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J. Biol. Chem., Vol. 281, Issue 21, 14700-14710, May 26, 2006

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Attenuation of Peroxisome Proliferator-activated Receptor (PPAR) Mediates Gastrin-stimulated Colorectal Cancer Cell Proliferation^*

From the Section of Gastroenterology, Boston University School of Medicine and Boston Medical Center, Boston, Massachusetts 02118

Received for publication, March 20, 2006

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Peroxisome proliferators-activated receptor (PPAR) has been shown to suppress cell proliferation and tumorigenesis, whereas the gastrointestinal regulatory peptide gastrin stimulates the growth of neoplastic cells. The present studies were directed to determine whether changes in PPAR expression might mediate the effects of gastrin on the proliferation of colorectal cancer (CRC). Initially, using growth assays, we determined that the human CRC cell line DLD-1 expressed both functional PPAR and gastrin receptors. Amidated gastrin (G-17) attenuated the growth suppressing effects of PPAR by decreasing PPAR activity and total protein expression, in part through an increase in the rate of proteasomal degradation. G-17-induced degradation of PPAR appeared to be mediated through phosphorylation of PPAR at serine 84 by a process involving the biphasic phosphorylation of ERK1/2 and activation of the epidermal growth factor receptor (EGFR). These results were confirmed through the use of EGFR antagonist AG1478 and MEK1 inhibitor PD98059. Furthermore, mutation of PPAR at serine 84 reduced the effects of G-17, as evident by inability of G-17 to attenuate PPAR promoter activity, degrade PPAR, or inhibit the growth suppressing effects of PPAR. The results of these studies demonstrate that the trophic properties of gastrin in CRC may be mediated in part by transactivation of the EGFR and phosphorylation of ERK1/2, leading to degradation of PPAR protein and a decrease in PPAR activation.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Colorectal cancer (CRC)² is the third most common cause of cancer-related death in the United States, accounting for 10% of all deaths caused by malignancy. The American Cancer Society estimated that in 2005, 145,000 Americans would be diagnosed with CRC, of whom 56,000 would die. CRC generally arises from benign adenomas, which progress into malignant adenocarcinomas (2, 3). The development of CRC appears to involve a multistep process of genetic mutations combined with largely undefined environmental factors whereby normal epithelial cells undergo dysplastic transformation, followed by proliferation and eventual histological progression to neoplasia (2). This transition appears to require multiple genetic alterations, such as mutations in the adenomatous polyposis coli (APC) gene. After the initial mutations have established a neoplastic phenotype, other factors appear to promote proliferation and neoplastic progression (2).

One such potential growth factor is the polypeptide hormone gastrin. In addition to its recognized role in the physiological regulation of acid secretion (4–7), another biological property attributed to gastrin is its trophic effect on the gastrointestinal (GI) mucosa (6–8). Numerous studies have provided evidence that gastrin peptides play an integral role in promoting colorectal tumor growth (3), as well as other malignancies throughout the GI tract (8, 9). In a well controlled, elegantly designed epidemiologic study of nearly 130,000 individuals, Thorburn et al. (10) found that prolonged hypergastrinemia comprised a risk factor for the development of CRC. This study found a 3.9-fold increased risk of CRC due to elevated circulating gastrin levels, prompting the authors to conclude that 8.9% of all CRCs may be attributed to hypergastrinemia (10). Studies in transgenic mice overexpressing gastrin have demonstrated increased proliferation of the gastric and colonic epithelium after eight months. When these mice were followed for longer periods (20 months), an increased tendency to develop neoplasia was observed (11).

Several molecular forms of gastrin are synthesized and released into the circulation, with the predominant peptide being -amidated gastrin-17 (G-17) (9). Progastrin precursor peptides such as glycine-extended gastrin (Gly-G) appear to affect CRC in vivo principally via autocrine pathways, while fully processed gastrin also utilizes endocrine pathways (12–16). Smith and Watson (17) reported that 80% of all colorectal adenomas express gastrin/cholecystokinin-2 receptors (CCK-2R), and Ciccotosto et al. (18) found that whereas 70% of CRC expressed fully processed -amidated gastrin, 100% produced progastrin precursor peptides. These studies all strongly suggest a role for gastrin in GI cell proliferation and carcinogenesis.

Although the trophic effects of gastrin are well recognized, the molecular and intracellular mechanisms by which gastrin modulate cell growth in the GI tract have not been fully elucidated. Previous studies have reported that stimulation of the CCK-2R by gastrin activates various signal transduction pathways implicated in cell proliferation, such as the mitogen-activated protein kinases (MAPK), which include ERK, JNK, and p38 kinase (8). One potential downstream target of the MAPKs is peroxisome proliferator-activated receptor (PPAR) (19–24). The possibility of a functional relationship between gastrin and PPAR has not been previously evaluated.

PPAR, a member of the nuclear hormone receptor family, functions as a transcription factor that regulates several biological processes, including growth and differentiation (25, 26). In addition to its recognized role in adipogenesis (31), PPAR has been shown to modulate the growth of cells in various organs. This trophic effect is most evident in the colon, where normal human colonic mucosa, colon adenocarcinoma, and cultured CRC cells express levels of PPAR1 equivalent to that detected in adipocytes (27, 28). Activation of PPAR in cultured colon cells inhibits growth and induces differentiation, reverses the malignant phenotype, and promotes apoptosis (27, 29–32). In CRC cells, PPAR activation results in both an increase in the cyclin-dependent kinase inhibitors, p21 and p27 (33), which repress cell cycle progression, leading to a decrease in cell growth and an increase in the differentiation of cancer cells, and up-regulation of caspase activity (34, 35), resulting in DNA fragmentation and apoptosis. Moreover, a recent study demonstrated that 8% of primary colorectal tumors harbor a functional mutation in one allele of the PPAR gene, further supporting the role of PPAR as a tumor suppressor in humans (36).

