Background
Caveolae are flask-shaped vesicular invaginations of the plasma membrane characterized by the existence of integral membrane proteins termed caveolins. Caveolae is implicated in many cellular functions, including membrane trafficking, endocytosis, lipid metabolism, cell adhesion, signal transduction in cellular proliferation and apoptosis [
1]. Caveolins are a family of proteins composed of three isoforms, Caveolin (Cav)-1, −2, and −3. Among the three caveolins, Cav-1 is a principal structural component of caveolae and forms a high molecular complex of homo-oligomer or hetero-oligomer with Cav-2. A scaffolding domain within Cav-1 allows this protein to interact with signaling molecules, including growth factor receptors, G-protein coupled receptors, small GTPases, Src kinases, nitric oxide synthases, and integrins [
2]. Integrations and complex formation of Cav-1 with signaling molecules functionally affect the activity of these molecules.
Despite a growing body of evidence on Cav-1 implication in tumorigenesis, its role in tumor growth and underlying molecular mechanisms remain largely undefined. Both tumor suppression and promotion roles of Cav-1 have been proposed on the basis of its expression status detected in cancers. Cav-1 expression is frequently down-regulated in many human cancers mainly due to promoter hypermethylation whereas its elevation correlates with enhanced progression, multidrug resistance, and metastatic potentials of certain tumors [
3‐
6]. Furthermore,
Cav-1 gene amplification and mutation were reported in a subgroup of breast cancers [
7,
8]. These findings demonstrate that Cav-1 has differential functions in tumorigenesis depending on the types, origins, or genetic contexts of tumors.
Caveolae have been proposed to be the site of epidermal growth factor receptor (EGFR) signaling, including EGFR autophosphorylation [
9]. EGF-induced tumor cell proliferation and migration is suppressed when Cav-1 binds to the EGFR, suggesting that Cav-1 may play a role in maintaining the EGFR in an inactive state, with dissociation from Cav-1 promoting EGFR activation [
10]. It was also shown that many components of Ras signaling, including RAF, MEK, and ERK appear to be compartmentalized within caveolin-rich membrane domains and that Cav-1 downregulation results in constitutive activation of ERK signaling while activation of Ras-ERK signaling causes Cav-1 reduction [
11,
12]. In contrast, Cav-1 appears to promote metastasis of Ewing sarcoma and the proliferation of metastatic lung cancer cells through activation of the MAPK-ERK pathway [
13,
14]. A recent study also showed that Cav-1 is required for kinase suppressor of Ras 1 (KSR1)-mediated ERK1/2 activation, Ras-induced senescence, and transformation [
15]. These findings thus indicate that Cav-1 functions as an endogenous inhibitor or stimulator of the Ras-ERK cascade. Nevertheless, the molecular basis for the opposite effects of Cav-1 on EGFR and Ras-MAPK signaling and its implication in tumorigenesis remains largely undefined.
Gastric cancer is one of the most commonly diagnosed malignancies worldwide and a leading cause of cancer mortality in certain areas such as Korea, Japan, South America, and Eastern Europe [
16,
17]. Although a number of study indicates that genetic and/or epigenetic alterations of multiple genes, such as
p53,
K-Ras, and
E-Cadherin are associated with the development and progression of gastric cancers, molecular events that drive the neoplastic process remain to be characterized [
18]. In this study, we found that
Cav-1 is abnormally down- and up-regulated in a considerable fraction of gastric cancers due to promoter hyper- and hypo-methylation, respectively. In low- and high-expressing tumor cells, Cav-1 evokes the opposite effects on cell proliferation and colony formation through the reciprocal control on the RAF-ERK negative feedback loop. Therefore, our study demonstrates that Cav-1 acts as a positive or negative regulator of the RAF-ERK feedback loop and that the mitogenic switch of Cav-1 function is highly associated with bidirectional alteration of its expression in tumor progression.
