alone. Thus, the LPA-induced attenuation of p53 signaling provides an additional mechanism that can contribute to LPA-dependent cell survival.


Figure 7
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  		FIGURE 7. LPA partially protects A549 cells from actinomycin D–induced cell death and increased p53 abundance. A. A549 cells were grown in either SFM, 10 µmol/L LPA, 2 µg/mL actinomycin D, or both 10 µmol/L LPA and 2 µg/mL actinomycin D for 24 h. Cell proliferation was measured using WST-1 cell proliferation reagent as described in Materials and Methods. The results were normalized to cells grown in SFM alone (control). Columns, mean of six replicates per condition from a representative experiment that was repeated thrice with similar results; bars, SE. B. Western blotting. A549 cells were grown in either SFM, 10 µmol/L LPA, 2 µg/mL actinomycin D, or both 10 µmol/L LPA and 2 µg/mL actinomycin D for 24 h. Whole-cell extracts were prepared and processed for Western blot detection of total p53, phosphorylated p53, and glyceraldehyde-3-phosphate dehydrogenase as described in Materials and Methods. The numbers above the lanes indicate the band intensities as a fraction of the p53 band observed in untreated control cells.

 

   Discussion
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
Materials and Methods
 Acknowledgements
 References
 
In this study, we describe a novel target of LPA-induced cell signaling, namely that LPA through stimulation of its receptors reduces the abundance of the p53 tumor suppressor, which in turn decreases p53-dependent transcription. The LPA-dependent reduction of p53 is caused by enhanced degradation because inhibition of proteasomal action prevented the loss of p53. The signaling pathway leading from LPA receptors to p53 degradation involves PI3K. This novel action of LPA contributes to the cell survival and growth-promoting effects of LPA. It helps to explain how increased LPA action promotes tumor progression and protects tumors against chemotherapy.

p53 is degraded both in the cytoplasm and in the nucleus through a proteasomal pathway (42, 56, 57), which we now show can be activated by LPA. We found that LPA stimulation of A549 lung carcinoma cells led to a rapid loss of p53 protein from the nucleus of these cells and a concurrent decrease in the cellular levels of total p53 (Figs. 1, 2, and 5). p53 contains several nuclear import signals as well as multiple nuclear export signals (57-59). In nontumorigenic cells, association of p53 with the E3 ubiquitin ligase MDM2 promotes ubiquitylation and nuclear export of p53 with subsequent degradation of the protein by cytoplasmic proteasomes (40, 41, 60). However, p53 can also be degraded in the nucleus by nuclear proteasomes (42-44). Our data show that LPA stimulation leads to a dose-dependent, LPA-induced decrease of nuclear p53 (Fig. 1B). Furthermore, the proteasomal inhibitor lactacystin prevented the LPA-induced loss of nuclear p53 (Fig. 2B). Given that lactacystin inhibits both nuclear and cytoplasmic degradation (44), the accumulation of p53 in the nucleus in the presence of LPA and lactacystin suggests that LPA may also stimulate p53 degradation in the nucleus.

Consistent with the LPA-induced loss of nuclear p53 protein, we observed a corresponding decrease in the expression of the endogenous gene target, p21CIP1 (Fig. 3). One of the consequences of p53 mobilization is to stimulate the transcription of the cyclin inhibitor p21CIP1, which in turn promotes G1 cell cycle arrest (46). The concurrent loss of p21 suggested that the LPA-induced loss of p53 led to decreased p53-dependent transcription. In support of this hypothesis, we found that LPA signaling, either by endogenous or overexpressed LPA receptors (Figs. 4 and 5), inhibited the expression of a p53-dependent luciferase transcriptional reporter gene.