The present studies were directed to determine whether changes in PPAR expression might mediate the effects of gastrin on the proliferation of CRC. We have demonstrated that gastrin-stimulated proliferation of CRC cells is associated with a significant concomitant decrease in cellular PPAR levels. Moreover, gastrin attenuates the inhibition of cell growth induced by PPAR agonists. Finally, our studies demonstrate that the trophic properties of gastrin may be mediated in part by transactivation of the epidermal growth factor receptor (EGFR) and phosphorylation of ERK1/2, leading to degradation of PPAR protein and a decrease in PPAR activation.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Materials—DLD-1 human adenocarcinoma cells were obtained from the American Type Culture Collection, SW48 human colon adenocarcinoma cells were kindly donated by Dr. B. Vogelstein (Baltimore, MD), and MC-26 cells, a transplantable mouse CRC cell line, were obtained from Dr. K. K. Tanabe (Boston, MA). The PPAR monoclonal (E-8, no. sc-7273) antibody was purchased from Santa Cruz Biotechnology (Santa Cruz, CA), and -amidated gastrin-17 (G-17) and Gly-G were purchased from Bachem (King of Prussia, PA) and Anspep (Parkville, Victoria, Australia), respectively. The phosphospecific PPAR polyclonal antibody (clone AW504) and the phosphospecific monoclonal antibody to the epidermal growth factor receptor (pTyr¹¹⁷³) were purchased from Upstate%20Biotechnology">Upstate Biotechnology (Waltham, MA). The phosphospecific monoclonal antibody to ERK1/2 (Thr²⁰²/Tyr²⁰⁴) was purchased from Cell Signaling Technologies (Beverly, MA), and the proteasome inhibitor, MG132 (N-carbobenzoxyl-Leu-Leu-Leucinal), cycloheximide, and thymidine were purchased from Sigma. [³H]Thymidine was purchased from PerkinElmer Life Sciences (Boston, MA). FuGENE 6 and Complete Protease Inhibitor mixture tablets were purchased from Roche, and the thiazolidinediones, ciglitazone and rosiglitazone, which stimulate PPAR activity, were purchased from Cayman Chemicals (Ann Arbor, MI). The pHD(x3)Luc vector was a kind gift from Dr. John Capone of McMaster University (Hamilton, ON). This vector contains three tandem repeats of the peroxisome proliferators response element from the promoter of the rat hydratase-dehydrogenase gene subcloned into the BamHI site of pCPSluc located immediately upstream of the carbamoyl-phosphate synthetase promoter. L-365,260 was generously provided by Dr. L. Iverson (Oxford, UK), pCMV--gal was purchased from Invitrogen, and PD98059, SB203580, and lactacystin were purchased from Calbiochem. The RNAeasy Mini kit and the SYBR green quantitative PCR master mix were purchased from Qiagen (Valencia, CA), and pTracerA/Bsd plasmid and ThermoScript^TM Reverse Transcriptase were purchased from Invitrogen. The Dual-Luciferase® Reporter assay system, the -Galactosidase Enzyme assay system, and the CellTiter 96® AQ_ueous One Solution Cell Proliferation (MTT) assay kit were purchased from Promega. The QuikChange XLII site-directed mutagenesis kit was purchased from Stratagene (La Jolla, CA). KpnI, XbaI, and the Quick Ligase kit were purchased from New England Bio-labs (Ipswich, MA).

Site-directed Mutagenesis and Generation of PPAR Phosphorylation Mutants—Site-directed mutagenesis was performed utilizing the QuikChange XLII kit with the forward primer, 5-GTGGAGCCTGCAGCTCCACCTTATTATTCTG-3, and the reverse primer, 3-CACCTCGGACGTCGAGGTGGAATAATAAGAC-5, and the pcDNA3-FLAG-wtPPAR vector as a template to generate a substitution of Ser⁸⁴ to Ala. An initial denaturation step was performed at 95 °C for 2 min. and followed by 20 cycles at 95 °C for 1 min, annealing at 57 °C for 1 min, and extension at 68 °C for 7.5 min. A final extension phase was performed at 68 °C for 7 min. The DNA sequence was confirmed by the Tufts University Core Facilities. PPAR sequence inserts were double-digested with KpnI and XbaI and subcloned into the pTracerA/Bsd vector utilizing the Quick Ligase Kit according to the manufacturer's instructions, at a 3:1 ratio of PPAR insert to pTracer. Stable cell lines expressing pTracer, pTracerFLAG-wtPPAR, or pTracerFLAG-mutPPAR were generated by blasticidin selection (5 µg/ml) and confirmed by Western analysis using the mouse monoclonal anti-PPAR antibody and the mouse monoclonal anti-green fluorescent protein (GFP) antibody, and by fluorescent light microscopy.

Cell Culture—The human CRC cell line DLD-1 and the mouse CRC cell line MC-26 were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum (FBS) (Invitrogen) and 1% penicillin/streptomycin (Invitrogen). The human CRC cell line SW48 was maintained in McCoy's medium (Invitrogen) supplemented with 10% FBS and 1% penicillin/streptomycin. All cell lines were kept in a humidified, 5% CO₂ environment. 24 h prior to experimentation, serum-containing media were replaced with serum-free media.

[³H]Thymidine Incorporation—Equal amounts of cells (3 x 10⁵ cells/well) were plated in 6-well plates. After 24 h of serum starvation in 0.5 mM thymidine-containing medium, cells were gently washed with phosphate-buffered saline (PBS) and then incubated for 22 h in fresh serum-free medium in the presence of G-17 or vehicle alone. In addition, to examine the involvement of the CCK-2R in DNA synthesis, 500 nM L-365,260, a specific CCK-2/gastrin receptor antagonist, was added to the culture medium. 4 h prior to termination of the experiment, cells were pulsed with [³H]thymidine at a final concentration of 1 µCi/well. At the end of the incubation period, the radioactive medium was aspirated, and cell monolayers were gently rinsed with PBS at 4 °C. Ice-cold 10% trichloroacetic acid was then added, and cells were gently rocked at 4 °C for 30 min. Cell monolayers were washed again with PBS, followed by incubation for 30 min at 25 °C in 1 N NaOH. Scintillation fluid was added, and samples were transferred into scintillation vials for measurement of [³H]thymidine using an automatic Beckman liquid scintillation counter.

