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Department of Biology, University ‘Roma Tre’, Viale G Marconi, 446,I-00146 Roma, Italy

(Requests for offprints should be addressed to M Marino; Email: m.marino{at}uniroma3.it)

	Abstract

Top Abstract Introduction Materials and methods Results Discussion References

 
The cellular functions regulated by 17ß-estradiol (E2) start after the hormone binds to its receptors (i.e., ER and ERß ). These act as ligand-dependent transcription factor transactivating target genes. In addition, E2 induces non-genomic actions, whose activation is triggered by a fraction of the ERs localized at the plasma membrane. Palmitoylation allows ER to localize at the plasma membrane, to associate with caveolin-1, and, upon E2 stimulation, to activate rapid signals relevant for cell proliferation. The existence of a mechanism, which allows ERß localization at the plasma membrane and its putative role in anti-proliferative E2 effects is completely unknown. Here, the susceptibility of ERß to undergo palmitoylation and the role played by this process has been analyzed in DLD-1 containing endogenous ERß or in HeLa cells transiently transfected with ERß or ER expression vectors. As for ER , palmitoylation is necessary for ERß localization at the plasma membrane and its association with caveolin-1 but, in contrast to ER , the E2 binding increases ERß association with caveolin-1 and the p38 member of MAPK family. Moreover, the palmitoyl acyl transferase (PAT) inhibitor blocks the ability of ERß –E2 complex to activate p38 impairing the receptor-dependent activation of downstream proapoptotic cascade (i.e., caspase-3 activation and poly(ADP-ribose)polymerase (PARP) cleavage). Consequently, palmitoylation must be considered to be a molecular device for ERß , which allows these receptors to interact with the plasma membrane and to regulate E2-induced non-genomic functions relevant to the anti-proliferative effect of this hormone.

	Introduction

Top Abstract Introduction Materials and methods Results Discussion References

 
Knowledge of the pleiotropic functions regulated by 17ß-estradiol (E2) in different tissues and organs has evolved rapidly during the past decade. It is now well established that the E2 effects in living cells are mediated by a complex signaling network rather than a single uniform mechanism. In fact, in addition to its role as gene transcription regulator (Acevedo & Kraus 2004), sustained by the two estrogen receptors (ER and ERß ), E2 induces rapid, membrane starting, non-genomic actions (Levin 2005, Marino et al. 2005, Song et al. 2005, Leclercq et al. 2006).

Among other physiological roles, there is growing evidence that the rapid actions activated when E2 binds to the isoform of ER are important for cell proliferation. The E2 treatment of mammary-derived MCF-7 cells triggers ER association with Src, and the regulatory subunit of phosphatidyl inositol 3-kinase, (PI3K) leading to DNA synthesis (Castoria et al. 2001). Moreover, in HepG2 cells, multiple and parallel membrane starting pathways are rapidly activated by the ER –E2 complex (Marino et al. 1998, 2002, 2003). The blocking of phospholipase C/protein kinase C, mitogen-activated protein kinase (MAPK)/extracellular regulated kinase (ERK), and PI3K/AKT pathways completely prevents the E2-induced DNA synthesis (Marino et al. 2002, 2003). All these results point to the belief that membrane localized ER is the primary endogenous mediator of rapid E2 actions important for cell proliferation and survival.

However, recent reports have demonstrated that E2 could even decrease cell growth by significantly increasing apoptosis in several cell types (see Song & Santen 2003 for review). In particular, epidemiologic, clinical, and experimental evidence show that E2 confers protection against prostate and colon cell proliferation (Horvath et al. 2001, Konstantinopoulos et al. 2003, Bardin et al. 2004, Koehler et al. 2005).

ERß seems to participate in these E2-induced blockages of cell proliferation. A progressive decline of ERß expression has been reported in multistage mammary carcinogenesis (Roger et al. 2001) and prostate cancer (Horvath et al. 2001); moreover, ERß is the predominant ER expressed in colonic tissues and its expression is selectively lost in human malignant colon tissue (Konstantinopoulos et al. 2003, Wada-Hiraike et al. 2006). It has been proposed that ERß could act as a dominant regulator of E2 signaling (Koehler et al. 2005), and when co-expressed with ER , it would cause a concentration-dependent reduction in ER-mediated transcriptional activation (Pettersson et al. 1997). The ERß-directed repression of ER-mediated effects includes cell proliferation (Imamov et al. 2005, Koehler et al. 2005). Data from gene expression in cell cultures and knockout mice, clearly indicate that E2-activated ERß must have some function as a tumor suppressor by modulating the proliferative effects of ER (Couse & Korach 1999, Cheng et al. 2004, Paruthiyil et al. 2004, Strom et al. 2004).