It was intriguing that overexpression of LPA receptors alone was sufficient to promote the inhibition of p53-dependent transcription. However, several lines of evidence support the specificity of this response. First and most important, stimulation of native, untransfected A549 cells with LPA alone inhibited p53-dependent transcription (Fig. 4A, control cells). This shows that ligand stimulation of endogenous LPA receptors is sufficient to reduce p53. Second, treatment of native A549 cells with the LPA3-selective agonist OMPT (36) also potently reduced nuclear p53 localization (Fig. 2B). Third, treatment of cells with the LPA1/LPA3-selective antagonist VPC32183(S) suppressed the LPA-induced loss of p53 from the nucleus of A549 cells (Fig. 6A). Fourth, neither overexpression of inactive LPA1 R124A nor Gi-coupled M2 mAChRs inhibited p53-mediated transcription (Fig. 5C). Finally, we observed that the inhibition of p53-dependent transcription in cells by LPA receptor overexpression is prevented by incubation with exogenous phospholipase B, which degrades extracellular LPA that is likely generated by secreted autotaxin, and thus inhibits LPA signaling (Fig. 4B; refs. 21, 22, 35, 48).

There are two possible explanations for the LPA-independent reduction of p53-mediated transcription by overexpressed LPA receptors. First, it is quite possible that overexpressed LPA receptors signal in a ligand-independent manner. There are numerous examples of G protein–coupled receptors that are constitutively active, especially those encoded by human viruses, such as Kaposi's sarcoma virus, EBV, and cytomegalovirus (61-63). Indeed, overexpression of LPA1 in Schwann cells protects them from apoptosis induced by serum deprivation, with addition of LPA having little additional effect (64). Furthermore, overexpression of LPA1 in RH7777 hepatoma cells can stimulate actin stress fiber formation even in the absence of LPA, although addition of LPA does potentiate this effect (64). Another piece of data from our current work that is consistent with this hypothesis is the finding that overexpression of neither inactive LPA1 R124A nor M2 mAChRs reduced p53 activity (Fig. 5C). A second possibility is an autocrine mechanism involving a combination of the intracellular production of LPA and the conversion of extracellular lysophosphatidylcholine to LPA by autotaxin, which is expected to partially activate the endogenous LPA receptors in A549 cells and the overexpressed receptors in HepG2 cells. Consistent with this hypothesis, we found that treatment of control cells either with phospholipase B or with the LPA receptor antagonist VPC32183(S) (Figs. 4B and 6) increased basal p53-mediated transcription and in nuclear p53 labeling, which would be expected if autocrine stimulation of endogenous LPA receptors was occurring. Regardless of the mechanism, our data indicate that LPA signaling promotes the reduction of cellular p53 abundance.

What are the LPA signaling pathways that are responsible for the reduction of p53? LPA receptors stimulate multiple G protein–mediated signaling pathways (18, 64, 65). Our data indicate that PI3K is involved in the LPA-dependent reduction of cellular p53 (Fig. 6). The PI3K/Akt signaling pathway is important for cell survival of normal and tumorigenic cells (3, 28, 66, 67). Downstream of PI3K, Akt phosphorylates the E3 ubiquitin ligase MDM2, which enhances both MDM2's nuclear import and its ubiquitin ligase activity. Thus, LPA stimulation of PI3K may lead to increased p53 degradation via enhanced MDM2 activity.

What are the consequences of LPA-induced attenuation of p53? LPA promotes the proliferation of a variety of normal and tumorigenic cells and it stimulates both cell proliferative and antiapoptotic signaling pathways (20). Studies on the cellular mechanisms involved in mediating the growth-promoting effects of LPA have focused on the activation of Ras/mitogen-activated protein kinase and Rho GTPase signaling pathways as well as stimulation of the serine/threonine kinase Akt. However, relatively little is known about the effects of LPA signaling on cell cycle regulators, such as the p53 tumor suppressor. Our studies highlight a previously unappreciated effect of LPA signaling, namely that it decreases p53 concentrations and potently inhibits transcriptional activity of p53. In normal cells, activated p53 promotes G1-S cell cycle arrest and/or apoptosis in response to DNA damage (45). p53 is either mutated or inactivated in 50% of all cancers, which enhances the ability of cancer cells to evade cell cycle arrest and apoptosis. In many tumor cells that contain wild-type p53, key downstream effectors, such as the cyclin kinase inhibitor p21CIP1, are often mutationally inactivated.