Cell Counting—Equal amounts of cells (3 x 10⁵ cells) were plated onto 10-cm plates. After 24 h of serum starvation in 0.5 mM thymidine-containing medium, cells were gently washed with PBS, and the medium was replaced with 1% FBS-containing medium with or without G-17. After a 6-day incubation period, cells were trypsinized, harvested, and resuspended in equal volumes. Cell number was determined by counting with the aid of a hemacytometer under an inverted light microscope.

MTT Assays—MTT (3-(4,5-dimethylthiazoyl)-2,5-diphenyltetrazolium bromide) assays (Promega) were also performed to measure cell growth. This method measures the quantity of the formazan product, as measured by the ratio of 490/630 nm absorbance, which is proportional to the number of living cells in culture. Cells were seeded onto 96-well plates at a density of 5 x 10³ cells per well and incubated overnight in 10% FBS medium. The medium was then replaced with serum-free medium the following day and incubated for 24 h. To determine the effect of gastrin on cell growth, the medium were then replaced with medium containing 1% FBS and increasing concentrations of G-17 or Gly-G, and cells were grown for 4 days. To determine the effect of PPAR ligands on cell growth, ciglitazone and rosiglitazone, two PPAR agonists, were added after serum starvation in medium containing 1% FBS, and cells were grown for an additional 3 days. To determine the effects of gastrin on cell growth in the presence of PPAR ligands, DLD-1 cells were preincubated with gastrin peptides for 12 h prior to addition of ciglitazone or rosiglitazone. At the end of each experiment, 20 µl of CellTitre96® Aqueous Solution Reagent were added to each well, and the plate was incubated for 30 min. The absorbance ratio (490/630 nm) was recorded using a 96-well El_x800 universal plate reader (BIO-TEK Instruments, Inc., Winooski, VT).

Transient Transfection and Luciferase Assays—DLD-1 cells (4 x 10⁴ cells/well) were plated in 24-well plates. After an overnight incubation, cells were transiently transfected with the pHD(x3)luc vector in Opti-MEM for 16 h. To normalize for transfection efficiency, the cells were co-transfected with a pCMV--gal reporter construct. FuGENE 6 was used according to the manufacturer's instructions, and a FuGENE6 to DNA ratio of 3:1 was used in each transfection experiment. After a 16-h transfection period, the medium was replaced with medium containing 1% FBS in the presence or absence of 200 nM gastrin for 12 h. The cells were then treated with various concentrations of ciglitazone for an additional 16 h, after which cell lysates were collected and PPAR reporter activity measured using the luciferase assay system. Values were normalized to -galactosidase activity.

RNA Extraction and RT-PCR Analysis—DLD-1 cells (1.50 x 10⁶ per plate) were plated onto 10-cm diameter plates and incubated overnight. After 24 h of serum starvation, DLD-1 cells were incubated in the presence or absence of G-17. At the indicated time points, RNA was extracted using the RNAeasy Mini kit. Total RNA was measured, and 1 µg of total RNA was reverse-transcribed using the ThermoScript reverse transcriptase. The reverse transcriptional reaction was carried out at 50 °C for 60 min and 85 °C for 5 min. To quantify the amount of PPAR cDNA, all samples were subjected to PCR amplification using the QuantiTect SYBR Green PCR Kit (Qiagen). The forward primer PPAR-F, 5-TCTCTCCGTAATGGAAGACC-3 and reverse primer PPAR-R, 5-GCATTATGAGACATCCCCAC-3, were used according to the method of Terashita et al. (37). The PCR protocol was as follows: initial denaturation at 95 °C for 15 min, followed by 35 cycles at 94 °C for 15 s, annealing at 55 °C for 30 s, and extension at 72 °C for 30 s. The PCR product was quantified by the intensity of SYBR Green I fluorescence at 83 °C.

Western Blot Analysis—Cell monolayers were rinsed twice with 1x PBS, directly lysed in the plate on ice with radioimmune precipitation assay buffer containing Tris-HCl (50 mM, pH 7.4), NaCl (150 mM), Nonidet P-40 (1%), sodium deoxycholate (0.5%), SDS (0.1%), 1 µM phenylmethylsulfonyl fluoride, and complete Protease Inhibitor Mixture. Cell debris was pelleted by centrifugation at 14,000 rpm for 15 min, and the supernatant was collected for protein quantification. The bicin-choninic acid protein assay (Pierce) was used to estimate protein concentration according to the manufacturer's instructions. 50 µg of protein were diluted with 4x SDS sample loading buffer, boiled for 5 min, and separated by SDS-polyacrylamide gels. Following electrophoresis, separated proteins were transferred onto nitrocellulose membranes. The membranes were then blocked with 5% milk/PBS and incubated with the indicated primary antibodies. After incubation with the primary antibodies, membranes were washed thoroughly in TBS-Tween buffer (25 mM Tris, pH 8.0, 125 mM NaCl, 0.1% Tween 20). Appropriate secondary antibodies conjugated with horseradish peroxidase were used to detect the primary antibodies. Immunoreactive bands were visualized by chemiluminescence in signaling solution (Pierce).

ERK1/2 Activation, EGF Receptor Activation, PPAR Phosphorylation—To investigate the effects of gastrin on ERK1/2 activation, EGFR transactivation, or PPAR phosphorylation, DLD-1 cells were incubated in the presence or absence of 200 nM G-17. At different time points, whole cell lysates were collected, and Western analysis was performed with specific antibodies. For identification of the active form of ERK1/2, an antibody specifically recognizing ERK1 and ERK2 phosphorylated at Thr²⁰² and Tyr²⁰⁴, respectively, was utilized. To determine the involvement of the EGFR in ERK1/2 activation, DLD-1 cells were preincubated for 30 min with the EGFR antagonist AG1478 prior to the addition of G-17. To evaluate EGFR transactivation by gastrin, Western analysis was performed with a monoclonal antibody specifically immunoreactive with the EGFR phosphorylated at Tyr¹¹⁷³ to detect the activated form of the EGFR. To examine PPAR phosphorylation, Western analysis was performed using a rabbit polyclonal antibody specifically immunoreactive with PPAR phosphorylated at Ser⁸⁴. Protein loading was normalized by the measurement of -actin.