These studies support a functional antagonism between ER and ERß with respect to the E2-induced cell proliferation, but failed either to ascertain the putative role of ERß in E2-induced apoptosis or to identify the signal transduction pathways involved. However, the ability of the ERß –E2 complex to activate rapid non-genomic mechanisms has been reported (Castoria et al. 2001, Kousteni et al. 2001, Geraldes et al. 2003, Mori-Abe et al. 2003). We recently demonstrated that E2-induced rapid signal transduction pathways in ERß-transfected HeLa cells appear to play a major role in mediating anti-proliferative properties of this steroid hormone. The action of E2 in these cells results from binding to ERß which, in turn, acutely promotes the rapid and persistent phosphorylation of the p38 member of MAPK family, thus triggering downstream activation of a proapoptotic cascade (Acconcia et al. 2005a). The rapidity by which these cellular cascades are activated raises the need for a receptor localized at the plasma membrane. Although a subpopulation of ERß localized within caveolar rafts, responsible for rapid endothelial nitric oxide synthase stimulation by E2 has been reported in the plasma membrane of endothelial cells (Chambliss et al. 2002), the mechanism allowing ERß localization at the plasma membrane and its putative involvement in the anti-proliferative effect mediated by ERß –E2 complex is completely unknown. We previously demonstrated that the Cys447 residue present in the ER ligand binding domain (LBD) is palmitoylable and this lipid modification is necessary for the induction of the non-genomic ERK/MAPK signal transduction pathway, which is relevant to E2-induced cell proliferation (Acconcia et al. 2005b). Although, the homology between ER and ERß LBDs is only 59% (Ascenzi et al. 2006), the amino acid sequence encompassing the palmitoylated Cys447 and Cys132 residues of ER and caveolin-1, respectively, is highly homologous to that surrounding the Cys399 residue of ERß (Acconcia et al. 2003). Based on these findings, we postulated that ERß serves as a palmitoyl acyl transferase (PAT) substrate and that the receptor palmitoylation is important for the localization at the membrane and for the activation of the E2-dependent proapoptotic cascade in colon cancer cells. Our results indicate that ERß palmitoylation is a major determinant for ERß –membrane and ERß –protein interaction (i.e., caveolin-1 and p38/MAPK) important for the E2-induced protective effect against colon cancer.

	Materials and methods

Top Abstract Introduction Materials and methods Results Discussion References

 
Reagents

E2, gentamicin, penicillin, GenElute plasmid maxiprep kit, Dulbecco Modified Eagle Medium (DMEM), RPMI-1640 medium (without phenol red), charcoal-stripped fetal calf serum, and the PAT inhibitor 2-bromohexadecanoic acid (2-bromopalmitate; 2-Br) (IC₅₀ 4.0 µ M; Varner et al. 2003), were purchased from Sigma-Aldrich (St Louis, MO, USA). The p38/MAPK inhibitor, SB 203 580 (SB), was obtained from Calbiochem (San Diego, CA, USA). The ER inhibitor ICI 182 780 (ICI) was obtained from Tocris (Ballwin, MO, USA). 9,10-[³H]-palmitic acid (specific activity 57.00 Ci/mmol) was purchased from Dupont-NEN (Boston, MA, USA). Lipofectamine reagent was obtained from Gibco-BRL Life-technology. The luciferase kit was obtained from Promega. Bradford Protein Assay was obtained from Bio-Rad Laboratories. The monoclonal anti-phospho-ERK, anti-AKT, anti-Src, anti-caspase-3, anti-poly(ADP-ribose)polymerase (PARP), and anti-ß-actin as well as the polyclonal anti-ERK, anti-caveolin-1, anti-ER MC20 (C-terminus), and anti-ERß L20 (C-terminus) antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). The monoclonal anti-ERß 14C8 (N-terminus) antibody was purchased from Genetex (San Antonio, TX, USA). The policlonal anti-phospho-AKT, anti-phospho-p38, and anti-p38 antibodies were obtained by New England Biolabs (Beverly, MA, USA). The modulator of non-genomic activity of estrogen receptor (MNAR, also named Pro-, Glu-, and Leu-rich protein-1; PELP1) polyclonal antibody was purchased from Benthyl Laboratories (Montgomery, TX, USA). CDP-Star, chemiluminescence reagent for Western blot was obtained from NEN. All the other products were from Sigma-Aldrich. Analytical or reagent grade products, without further purification, were used.