Our results show that LPA receptor-mediated inhibition of p53 activity can enhance cancer cell survival by preventing the induction of apoptosis. We showed that LPA could partially protect A549 cells from actinomycin D–induced cell death and suppressed the actinomycin D–stimulated increase in total and phosphorylated p53 abundance (Fig. 7). Actinomycin D inhibits RNA polymerase activity (68), which in turn activates p53 (69). This action could also contribute to the observed effects of LPA in protecting cancer cells from cell death caused by chemotherapeutic agents (54). The action of LPA in increasing cyclin D1 expression (24) would favor S-phase entry when LPA also decreases p53 expression. This would decrease the normal effects of p53 in blocking the transition at G1-S and cell cycle progression. This unscheduled (by increased cyclin D) and unrestricted (by decreased p53) S-phase entry could lead to the accumulation of DNA damage after many cell generations and ultimately favor cell transformation.

Furthermore, several recent studies have shown the transcription-independent induction of apoptosis by cytosolic p53 and p53 localized to mitochondria (70-73). Cytosolic p53 directly binds and activates the proapoptotic protein Bax, which induces the permeabilization of the outer mitochondrial membrane, cytochrome c release, and caspase-3 activation (73). Alternatively, a fraction of the total cellular p53 is translocated to the outer mitochondrial membrane in response to apoptotic stimuli (71, 72). At the mitochondrial membrane, p53 interacts with Bcl-2 family members to promote cytochrome c release and caspase-3 activation. Thus, the LPA-mediated reduction of cellular p53 is expected to also diminish the ability of p53 to directly promote apoptosis in cancer cells through its interactions with proapoptotic and antiapoptotic Bcl-2 family proteins. Indeed, this may contribute to the observed protection of A549 cells from actinomycin D–induced apoptosis by LPA. Taken together, these actions of LPA on p53 along with other proliferative signals through the Ras/Raf/extracellular signal-regulated kinase pathway can contribute to the actions of LPA in promoting tumor growth.

Our work, therefore, identifies LPA as a physiologic agonist that decreases p53 expression and thus shows a further dimension whereby LPA can both decrease apoptosis and increase cell division. The present results provide further insights into the role of LPA in promoting cell division versus apoptosis and thereby cell proliferation. The LPA-induced degradation of p53 and its consequent decreased control of cell cycle progression and cell death provide a novel dimension for understanding how LPA promotes tumor progression and protects tumor against chemotherapy.


   Materials and Methods
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
Acknowledgements
 References
 
Cell Culture
HepG2 cells were obtained from Dr. Athanassios Sambanis (Georgia Institute of Technology, Atlanta, GA) and maintained as described previously (51). A549 cells were purchased from the American Type Culture Collection and grown at 37°C and 5% CO2 in F12K Kaighn's modification medium (Mediatech) supplemented with 10% fetal bovine serum, 100 IU/mL penicillin, 100 µg/mL streptomycin (HyClone), and 1.5 g/L NaHCO3 (Biosource International).

Reagents
LPA (18:1; 1-oleoyl-2-hydroxy-sn-glycero-3-phosphate), OMPT, and VPC32183(S) [(S)-phosphoric acid mono-{2-octadec-9-enoylamino-3-[4-(pyridine-2-ylmethoxy)-phenyl]-propyl} ester; Avanti Polar Lipids] were reconstituted in 1% fatty acid–free, charcoal-stripped bovine serum albumin as described previously (74, 75). Mouse anti-p53 antibody (DO-1), mouse anti-phosphorylated Ser15 p53 antibody, and rabbit anti-p53 (FL-393), which both detect total p53, were from Santa Cruz Biotechnology, Inc. Mouse anti-paxillin and mouse anti-Bip antibodies (BD Biosciences) were used to monitor loading accuracy. Mouse anti-p21 antibodies were purchased from BD Biosciences and Calbiochem, respectively. All other reagents were from Sigma Chemical Co., including phospholipase B, which was isolated from Vibrio species; Sigma has discontinued this product.