PPAR Stability—To examine the effects of gastrin on PPAR protein stability, DLD-1 cells were incubated with the protein synthesis inhibitor cycloheximide (50 µg/ml) in the presence or absence of 200 nM G-17. At different time points, whole cell lysates were collected, and Western analysis was performed using the anti-PPAR antibody to detect PPAR, with -actin measured as a loading control. Linear regression analysis was performed to determine the half-life of PPAR in the presence and absence of gastrin. Values were normalized to -actin and plotted on a log versus time scale. To ascertain whether PPAR protein was degraded through the proteasomal pathway, cells were preincubated with the proteasomal inhibitor, MG132 (20 µM) or lactacystin (5 µM), for 30 min prior to the addition of 200 nM G-17. To determine whether ERK1/2 and EGFR activation might affect PPAR half-life, DLD-1 cells were preincubated with 40 µM PD98059, a potent MEK1 inhibitor, to inhibit the downstream activation of ERK1/2, with 100 nM AG1478, or with 10 µM SB20358, the p38 MAPK inhibitor, for 30 min prior to addition of cycloheximide.

Statistical Analysis—All results are expressed as the mean ± S.D. Statistical analysis was performed using analysis of variance and Student's t test. A p value <0.05 was considered to be statistically significant.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Effects of PPAR and Gastrin on Cellular Proliferation—To determine the effect of gastrin on cellular proliferation, DLD-1 cells were incubated in the presence and absence of 50 or 200 nM G-17 for 24 h, and [³H]thymidine uptake was measured. [³H]Thymidine uptake was increased by 35.4 ± 7.3% (p < 0.05) and 51 ± 12.4% (p < 0.01) by 50 nM and 200 nM G-17 treatment, respectively. In the presence of the CCK-2R antagonist, L-365,260, the increase in cell proliferation induced by 200 nM G-17 was attenuated by 60% to 20 ± 9.2% of control values (p < 0.01 compared with 200 nM G-17, p < 0.01 compared with vehicle treatment alone) (Fig. 1). Similar results were obtained from cell counting experiments and the MTT proliferation assay (data not shown).

In separate experiments, DLD-1 cells were incubated in the presence of increasing concentrations of two known PPAR agonists, ciglitazone and rosiglitazone, to determine the effect of PPAR activation on cellular proliferation. Both ciglitazone and rosiglitazone treatment suppressed cell growth in a concentration-dependent manner. Rosiglitazone treatment decreased cellular proliferation at concentrations as low as 1 µM (by 18.4 ± 6.7%, p < 0.05), and maximal inhibition began to plateau from 5 µM (69.4 ± 10.2% of control, p < 0.01) to 10 µM (64.8 ± 8.6% of control, p < 0.01) (Fig. 2a). Similarly, ciglitazone treatment decreased cellular proliferation at concentrations as low as 1 µM (by 13.8 ± 6.6%, p < 0.05), with maximal inhibition achieved at a concentration of 10 µM (71.3 ± 3.4% of control, p < 0.01) (Fig. 2b). In the presence of 200 nM G-17, PPAR growth suppression induced by 10 µM ciglitazone and 10 µM rosiglitazone was significantly attenuated to 93.8 ± 5.3% of control (p < 0.05) (Fig. 2b) and 89.9 ± 6.8% of control (p < 0.05) (Fig. 2a), respectively.

View larger version (15K): [in this window] [in a new window]   FIGURE 1. Gastrin promotes DLD-1 CRC cell proliferation. Gastrin stimulates DNA synthesis, as measured by [3H]thymidine uptake. Following serum starvation, DLD-1 cells were incubated in the presence of 50 nM G-17, 200 nM G-17, or vehicle alone for 22 h. To determine the involvement of the CCK-2R, DLD-1 cells were preincubated with L-365,260 30 min prior to addition of 200 nM G-17. Values are expressed as a mean percent of control (vehicle) ± S.D. of at least three independent experiments performed in triplicate. *, p < 0.05; , p < 0.01.

View larger version (27K): [in this window] [in a new window]   FIGURE 2. Ligand-dependent PPAR growth suppression is attenuated by pretreatment with gastrin. DLD-1 cells were serum-starved for 24 h followed by 12 h incubation in medium containing 200 nM G-17 or vehicle alone. Following the 12-h incubation period, rosiglitazone (0–20 µM) in a, or ciglitazone (0–20 µM) in b, was added and incubated for 72 h, at which time cell proliferation was measured by the MTT assay method, as described under "Experimental Procedures." Rosiglitazone in a, and ciglitazone in b, dose-dependently suppressed DLD-1 cell proliferation. In the presence of 200 nM G-17, PPAR growth suppression was attenuated. Data are represented as mean percent of control (no rosiglitazone or ciglitazone) ± S.D. of three independent experiments performed in sixteen replicates. *, p < 0.05 compared with control; , p < 0.01 compared with control. +, p < 0.05 compared with 10 µM rosiglitazone or ciglitazone.

View larger version (17K): [in this window] [in a new window]   FIGURE 3. Gastrin attenuates ligand-dependent PPAR activation. DLD-1 cells were co-transfected with the pHD(x3)Luc and pCMV--gal. Then DLD-1 cells were incubated for 12 h in the presence or absence of 200 nM G-17 in medium containing 1% FBS. Ciglitazone (0–10 µM) was then added and incubated for 16 h, after which luciferase and -galactosidase activities were measured. Ciglitazone promoted a dose-dependent increase in luciferase activity, which was attenuated by G-17 pretreatment. Data are expressed as mean-fold activation ± S.D. compared with control (no ciglitazone) of three independent experiments performed in triplicate. *, p < 0.001 compared with control; , p < 0.01 compared with respective ciglitazone concentrations (1 µM, 5 µM, 10 µM).