Cell culture and count

The ERs-devoid human cervix epithelioid carcinoma cell line (HeLa; Marino et al. 2002) and the ERß containing human colon adenocarcinoma cells (DLD-1; Marino et al. 2006) were used as experimental models. Cells were routinely grown in air containing 5% CO₂ in modified, phenol red-free, DMEM (HeLa cells) or RPMI-1640 (DLD-1 cells) media, containing 10% (v/v) charcoal-stripped fetal calf serum, L-glutamine (2 mM), gentamicin (0.1 mg/ml), and penicillin (100 U/ml). Cells were passaged every 2 days and media changed every 2 days.

DLD-1 cells were grown to approximate 70% confluence in six-wells plates then stimulated. Thirty hours after treatment (10 nM E2, 1 µ M ICI 182 780, 10 nM E2–BSA), cells were harvested with trypsin and centrifuged. Cells were stained with the Trypan blue solution and counted in a hemocytometer (improved Neubauer chamber) in quadruplicate.

Plasmids

The reporter plasmid containing the promoter of complement component 3 gene, retaining a natural estrogen responsive element (ERE), linked to the gene of luciferase (pC3), the reporter plasmid containing the promoter of cyclin D1 pXP2-D₁– 2966-luciferase (pD1), the expression vector pCR3.1-ß-galactosidase, wild type human ER pSG5-HE0, human ERß (pCNX2-ERß ) have been described elsewhere (Herbert et al. 1994, Marino et al. 2002, Acconcia et al. 2004, 2005a). A luciferase dose-response curve showed that the maximum effect was obtained when 1.0 µ g of plasmids was transfected together with 1.0 µ g of pCR3.1-ß-galactosidase to normalize for transfection efficiency (approximate 55–65%). Plasmids were purified for transfection using the GenElute plasmid maxiprep kit according to the manufacturer’s instructions.

Transfection and luciferase assay

HeLa and DLD-1 cells were grown to ~70% confluence and then transfected using lipofectamine reagent according to the manufacturer’s instructions. Six hours after transfection, the medium was changed and 24 h after the cells were stimulated with 10.0 nM E2 for 6 h. The cell lysis procedure as well as the subsequent measurement of luciferase gene expression was performed using the luciferase kit according to the manufacturer’s instructions with an EC & G Berthold luminometer (Bad Wildbad, Germany). When indicated, the PAT inhibitor 2-Br (final concentration 10.0 µ M) was added 15 min before E2 administration.

Cell labelling with [³H]-palmitate and immunoprecipitation

Twenty-four hours after transfection with either plasmid containing ER or ERß , HeLa cells and untransfected DLD-1 cells were incubated with 0.5 mCi/ml [³H]-palmitate at 37 ° C for different times ranging between 0 and 240 min. DLD-1 and HeLa cells were stimulated with different concentrations of E2 (0.1, 1.0, 10.0, and 100.0 nM) for different times (10, 60, and 240 min) in the presence of [³H]-palmitate. Cells were then washed in ice-cold PBS, harvested with trypsin (1%, v/v), and lysed in 50 µ l lysis buffer (10.0 mM Tris, pH 7.5, 1.0 mM EDTA, 0.5 mM EGTA, 10.0 mM NaCl, 1% (v/v) Triton X-100, and 1% (w/v) sodium cholate) containing protease inhibitors (0.5 mM phenylmethylsulfonyl fluoride, 1.0 µ g/ml leupeptin, and 5.0 µ g/ml aprotinin). The cell lysates were then clarified by centrifugation and immunoprecipitated as described previously (Acconcia et al. 2005b). Briefly, equal amounts of soluble cell extracts were incubated with different anti-ERß antibodies. The optimal signal was obtained using 1.0 µ g anti-ERß L10 (C-terminus) together with 1.0 µ g of anti-ERß 14C8 (N-terminus). The lysates and antibodies were incubated for 90 min at 4 ° C, and then 20 µ l of protein-A agarose were added for 30 min at 4 ° C. After centrifugation (50 000 g for 15 min), the supernatant and the immunoprecipitated proteins were separated in 7–10% SDS-PAGE. In some experiments, the radioactivity present in immunoprecipitated proteins and in the supernatant was monitored by counting with a Camberra Packard Liquid ß-counter (Milan, Italy).