Methods
Tissues specimens and cell lines
Total 180 gastric tissues including 100 primary carcinomas, 4 adenomas, 6 hamartomas, 6 hyperplastic polyps, and 64 normal gastric tissues were obtained were obtained from 100 gastric cancer patients and 80 noncancer patients by surgical resection in the Kyung Hee University Medical Center (Seoul, Korea). Signed informed consent was obtained from each patient. Tissue specimens were snap-frozen in liquid N2 and stored at −70 °C until used. Tissue slices were subjected to histopathological review and tumor specimens composed of at least 70% carcinoma cells and adjacent tissues found not to contain tumor cells were chosen for molecular analysis. Fourteen human gastric cancer cell lines (SNU5, SNU16, SNU216, SNU484, SNU601, SNU620, SNU638, SNU719, MKN1, MKN28, MKN45, MKN74, AGS, and KATO-III) were obtained from Korea Cell Line Bank (Seoul, Korea) or American Type Culture Collection (Rockville, MD).
Quantitative RT- and genomic PCR
RNA extraction and cDNA synthesis were performed as described previously [
19]. Briefly, 1 μg of total cellular RNA was converted to cDNA using random hexamer primers and M-MLV reverse transcriptase (Invitrogen Corporation, Carlsbad, CA). PCR was initially carried out over 24–40 cycles and 12.5 ng cDNA (50 μl PCR reaction) undergoing 30–36 cycles showed logarithmic amplification with primers Cav-1S/Cav-1AS for
Cav-1, C1αA/C1αAS for
Cav-1α, C1β/C1βAS for
Cav-1β, Cav-2S/Cav-2AS for
Cav-2, C2αA/C2αAS for
Cav-2α, C2βA/C2βAS for
Cav-2β, and G2/G3 for an endogenous expression standard gene
GAPDH (Table
1). PCR was done in 1.5 mM MgCl
2-containing reaction buffer (PCR buffer II) (Perkin Elmer, Branchburg, NJ) and 10 μl of PCR products were resolved on 2% agarose gels. Quantitation was achieved by densitometric scanning of the ethidium bromide-stained gels. Integration and analysis was performed using Molecular Analyst software program (Bio-Rad, Richmond, CA). For genomic PCR, intron 2 regions of
Cav-1 and
Cav-2 and intron 5 region of
GAPDH were amplified with intron-specific primers RF2S/RF2AS and G3/G5, respectively (Table
1). Quantitative PCR was repeated at least three times for each specimen and the mean was obtained.
Table 1
Primers used for PCR, LOH and bisulfite sequencing analysis
Cav1 | Cav-1S | TCTGGGGCGTCGTGCGCAAA |
Cav-1AS | GAACCTTGATGAAGCCTGTG |
C1⍺A | AGTTTTCATCCAGCCACGGG |
C1⍺AS | TCTTGACCACGTCATCGTTGAG |
C1βA | CATTTTTCCTCCCACCGCCGTT |
C1βAS | AAAACTGTGTGTCCCTTCTG |
RF2S | ATGTATATGTACATCAGGGA |
RF2AS | CAGGCACATAGCTGGGTACC |
SNP-1 | GGCTCAACATTGTGTTCCCATTTCAGC |
SNP-2 | GTGTCAGGAAGACTGGAAGAGGCA |
P1 | TGTGTATTTTGTAAATATGGTATA |
P2 | AAGTTAAAGATTTTTATTTTTTATT |
Cav2 | Cav-2S | ATCTGCAGCCATGCCCTCTTTG |
Cav-2AS | GGGTCCAAGTATTCAATCCTGG |
C2⍺A | ATGGGGCTGGAGACGGAGAA |
C2⍺AS | ACTGAAGGCAGAACCATTAGGCA |
C2βA | TGCGTCCTGTCTCCTCAGCT |
C2βAS | ACTGAAGGCAGAACCATTAGGCA |
GAPDH | G2 | CATGTGGGCCATGAGGTCCACCAC |
G3 | AACCATGAGAAGTATGACAACAGC |
G5 | GAGTCCTTCCACGATACCAAAG |
Loss of heterozygosity (LOH) analysis
LOH of the
Cav-1 gene was determined using an intraexonic SNP (5′-AGCATC
C/
T-3′) located at +2061 nucleotide (exon 3) from the transcription start site. PCR was performed on each tumor and normal DNA sample pair obtained from 50 patients using primers SNP-1/SNP-2 (Table
1). Five μl of the PCR products were used for cutting with the endonuclease
BtsCI (NEB, Beverly, MA) and enzyme-digested PCR products were electrophoresed on 2% agarose gels. Signal intensity of fragments and the relative ratio of tumor and normal allele intensities were determined by scanning densitometry.