Effects of Gastrin on PPAR Expression—We next examined whether gastrin might affect PPAR protein expression. DLD-1 cells were incubated in the presence or absence of G-17 (50 nM or 200 nM) for 12, 24, and 48 h, at which time PPAR protein levels were measured by Western analysis. As shown in Fig. 4a, in response to the incubation of DLD-1 cells in the presence of G-17, PPAR protein levels decreased when compared with control levels at all indicated time points. In addition, the decrease in PPAR protein levels was concentration-dependent, with more pronounced reductions detected in the presence of 200 nM G-17. To determine whether the gastrin-promoted decrease in PPAR protein concentrations occurred as a result of a decrease in PPAR gene expression, DLD-1 cells were incubated in the presence of 50 nM and 200 nM G-17 or vehicle alone for 3, 6, 12, and 24 h, at which time total RNA was extracted and quantitative PCR analysis performed. No significant differences in PPAR gene expression were detected in the presence of G-17 when compared with vehicle treatment at any of the above time points (data not shown).

Two additional CRC cell lines, SW48 and MC-26, were employed to determine whether the reduction in PPAR protein levels induced by G-17 was cell-specific. As shown in Fig. 4b, 24-h incubation with 200 nM G-17 significantly diminished PPAR protein levels in both SW48 and MC-26 CRC cells. Therefore, the effects of gastrin on PPAR expression observed in DLD-1 cells, a human CRC cell line possessing an APC mutation, appear to extend to a murine CRC cell line (MC-26) and to a human CRC cell line possessing the wild-type APC phenotype (SW48).

Because gastrin decreased PPAR protein levels in the absence of exogenous PPAR ligands, we next examined the effect of gastrin on PPAR protein levels in the presence of the PPAR ligand, rosiglitazone. When DLD-1 cells were incubated with 5 µM rosiglitazone, PPAR protein levels slightly increased when compared with untreated cells (Fig. 4c). The addition of 200 nM G-17 to the culture medium containing 5 µM rosiglitazone significantly diminished PPAR protein levels, whereas PPAR levels decreased to a greater extent when cells were incubated in the presence of 200 nM G-17 alone (Fig. 4c).

View larger version (70K): [in this window] [in a new window]   FIGURE 4. Gastrin promotes a concentration-dependent decrease in PPAR protein expression. a, following serum starvation, DLD-1 cells were incubated in the presence of vehicle alone, 50 nM G-17, or 200 nM G-17 for the indicated time periods. b, following serum starvation, SW48 and MC-26 cells were incubated in the presence of vehicle alone or 200 nM G-17 for 24 h. c, following serum starvation, DLD-1 cells were incubated in the presence of vehicle alone or 200 nM G-17 and in the presence of absence of 5 µM rosiglitazone for 24 h. At the indicated time points, cell lysates were collected, resolved by SDS-PAGE, and immunoblotted with anti-PPAR antibody. -Actin was used as a control for loading. Top immunoblots are representative of at least three independent experiments.

View larger version (36K): [in this window] [in a new window]   FIGURE 5. Gastrin decreases the stability of PPAR protein. a, following serum starvation, DLD-1 cells were incubated with vehicle alone or 50 nM, 100 nM, or 200 nM G-17 in the presence of 50 µg/ml cycloheximide for 12 h. Whole cell lysates were then collected, resolved by SDS-PAGE, and immunoblotted for PPAR. -Actin levels were measured as a control for loading. G-17 increased PPAR protein degradation in a concentration-dependent manner. b, following serum starvation, DLD-1 cells were incubated over a 12-h period in the presence or absence of 200 nM G-17 in medium containing 50 µg/ml cycloheximide. Whole cell lysates were collected at the indicated time points, resolved by SDS-PAGE, and immunoblotted with the anti-PPAR antibody. -Actin levels were measured as a control for loading. As shown in the lower panel, gastrin decreased the half-life of PPAR when compared with untreated cells represented in the upper panel. c, linear regression analysis of data from six experiments was plotted on a log scale to determine the half-life of PPAR in the presence (*, p < 0.05) or absence of 200 nM G-17. The t of PPAR protein is decreased from 11.3 h (sl–G-17 = –4.43, r2 = 0.993) in the absence of G-17 to 7.1 h in the presence of 200 nM G-17 (sl+G-17 = –7.02, r2 = 0.981, p < 0.05 versus sl–G-17).

To further investigate the role of the proteasome in PPAR degradation, DLD-1 cells were incubated in the presence of two 26 S proteasomal inhibitors, lactacystin or MG132. In the presence of 5 µM lactacystin or 20 µM MG132, both basal and gastrin-induced PPAR degradation were markedly inhibited (Fig. 6). These results are consistent with the hypothesis that a decrease in PPAR protein levels following gastrin treatment is mediated by targeting PPAR for proteasomal degradation.

Gastrin Transactivation of the EGF Receptor—Earlier studies have demonstrated the involvement of many growth factors in the transactivation of the EGFR (41–49). However, gastrin transactivation of the EGFR has not been previously examined in CRC. To evaluate the role of EGFR transactivation by gastrin in CRC, DLD-1 cells were incubated in the presence of 200 nM G-17 for 2 h, and whole cell lysates were collected at various time points. The lysates were evaluated by Western analysis using an antibody against the phosphorylated, active form of the EGFR. As shown in Fig. 7, G-17 promoted a biphasic activation of the EGFR. An initial increase in EGFR phosphorylation was observed after 5 min, which then declined after 30 min to a level slightly above control. The second component of the biphasic activation of the EGFR was detected at 60 min and remained steadily up-regulated for up to 2 h.

View larger version (42K): [in this window] [in a new window]   FIGURE 6. Gastrin promotes PPAR degradation through the proteasomal pathway. Following serum starvation, DLD-1 cells were preincubated with or without the proteasomal inhibitors, MG132 and lactacystin. Cells were then treated with 200 nM G-17 or vehicle alone in medium containing cycloheximide for 12 h. Top immunoblot is representative of three independent experiments.