True-blot immunoprecipitation

The cell lysates, prepared as described above, were clarified by centrifugation and immunoprecipitated with TrueBlot (eBioscience, San Diego, CA, USA), which preferentially detects the native disulfide form of mouse IgG or rabbit IgG reducing interference by approximate 55 kDa heavy and approximate 23 kDa light chains of the immunoprecipitating antibody. Briefly, after stimulation, equal amounts of soluble cell extracts were incubated with either 2.0 µ g of anti-caveolin-1 or anti-p38 or anti-Src or anti-MNAR or anti-14C8 ERß antibody. The lysates and antibodies were incubated at 4 ° C for 1 h, then 20 µ l of Anti Mouse IgG Beads (eBioscience) were added and samples incubated for 1 h on a rocking platform at 4 ° C. Samples were centrifuged at 50 000 g for 10 min, the supernatant was removed completely and beads (pelleted) were washed three times with 100 µ l of lysis buffer. SDS-reducing sample buffer (20 µ l, containing 50 mM DTT) was added and samples were boiled at 100 ° C for 5 min. Proteins were resolved using 10% SDS-PAGE at 100 V for 1 h and then electrophoretically transferred to nitrocellulose for 45 min at 100 V at 4 ° C. The nitrocellulose was treated with 5% (w/v) non-fat dry milk (Bio-Rad Laboratories) in 150 mM NaCl, 50.0 mM Tris HCl (pH 8.0), 0.1% (w/v) Tween-20, and then probed at 4 ° C overnight with anti-ERß antibody. The antibody reaction was visualized with the chemiluminescence reagent for Western blot.

Immunofluorescence studies

DLD-1 cells were grown on coverslips, treated as above specified, and fixed in 2% (w/v) freshly de-polymerized p-formaldehyde (Sigma-Aldrich) and 0.2% (v/v) glutaraldehyde (Agar Scientific Ltd, Stanstead, UK) in PBS. Cells were then incubated with 14C8 anti-ERß mouse monoclonal antibody diluted 1:100 in PBS containing 1% (w/v) BSA and 0.1% (v/v) Triton X-100. After washing, cells were incubated with goat anti-mouse fluorescein isothiocynate (FITC)-conjugated antibody (Vector Laboratories, Burlingame, CA, USA; diluted 1:200 in PBS containing 1% (w/v) BSA and 0.1% (v/v) Triton X-100). Finally, slides were mounted with Vectashield (Vector Laboratories) and localization of the receptor was examined with a Zeiss LSM510 confocal microscope. Pictures were electrically captured and composed by Photoshop 7.0 software.

RNA isolation and quantitative RT-PCR analysis (qRT-PCR)

The sequences for gene-specific forward and reverse primers were designed using the OligoPerfect Designer software program (Invitrogen). The following primers were used: for human ER (ESR1/NR3A1, GeneBank Accession No. AY425004 [GenBank] ), 5'-TCCTAGCAGGGAGATGAGGA-3' (forward) and 5'-CCTTTATGGCCAGCAATCAT-3' (reverse), for human ERß (ESR2/NR3A2, GeneBank Accession No. AY785359 [GenBank] ), 5'-GGCGCGATCTTG-GCTCAC-3' (forward) and 5'-TGGCTGGACGT-GGTGGCA-3' (reverse), and for ß-actin (GeneBank Accession No. X00351 [GenBank] ), 5'-AGAAGGATTCC-TATGTGGGCG-3' (forward) and 5'-CATGTCGTC-CCAGTTGGTGAC-3' (reverse).

Total RNA was extracted from DLD-1 cells using TRIzol Reagent (Invitrogen) according to the manufacturer’s instructions. To determine hER-ß gene expression levels (as well to confirm the absence of hER- ), cDNA synthesis and qPCR were performed using a one-step qRT-PCR kit (‘SuperScript III Platinum – SYBR Green One-Step’ kit; Invitrogen) according to the manufacturer’s instructions. cDNA synthesis and qPCR was carried out in a ABI Prism 7900HT Sequence Detection System (Applied Biosystems, Foster City, CA, USA) as follows: first strand cDNA synthesis was performed at 50 ° C for 5 min, followed by an automatic hot-start Taq DNA Polimerase activation step at 95 ° C for 5 min, and then by 25 cycles of denaturation at 95 ° C for 15 s, annealing at 60 ° C for 30 s, and elongation at 40 ° C for 1 min. Gene expression was verified by electrophoresis on 2% agarose gel. Each sample was tested in duplicate and the experiment repeated four times.