To screen the presence of somatic mutations, RT-PCR-SSCP analysis of Cav-1 and Cav-2 was performed using 3 sets of primers that were designed to cover the entire coding region of the genes. Twenty μl of PCR products mixed with 10 μl of 0.5 N NaOH, 10 mM EDTA, and 15 μl of denaturing loading buffer (95% formamide, 20 mM EDTA, 0.05% bromophenol blue and 0.15% xylene cyanol). After heating at 95 °C for 5 min, samples were loaded in wells pre-cooled to 4 °C and run using 8% nondenaturing acrylamide gels containing 10% glycerol at 4–8 °C and 18–22 °C.
5-Aza-dC treatment and bisulfite DNA sequencing analysis
To assess re-activation of
Cav-1 expression, cells were treated with 5 μM of 5-Aza-dC (Sigma, St. Louis, MO) for 4 days. For bisulfite sequencing analysis, 1 μg of genomic DNA was incubated with 3 M sodium bisulfite (pH 5.0) and DNA samples were purified as described previously [
20]. Fifty ng of bisulfite-modified DNA were subjected to PCR amplification of the 37 CpG sites within the promoter and exon 1 using primers P1/P2 (Table
1). The PCR products were cloned into pCR
II vectors (Invitrogen Corporation, Alemeda, CA) and 5 clones of each specimen were subjected to DNA sequencing analysis to determine the methylation status.
Immunoblotting assay
Cells were lysed in a lysis buffer containing 60 mM octylglucoside, 20 mM Tris (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 2.5 mM sodium phosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 μg/ml leupeptin and 1 mM PMSF. Twenty μg of total protein were supplemented with Laemmli buffer and loaded on a 10% SDS-polyacrylamide gel for electrophoresis. Western analyses were performed using antibodies specific for Cav-1 (sc-894, Santa Cruz, CA), Cav-2 (610,685, BD bioscience, CA), EGFR (#4267, Cell Signaling, Danvers, MA), RAF (#2330, Cell Signaling), MEK1/2 (#9911, Cell Signaling), ERK (#9101, Cell Signaling), AKT (#4060, Cell Signaling), JNK (#4668, Cell Signaling), and β-tubulin (T8328, Sigma). Antibody binding was detected by enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ) using a secondary antibody conjugated to horseradish peroxidase.
Immunofluorescence and immunohistochemistry (IHC) assay
For immunofluorescence assay, cells were fixed with 4% formaldehyde, permeabilized with 0.2% Triton X-100 and blocked with 2% bovine serum albumin-PBS. Slides were incubated with anti-GFP antibody and fluorescent imaging was obtained with a confocal laser scanning microscope (Carl Zeiss, Jena, Germany). IHC study was carried out using tissue arrays (SuperBioChips Laboratory, Seoul, Korea) and Vectastain ABC (avidin-biotin-peroxidase) kit (Vector Laboratories) as described previously [
21]. Briefly, slides were incubated with Cav-1 antibody overnight using biotin-free polymeric horseradish peroxidase-linker antibody conjugate system. Slides were counterstained with hematoxylin, dehydrated and visualized using an Olympus CK40 microscopy (Tokyo, Japan). For the immunoreactive score, we established a 1- to 12-point system by multiplying the percentage of positive cells by the intensity of the staining score. Two pathologists performed the assessment of immunostaining sections. Immunoreactive scores of 0–5 were classified as negative, and scores of 6–12 were regarded as positive [
22].
Ras activity assay
Cells were lysed with Mg-containing lysis buffer containing 25 mM HEPES (pH 7.5), 150 mM NaCl, 1% Igeal CA-630, 10% Glycerol, 25 mM NaF, 10 mM MgCl2, 1 mM EDTA, 1 mM Sodium orthovanadate, 10 μg/ml Leupeptin, 10 μg/ml Aprotinin, and 1 mM PMSF. Cell lysates were mixed with RAF-1 RBD agarose (Millipore, Billerica, MA) and the reaction mixture were rocked gently at 4 °C for 30 min. Agarose beads were collected by centrifugation, washed 3 times with lysis buffer, and resuspended in 2X Laemmli sample buffer. Samples were electrophoresed on SDS-PAGE and immunoblotted.