View larger version (47K): [in this window] [in a new window]   FIGURE 7. Gastrin promotes transactivation of the EGF receptor. a, shown is a time course of EGFR transactivation by 200 nM G-17. Following serum starvation, DLD-1 cells were incubated in the presence of 200 nM G-17 over a 2-h period. At the indicated time points, whole cell lysates were collected, separated by SDS-PAGE, and immunoblotted with the phosphospecific anti-EGFR (pTyr1173) antibody. -Actin protein levels were measured as a control for loading. b, graphical representation of data from three experiments depicted in a. Values are normalized to -actin and are expressed as fold-activation ± S.D. of control (vehicle). *, p < 0.05 compared with control.

Past studies have demonstrated that ERK1/2 activation may occur, in part, through transactivation of the EGFR by G-protein-coupled receptors (10–18). Because ERK1/2 activation by gastrin nearly paralleled gastrin-stimulated EGFR transactivation in our study, we hypothesized that gastrin may promote activation of ERK1/2 through transactivation of the EGFR. To examine the role of gastrin-stimulated transactivation of the EGFR in ERK1/2 activation, DLD-1 cells were pretreated with the EGFR kinase inhibitor AG1478, which inhibited gastrin-stimulated EGFR phosphorylation (Data not shown). Moreover, in the presence of AG1478, activation of ERK1/2 by G-17 was nearly abolished at all observed time points when compared with the activated ERK1/2 levels in cells treated with G-17 alone (Fig. 8). A slight up-regulation of ERK2 activation still remained after 5 min and 120 min when compared with the levels evaluated at 0 min (Fig. 8). These results suggest that gastrin-induced EGFR transactivation may play a role in ERK1/2 activation.

View larger version (48K): [in this window] [in a new window]   FIGURE 8. Gastrin-stimulated ERK1/2 activation is mediated, in part, through the EGF receptor. After serum starvation, DLD-1 cells were incubated with 200 nM G-17 over 2 h. At the indicated time points, whole cell lysates were collected, separated by SDS-PAGE, and immunoblotted with the phosphospecific ERK1/2 monoclonal antibody (pThr202/Tyr204). To determine the involvement of the EGFR, cells were preincubated with 100 nM AG1478 prior to addition of 200 nM G-17. -Actin was measured as a control for loading. Top immunoblot is representative of at least three independent experiments.

Because gastrin stimulated ERK1/2 activation and appeared to promote EGFR transactivation, we examined the possibility that these two pathways may also play a role in gastrin-stimulated PPAR phosphorylation by utilizing AG1478 and PD98059 to inhibit EGFR and ERK1/2 activity, respectively. DLD-1 cells were pretreated individually with these inhibitors for 45 min prior to addition of 200 nM G-17. As depicted in Fig. 9b, in the presence of AG1478, gastrin-stimulated PPAR phosphorylation at Ser⁸⁴ was significantly diminished. Similarly, PPAR phosphorylation was abolished by co-incubation with PD98059 (Fig. 9b). Moreover, gastrin-induced ERK1/2 activation was abolished by the co-incubation of DLD-1 cells with either AG1478 or PD98059 (Fig. 9b). These results suggest that gastrin-enhanced PPAR phosphorylation at Ser⁸⁴ is mediated, in part, through EGFR transactivation and ERK1/2 activation.

The Roles of EGFR and ERK1/2 Activation in PPAR Degradation by Gastrin—Floyd and Stephens (40) have demonstrated that ERK1/2 activation is involved in IFN -promoted PPAR degradation in adipocytes. However, the role of MAPK phosphorylation in the regulation of PPAR has not been evaluated in neoplasia. To examine the roles of gastrin-stimulated EGFR and ERK1/2 activation in PPAR degradation, we employed PD98059 to inhibit ERK1/2 activation and AG1478 to inhibit EGFR activity and used Western analysis to measure PPAR protein levels. We also used SB203580, a p38 MAPK inhibitor, as a negative control because this MAPK has not been demonstrated to play a major role in PPAR phosphorylation (20). DLD-1 cells were pretreated with the inhibitors individually for 45 min prior to the addition of cycloheximide and vehicle alone or 200 nM G-17. In the presence of PD98059, G-17-induced PPAR degradation was abrogated when compared with G-17 treated cells alone (Fig. 10). In addition, EGFR inhibition by AG1478 also abolished gastrin-induced PPAR degradation (Fig. 10). As predicted, SB203580 had no significant effect on gastrin-stimulated PPAR degradation (Fig. 10). These results suggest that gastrin may promote PPAR degradation through activation of ERK1/2 and transactivation of the EGFR, without involvement of the p38 MAPK pathway.

View larger version (71K): [in this window] [in a new window]   FIGURE 9. Gastrin-stimulated PPAR phosphorylation is mediated, in part, through transactivation of the EGF receptor and activation of ERK1/2. a, following 24 h of serum starvation, DLD-1 cells were incubated with 200 nM G-17 over a 60-min period. b, to determine the EGFR and ERK1/2 involvement, DLD-1 cells were preincubated with 100 nM AG1478 or 40 µM PD98059, respectively, for 30 min prior to addition of 200 nM-G17. After 5 min of incubation with G-17, whole cell lysates were collected for a determination of phospho-PPAR and phospho-ERK1/2 protein levels. At the indicated time points, whole cell lysates were collected, separated by SDS-PAGE, and immunoblotted with the phosphospecific PPAR antibody (pSer84) and the phosphospecific ERK1/2 antibody (pThr202/Tyr204). -Actin was measured as a control for loading. Immunoblots are representative of at least three independent experiments.