Plasmids
The original plasmid encoding the human FLAG-LPA1 receptor was a kind gift from Dr. Junken Aoki (University of Tokyo, Tokyo, Japan), and the modified construction of the FLAG-LPA1 plasmid has been reported previously (76, 77). To enhance the cell surface expression of LPA2 and LPA3, PCR was used to attach a signal leader sequence from the influenza hemagglutinin protein onto the NH2 terminus of the plasmid preceding the FLAG epitope tag. The FLAG-LPA2 vector was constructed using the primers 5'-ATGCGGATCCATGGACTACAAAGACGAT-3' and 5'-GATCTCAGTCCTGTTGGTTGGG-3'. The FLAG-LPA3 vector was constructed by isolating total RNA from OVCAR-3 cultured cells with a GenElute Direct mRNA Miniprep kit (Sigma). Following isolation, reverse transcription-PCR was done using the following oligonucleotides: 5'-GATCATGAAGACCATCATCGCCCTGAGCTACATCTTCTGCCTGGTGTTCGCCGACTACAAGGACGATGATGACAAGATGAATGAGTGTCAC-3' and 5'-CGATTTAGGAAGTGCTTTTA-3'. The reverse transcription-PCR included 10 ng RNA, 10 µmol/L of sense and antisense primers, deoxynucleotides, reaction buffer, and 0.5 units of reverse transcriptase/Taq DNA polymerase (Invitrogen) in a final volume of 50 µL. The mixes were first incubated at 50°C for 30 min to allow mRNA to be copied into cDNA followed by a 2-min hold at 94°C and 40 cycles of 94°C for 15 s, 55°C for 30 s, and 72°C for 2 min. After the PCR, a single 1,062-bp band was observed, which was purified and subcloned into pcDNA3.1 V5/His mammalian expression vector (Invitrogen).

Transfections
For all transcriptional reporter gene assays, HepG2 cells (7 x 104) and A549 cells (3 x 104) were transiently transfected using Lipofectamine or Lipofectin (Invitrogen), respectively, according to the manufacturer's guidelines. Both A549 and HepG2 cells were transfected with 0.2 µg of plasmid DNA encoding empty vector, the indicated G protein–coupled receptors, and the p53 luciferase reporter construct; 0.8 ng of the control pRL-TK (Renilla luciferase) plasmid was used. Transient expression levels of FLAG-tagged LPA receptors were kept within 10% of each other (as assessed by immunofluorescence).

Immunoblotting
A549 cells were grown in 150-mm dishes for 24 h before washing with SFM and starving in SFM for 12 to 16 h before the treatments indicated above and in figure legends. Cells were rinsed with ice-cold PBS supplemented with phosphatase inhibitors (Active Motif), detached by scraping, and collected by centrifugation. Pellets were then solubilized on ice for 30 min with intermittent agitation in lysis buffer [10 mmol/L Tris (pH 7.4), 100 mmol/L NaCl, 1 mmol/L EDTA, 1 mmol/L EGTA, 1 mmol/L NaF, 20 mmol/L Na4P2O7, 2 mmol/L Na3VO4, 1% Triton X-100, 10% glycerol, 0.1% SDS, 0.5% deoxycholate (Biosource International)] supplemented with 1 mmol/L phenylmethylsulfonyl fluoride from a 0.3 mol/L stock and protease inhibitor cocktail (1:10 dilution; Sigma). Protein concentration was quantified using a bicinchoninic acid protein assay (Pierce). The samples (12-20 µg protein per lane) were then separated by 10% SDS-PAGE and transferred to nitrocellulose. The binding of primary antibodies was detected using SuperSignal West Pico Chemiluminescent Substrate (Pierce).