Expression plasmids, siRNA, shRNA, and transfection
GFP or Flag-tagged Cav-1 gene was cloned into the pcDNA3.1-V5-His (Invitrogen Corporation) and the pEGFP-N3 vector (Clontech, Mennheim Germany) using the Expand High Fidelity PCR system (Roche Molecular Biochemicals, Palo Alto, CA). siRNA against Cav-1 (5′-AACCAGAAGGGACACACAGUU-3′) and ERK2 (5′-CACCAUUCAAGUUCGACAUUU-3′) were synthesized by Dharmacon Research (Lafayette, CO). shRNA plasmid for Cav-1 (5′-caccACCTTCACTGTGACGAAATACTGGTTtctcAACCAGTATTTCGTCACAGTGAAGG-3′) was constructed by Genolution (Seoul, Korea). Transfection was performed using FuGENE 6 (Roche Molecular Biochemicals) or Oligofectamine (Invitrogen Corporation).
Cell proliferation, DNA synthesis, and colony formation assay
To measure in vitro cellular growth, cells were seeded at the density of 4 × 104 cells per well in triplicate and cell numbers were counted using a hemocytometer at 24 h intervals. For flow cytometry analysis, cells were fixed with 70% ethanol and resuspended in PBS containing 50 mg/ml RNase and 50 mg/ml propidium iodide (Sigma). The assay was performed on a FACScan flow cytometer (Becton Dickinson, San Jose, CA) and analyzed using Modfit software (Becton Dickinson). For DNA synthesis assay, cells were pulse-labeled for 4 h with 1 μCi/ml [3H] thymidine and harvested with lysis buffer (0.1 N NaOH, 1% SDS). The cell lysates were mixed with the liquid scintillation cocktail (ICN Inc., Irvine, CA) and [3H]thymidine incorporation was counted with Scintillation Counter (Wallac, Milton Keynes, UK). For colony formation assay, 1 × 105 cells per dish were maintained in the presence of G418 (1600 μg/ml) for 4–6 weeks. Selection medium were replaced every 2 days. Colonies were fixed with methanol for 15 min and stained with 0.05% crystal violet in 20% ethanol.
Statistical analysis
The results of cell growth, apoptosis and colony forming assays were expressed as mean ± SD. A student’s t-test was used to determine the statistical significance of the difference. The Chi-square test was used to determine the statistical significance of expression and methylation levels between tumor and normal tissues. A P value of less than 0.05 was considered significant.
Discussion
Cav-1 has opposite functions in tumorigenesis depending on the cellular contexts [
26]. However, the molecular mechanism underlying the differential effects of Cav-1 on tumor growth has been poorly defined. In the present study, we observed a bidirectional alteration of Cav-1 expression in gastric cancers, which is linked to the mitogenic conversion of its function. Our study provides evidence that the mitogenic conversion of Cav-1 function is associated with the switch of its role for the RAF-ERK feedback phosphorylation loop.
Previous IHC studies reported contrasting results on Cav-1 expression in gastric cancers [
27‐
30]. A study using frozen tissues and a monoclonal antibody revealed that Cav-1 is expressed in only a small fraction of intestinal type cancers [
28]. Meanwhile, a study using formalin-fixed specimens and a polyclonal antibody showed that Cav-1 is expressed in both diffuse and intestinal types at variable levels [
30]. In the present study, we identified that both intestinal and diffuse types of cancers express Cav-1 at highly variable levels. However, Cav-1 down-regulation was significantly more frequent in early versus advanced tumors while its up-regulation was more common in advanced versus early tumors and high versus low grade tumors, supporting that alteration of Cav-1 expression is associated with the oncogenic switch of its function [
29,
31]. Our data thus suggest that Cav-1 may act as a stage-specific growth modulator in gastric cancer, which is inactivated during the early stages of tumorigenesis and its subsequent elevation confers growth advantages and malignant progression [
26,
32].