View larger version (56K): [in this window] [in a new window]   FIGURE 10. Inhibition of ERK1/2 activation and EGF receptor transactivation attenuates gastrin-induced PPAR degradation. a, after serum starvation DLD-1 cells were preincubated with 100 nM AG1478, 40 µM PD98059, or 10 µM SB203580 for 30 min. 200 nM G-17 or vehicle alone was then added in the presence of the above inhibitors in medium containing 50 µg/ml cycloheximide. After 12 h, cell monolayers were lysed, resolved by SDS-PAGE, and immunoblotted with the anti-PPAR antibody and the phos-phospecific ERK1/2 antibody (pThr202/Tyr204). b, graphic representation of data of five experiments depicted in a. Values are normalized to -actin and are expressed as a mean percent ± S.D. of control (no treatment). *, p < 0.001 compared with control; , p < 0.001 compared with 200 nM G-17 treatment alone; N.S., not significant compared with 200 nM G-17 treatment alone.

Mutation of the Ser⁸⁴ Phosphorylation Site Attenuates the Effects of Gastrin on Ligand-dependent PPAR Growth Suppression—We next determined whether mutation of the Ser⁸⁴ phosphorylation site of PPAR might also affect the attenuation of PPAR-induced growth suppression by gastrin. 5 µM rosiglitazone decreased DLD-1 cell proliferation to 64% of control (p < 0.001) in DLD-1 cells overexpressing wtPPAR, and to 66% (p < 0.001) in cells overexpressing mutant PPAR S84A, when compared with their respective controls (Fig. 11d). Moreover, in response to gastrin incubation, rosiglitazone-induced growth suppression was diminished by 64% (p < 0.001 versus 5 µM rosiglitazone alone) in wtPPAR-overexpressing cells. In contrast, in the presence of 200 nM G-17, PPAR growth suppression was minimally attenuated in cells overexpressing mutPPAR S84A (Fig. 11d). These results indicate that gastrin appears to attenuate rosiglitazone-induced PPAR growth suppression, at least in part, through phosphorylation of PPAR at Ser⁸⁴.

View larger version (44K): [in this window] [in a new window]   FIGURE 11. Mutation of the Ser84 phosphorylation site reverses the attenuation of ligand-dependent PPAR activity and growth suppression by gastrin. a, confirmation of stable expression of PPAR variants. 1, parental DLD-1; 2, DLD-1 pTracer mutPPAR S84A; 3, DLD-1 pTracer wtPPAR; 4, DLD-1 pTracer empty vector. b, DLD-1 cells were co-transfected with the pHD(x3)Luc and pCMV--gal, after which DLD-1 cells were incubated for 12 h in the presence or absence of 200 nM G-17. Rosiglitazone (0–10 µM) was then added and incubated for 16 h, after which luciferase activity was measured. -Galactosidase activity was measured as a control. Rosiglitazone promoted an increase in luciferase activity, which was attenuated by G-17 pretreatment in cells overexpressing wtPPAR but not in cells overexpressing mutPPAR S84A. Data are expressed as mean-fold activation ± S.D. compared with control (no rosiglitazone) of three independent experiments performed in triplicate. c, DLD-1 cells overexpressing wtPPAR or mutPPAR S84A protein were serum-starved for 24 h and then incubated in the presence of 50 µg/ml of cycloheximide with or without 200 nM G-17. Cell monolayers were lysed, resolved by SDS-PAGE, and immunoblotted with the anti-PPAR antibody and -actin was measured as a loading control. Immunoblot is representative of three independent experiments. d, DLD-1 cells were serum-starved for 24 h, followed by a 6-h incubation in medium containing 200 nM G-17 or vehicle alone. Rosiglitazone (0–10 µM) was then added and incubated for 72 h, at which time cell proliferation was measured by the MTT assay method, as described under "Experimental Procedures." Rosiglitazone suppressed DLD-1 cell proliferation. In the presence of 200 nM G-17, PPAR growth suppression was significantly attenuated in cells overexpressing wtPPAR, but to a much lesser extent in cells overexpressing mutPPAR S84A. Data are represented as a mean percent of control (no rosiglitazone) ± S.D. of three independent experiments performed in 24 replicates. *, p < 0.001 compared with control; , p < 0.001 compared with 5 µM rosiglitazone. e, DLD-1 cells overexpressing pTracer alone or mutPPAR S84A were serum-starved for 24 h and then incubated with or without 200 nM G-17. Cell proliferation was measured by the MTT assay method, as described under "Experimental Procedures." Overexpression of PPAR attenuated G-17-induced cell proliferation. Data are represented as a mean percent of control (vehicle) ± S.D. of three experiments performed in 16 replicates. *, p < 0.01 compared to vehicle treatment; #, p < 0.01 compared with 200 nM G-17 alone.

Glycine-extended Gastrin Promotes Proliferation by Decreasing PPAR Protein Levels—Studies were next performed to determine whether, similar to -amidated gastrin, glycine-extended gastrin might also promote DLD-1 cell proliferation by decreasing PPAR protein levels. DLD-1 cells were incubated in the presence of 1 nM – 100 nM Gly-G. As shown in Fig. 12a, Gly-G promoted a concentration-dependent increase in DLD-1 cell proliferation, as determined using the MTT proliferation assay. DLD-1 cell growth increased by 15.0 ± 2.2% and 26.1 ± 2.2% in the presence of 1 nM and 10 nM Gly-G, respectively, with maximal proliferation detected with 50 nM Gly-G (49.5 ± 1.5%) and 100 nM Gly-G (50.1 ± 1.8%). To determine whether the mitogenic properties of Gly-G might be mediated, in part, by decreasing PPAR levels, DLD-1 cells overexpressing the empty vector, wild-type PPAR, or mutPPAR S84A were incubated in the presence of 50 nM Gly-G for 24 h. As depicted in Fig. 12b, 50 nM Gly-G decreased PPAR protein levels in the cell lines overexpressing the empty vector or wild-type PPAR. In contrast, no change in PPAR protein levels was observed in the cells overexpressing mutPPAR S84A. These observations suggest that Gly-G may likewise promote the growth of DLD-1 colorectal cancer cells, in part, by decreasing PPAR protein levels through the promotion of PPAR phosphorylation at Ser⁸⁴.