Indirect Immunofluorescence and Quantification
Cells were grown on glass coverslips, treated as described, and fixed in 2% formaldehyde in PBS. After blocking nonspecific sites with 10% FCS (PBS-serum), the cells were permeabilized with 0.02% saponin and incubated with primary antibodies. Coverslips were subsequently rinsed thrice with PBS-serum and incubated with fluorescently labeled secondary antibodies. During the final wash, Hoechst 33342 dye (Molecular Probes) was added to label the DNA. Samples mounted on glass slides were observed with an Olympus BX40 epifluorescence microscope equipped with a 60X plan-apochromat lens and digital photomicrographs were obtained with a MagnaFire SP digital camera. All photographs were obtained using the same exposure time.

For quantification of fluorescence intensity, photomicrographs of 15 to 25 cells per time point or experimental treatment were obtained from each experiment, which itself was repeated at least three independent times. The fluorescence intensity of the nuclear-localized p53 was measured in each cell using MetaMorph Imaging Software (Universal Imaging Corp.). The p53 fluorescence intensity was normalized by subtracting the background and then dividing that value by the fluorescence intensity of DNA labeled with Hoechst dye for each cell. Normalized data averages of each photomicrograph from all experiments were combined to obtain a grand average throughout. The data are presented as the mean ± SE.

MetaMorph image analysis showed that cells with bright nuclear p53 had an average fluorescence intensity of 0.83 ± 0.06 (n = 15 cells), whereas p53NUC-diminished cells had an average fluorescence intensity of 0.38 ± 0.03 (n = 15 cells). Changes in the proportion of p53NUC-bright cells were quantified by scoring cells whose nuclear p53 fluorescence intensity was judged as being approximately equal to the average fluorescence intensity of cells with bright nuclear p53 staining (e.g., ~0.8). One hundred cells per sample were randomly selected and scored as exhibiting either bright nuclear p53 staining or diminished nuclear p53 staining based on relative fluorescence intensity. The data are presented as the mean percentage of p53NUC-bright cells ± SE.

Luciferase Reporter Gene Assay
Both firefly luciferase and Renilla (pRL-TK) luciferase activities were measured 36 h after transfection using a dual luciferase assay kit (Promega) as described previously (51, 76). The pp53-TA-luc vector contains a p53 response element that is upstream of the luciferase reporter (BD Biosciences Clontech). This plasmid, therefore, provides a measure of the transcriptional activity of p53. The plasmid, pRL-TK, which constitutively expresses Renilla luciferase (Promega), was used to normalize for differences in transfection efficiency between samples. The normalized value is defined as the ratio of the p53-induced firefly luciferase activity to Renilla luciferase. The data are presented as percent of control and are compared with untreated controls. They are shown as the mean ± SE of triplicate measurements from a representative experiment that was repeated at least thrice.

Cell Proliferation
A549 cells (1 x 104) were grown in the wells of a 96-well plate for 12 h in complete medium and then rinsed thrice in SFM before incubating in either SFM, 10 µmol/L LPA, 2 µg/mL actinomycin D, or both 10 µmol/L LPA and 2 µg/mL actinomycin D for 24 h. After this incubation, 20 µL of WST-1 cell proliferation reagent (Roche) were added to each well for 1 h at 37°C. This assay quantifies cell proliferation and cell viability based on the cleavage of the tetrazolium salt WST-1 to formazan. The absorbance of each well was measured at 450 nm. The results were normalized to cells grown in SFM alone (control) and are the mean ± SE of six replicates per condition from a representative experiment that was repeated thrice with similar results.

Statistical Analysis
The data were analyzed using either a single-factor or two-factor ANOVA followed by a Tukey's statistical test.


   Acknowledgements
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
References
 
We thank Dr. Nael McCarty for critical reading of the manuscript.


   Notes
Top
 Notes
Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
Grant support: Public Service grant HL-16734, Georgia Cancer Coalition grant G-32-6CM (H. Radhakrishna), and Canadian Institutes of Health Research grant MOP81137 (D.N. Brindley).

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.

Note: M.M. Murph and J. Hurst-Kennedy contributed equally to this work.

Received 10/ 6/06; revised 7/24/07; accepted 8/ 8/07.


   References
Top
 Notes
 Abstract
 Introduction
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 

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