It is becoming clear that altered expression of Cav-1 in tumor stroma, particularly in cancer-associated fibroblasts (CAFs), is linked to the malignant progression of various types of human cancers [
33,
34]. A study showed that loss of CAFs Cav-1 promotes tumor microenvironment remodeling and tumor development [
35]. However it was also reported that stromal Cav-1 favors tumor invasion and metastasis [
36]. Therefore, the role for CAFs Cav-1 in tumorigenesis remains largely undefined. It was demonstrated that Cav-1 is not expressed in the epithelial compartment in normal gastric mucosa and in the metaplastic intestinal epithelium while its expression is significantly higher in advanced versus early cancers and an independent prognostic factor of poor survival [
29]. In contrast, a recent study using quantum dots immunofluorescence histochemistry identified that epithelial Cav-1 expression gradually decreases with the progression of gastric cancer [
37]. Interestingly, this study also showed that low Cav-1 expression in CAFs rather than in tumor cells predicts recurrence and survival in cancer patients, suggesting that loss of stromal Cav-1 heralds poor prognosis of gastric cancer patients, which is consistent with the finding in breast and prostate cancer [
38,
39]. Although we did not characterize CAFs Cav-1 expression status in the current study, it was recognized that compared to normal gastric mucosa, noncancerous tissues adjacent to cancerous tissues exhibit much variable levels of Cav-1 mRNA. Further studies will be required to address whether Cav-1 gene expression in CAFs also shows a bidirectional alteration due to promoter hypo- and hyper-methylation during gastric tumor progression.
Mutational alteration of the
Cav-1 gene has been rarely found in human cancers. However,
Cav-1 mutations were reported in certain tumor types [
8]. A mutant Cav-1 (P132L) found in scirrhous breast cancer was identified to exert a dominant negative function by cytoplasmic retention [
8]. Interestingly, we detected three missence and one silent sequence alterations in
Cav-1 from 3 of 50 primary tumors and 1 of 14 cancer cell lines. Our preliminary data suggest that all of these mutants are expressed. The central α-helical region of Cav-1 protein interacts with the catalytic subunit of protein kinase A (PKAcat) through a hydrogen bond between its Y97 residue and W196 residue of PKAcat [
38]. Our finding of Y97N mutation in the SNU638 cancer cell line thus raises the possibility that Cav-1 regulation of PKA signaling might be altered in these gastric cancer cells.
Cav-1 null mice exhibit increases in tumor incidence, tumor area, and tumor number compared with wild-type counterparts [
39]. In contrast, prostate cancer cells secrete Cav-1, which stimulates clonal growth of tumor cells, and high Cav-1 expression exerts anti-apoptotic effect under clinically relevant circumstances [
40,
41]. Our previous study showed that high Cav-1 expression enhances the metastatic potential of gastric tumor cells by increasing the adhesion ability of the cells to endothelium through the regulation of cell surface VCAM [
42]. In the present study, we observed that Cav-1 provokes either a growth-inhibiting or growth-promoting effect in gastric cancers, and this property of Cav-1 is associated with its reciprocal regulation of ERK. Signaling components, including Ras, RAF, MEK, and ERK, are known to be compartmentalized within caveolin-rich membrane domains [
2]. However, there is a disagreement in the data concerning whether Cav-1 plays an inhibitory or stimulatory role in Ras-ERK signaling in cancer cells. We found that Cav-1 reciprocally regulates MEK and ERK in low- and high-expressing tumor cells. RAF is a crucial factor to transmit growth factor-induced Ras signals to MEK, and its activity is precisely regulated by differential phosphorylation of RAF isoforms and a negative feedback loop between RAF and ERK [
25]. We found that Cav-1 evokes opposite effects on the inhibitory phosphorylation of RAF in low- and high-expressing cells through the reciprocal control for the ERK-mediated inhibitory phosphorylation of RAF. We validated that in EGF-treated gastric tumor cells, RAF is inhibited by ERK feedback phosphorylation and this feedback loop is differentially regulated by Cav-1 in these two cell types. The opposite effects of Cav-1 on tumor cell growth were disrupted if ERK expression is depleted, supporting that the mitogenic conversion of Cav-1 effect is linked to its reciprocal regulation of the ERK feedback phosphorylation of RAF. Although further studies are required to understand the molecular mechanism underlying the Cav-1 regulation of the RAF-ERK negative feedback loop, studies suggest that Cav-1 may affect ERK feedback regulation of signaling components, including RAF, MEK, and KSR1 [
15,
43,
44]. It is thus conceivable that MEK-ERK signaling is activated through Cav-1 up- and down-regulation in early and advanced tumors, respectively.