View larger version (42K): [in this window] [in a new window]   FIGURE 12. Gly-G promotes cell proliferation, in part, by attenuation of PPAR. a, Gly-G stimulates DLD-1 cell proliferation, as measured by the MTT assay. Following serum starvation, DLD-1 cells were incubated in the presence of 1 nM to 100 nM Gly-G, and proliferation was measured by the MTT assay. Values are expressed as a mean percent of control (vehicle) ± S.D. of at least three independent experiments performed in 24 replicates. *, p < 0.001 compared with vehicle. b, DLD-1 cells overexpressiong wtPPAR or mutPPAR S84A protein were serum-starved for 24 h and then incubated in the presence or absence of 50 nM Gly-G. Cell monolayers were lysed, resolved by SDS-PAGE, and immunoblotted with the anti-PPAR antibody and -actin was measured as a loading control. c, DLD-1 cells were serum-starved for 24 h, followed by a 6-h incubation in medium containing 50 nM Gly-G or vehicle alone. Rosiglitazone (0–10 µM) was then added and incubated for 72 h, at which time cell proliferation was measured by the MTT assay method, as described under "Experimental Procedures." Rosiglitazone suppressed DLD-1 cell proliferation. In the presence of 50 nM Gly-G, PPAR growth suppression was significantly attenuated in cells overexpressing wtPPAR, but to a much lesser extent in cells overexpressing mutPPAR S84A. Data are represented as a mean percent of control (no rosiglitazone) ± S.D. of two independent experiments performed in 24 replicates. *, p < 0.01 compared with control; , p < 0.01 compared with 5 µM rosiglitazone; N.S., not significant compared with 5 µM rosiglitazone.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Recent reports have indicated that both hypergastrinemia and decreased expression of PPAR are associated with an increased risk for the development of CRC (10, 37, 50, 51). However, the possibility of a link between gastrin and PPAR has not been reported previously. In the present study, we have reported for the first time that ligand-dependent PPAR growth suppression is attenuated by gastrin, at least in part, through attenuation of PPAR activity and through an increase in PPAR protein degradation. In addition, gastrin-induced EGFR transactivation and ERK1/2 activation appear to play a central role in this process.

Gastrin has been implicated as a trophic factor for various neoplasms of GI origin, including CRC and those of pancreatic, gastric, and esophageal origin (52–62). Gastrin and gastrin receptors are expressed aberrantly in colorectal adenomas and malignant tumors and have been implicated in malignant progression (17, 18). While Smith and Watson (17) and Guo et al. (46) reported that a majority (up to 80%) of all colorectal adenomas express the CCK-2R, others have detected the receptor in only 11–38% of such tumors (15, 63). These discrepancies may be explained, at least to some extent, by the detection of splice variants of the CCK-2R and by differences in methods utilized by various investigators.

The results of the present study demonstrate that gastrin and its precursor, Gly-G, stimulated the growth of DLD-1 cells in a concentration-dependent manner. Interestingly, the specific CCK-2 receptor antagonist L-365,260 only partially attenuated G-17-stimulated growth of DLD-1 cells. These findings suggest that gastrin-stimulated cell growth in these cells may be mediated, at least in part, by other receptors that bind gastrin peptides. In addition to the "classical" gastrin receptor binding site, several alternative binding sites on DLD-1 cells have been proposed. Ahmed et al. (64) recently identified two gastrin binding sites with nanomolar and micromolar affinities, respectively, on the cell membrane, both of which played key roles in cell proliferation stimulated by amidated and precursor gastrin peptides. Furthermore, Yang et al. (65) identified a micromolar affinity site for both G-17 and the non-amidated glycine-extended gastrin precursors on DLD-1 cells. Thus, it appears that the activation of alternative gastrin receptors, in addition to the "classical" CCK-2R, might mediate gastrin-stimulated proliferation of DLD-1 cells.

The activation of ERK1/2 is a known pathway by which gastrin stimulates cell proliferation (42, 46, 66, 67). Although several studies have demonstrated that gastrin promotes ERK1/2 activation through transactivation of the EGFR, many of them have been performed in cell lines ectopically overexpressing the CCK-2R (42, 45, 46, 60, 62). The present study represents the first to demonstrate that gastrin promotion of ERK1/2 activation involves EGFR transactivation in human CRC cells expressing endogenous gastrin receptors. Our findings corroborate an earlier study by Guo et al. (46) who reported that gastrin activates ERK1/2 in a biphasic manner in rat intestinal epithelial (RIE) cell lines ectopically expressing the CCK-2R (46). However, in their study, EGFR transactivation appeared to promote only the later phase of ERK1/2 activation. In our study, EGFR transactivation played a key role in both early and late ERK1/2 activation, as demonstrated by significant inhibition of both phases in the presence of the EGFR kinase inhibitor, AG1478. In addition, the biphasic activation of ERK1/2, although not identical, paralleled that of EGFR transactivation in the presence of gastrin, further suggesting the involvement of the EGFR in the early and late phases of ERK1/2 activation. Moreover, these results are consistent with the hypothesis that gastrin-stimulated EGFR transactivation promotes ERK1/2 activation, which in turn appears to play a critical role in mediating the trophic properties of gastrin in CRC.

Nevertheless, the possibility of the involvement of additional pathways independent of the EGFR in the regulation of gastrin-enhanced phosphorylation of ERK1/2 cannot be excluded. As indicated above, gastrin-stimulated phosphorylation of the EGFR exhibited a similar profile to that of ERK1/2, but was not identical. For example, at 15 min, ERK1/2 phosphorylation was diminished while EGFR phosphorylation was increased. Furthermore, residual ERK1/2 activation was evident at 5 and 120 min the presence of AG1478. Thus, it is likely that gastrin promotes ERK1/2 phosphorylation by additional pathways not involving the EGFR, including direct stimulation of MAP kinase pathways (42, 46, 66, 67).

Although not examined in the present study, several mechanisms responsible for gastrin-stimulated EGFR transactivation have been suggested in other cell lines overexpressing the CCK-2R.<