Electrophoresis and immunoblotting

After treatment, cells were lysed and solubilized in 0.125 M Tris, pH 6.8, containing 10% (w/v) SDS, 1.0 mM phenylmethylsulfonyl fluoride, and 5.0 µ g/ml leupeptin; then the cell lysates were boiled for 2 min. In some experiments cells were homogenized using ten strokes of the pestle of Potter homogenizer until about 90% of the cells were broken. Homogenates were centrifuged at 1000 g for 10 min to pellet the nuclear fraction. Soluble and particulate fractions were obtained by centrifuging the supernatants at 100 000 g for 30 min. Proteins were solubilized as above described. Total proteins were quantified using the Bradford Protein Assay. Solubilized proteins (20 µ g) were resolved by 10% SDS-PAGE at 100 V for 1 h at 24 ° C and then electrophoretically transferred to nitrocellulose for 45 min at 100 V and 4 ° C. The nitrocellulose was treated with 3% (w/v) BSA in 138.0 mM NaCl, 25.0 mM Tris, pH 8.0, at 24 ° C for 1 h and then probed overnight at 4 ° C with either anti-phospho-ERK or anti-phospho-AKT or anti-phospho-p38 or anti-ERß antibodies. The nitrocellulose was stripped by Restore Western Blot Stripping Buffer (Pierce Chemical Company, Rockford, IL, USA) for 10 min at room temperature and then probed with either anti-ERK or anti-AKT or anti-p38 or anti-caspase-3 or anti-PARP and anti-ß-actin antibodies. Antibody reaction was visualized with chemiluminescence Western blotting detection reagent (Amersham Biosciences). The PAT inhibitor 2-Br (10.0 µ M) or the p38 inhibitor SB 203 580 (5 µ M) was added 30 min before E2 administration.

Statistical analysis

A statistical analysis was performed by utilising Student’s t-test with the INSTAT software system for Windows. Some data were analyzed by one-way ANOVA and post hoc Bonferroni’s test (INSTAT software system for Windows). In all cases probability (P) values below 0.05 were considered significant.

	Results

Top Abstract Introduction Materials and methods Results Discussion References

 
E2 reduces DLD-1 cell growth

The main difficulty in studying ERs action mechanism in cancer cells derives from the expression of a great number of ERs splice variants (Herynk & Fuqua 2004). To bypass this problem we chose human colon adenocarcinoma (DLD-1) cells as an experimental model, which contain only one ERß isoform corresponding to 54 kDa protein (Fig. 1A and B). As already reported (Marino et al. 2006), E2 stimulation decreases DLD-1 cell growth. This effect requires ERß , since it is completely prevented by ICI 182 780 (Fig. 1, panel C). Stimulation of DLD-1 cells with the E2 cell membrane impermeable E2-BSA, a well-known agent able to discriminate between non-genomic vs genomic effects of ER(s) (Marino et al. 2003), affected DLD-1 cell growth, as did E2 (Fig. 1, panel C). This indicates the pivotal role of plasma membrane-starting signals in E2-induced anti-proliferative effects.

View larger version (26K): [in this window] [in a new window]   Figure 1 Level of ERs in DLD-1 cells. (A) Western blot analysis of ER and ERß levels were performed in DLD-1 and HeLa cells. HeLa cells were untransfected (none) or transfected with ER or ERß expression vectors. Fifty four kilodaltons of human ERß recombinant protein (5 ng, Invitrogen) was used as control. (B) Identification of ERß mRNA by RT-PCR using specific sense/antisense primers. Amplification products were separated by electrophoresis on 2% agarose and identified by ethidium bromide staining. ERß =108 bp; ER =248 bp. (C) DLD-1 cells grown for 30 h in the presence of E2 (10 nM) and/or ICI 182 780 (ICI, 1 µ M) or E2–BSA (10 nM) and then counted. The data are the mean values ± S.D. of five independent experiments carried out in duplicate. P < 0.001, calculated with Student’s t-test, compared with respective unstimulated values (control) (*) or with E2-stimulated values (). For details see text.

We first verified the occurrence of ERß palmitoylation in DLD-1 cells. Cells were incubated with [³H]-palmitate at 37 ° C for different periods (from 0 to 240 min) and the amount of radioactivity in both the immunoprecipitate and the supernatant was determined (Fig. 2A). ERß and radioactivity were not detected in the supernatant fractions (Fig. 2A and data not shown). [³H]-palmitate incorporation in immunoprecipitated ERß was complete within 120 min and remained unchanged over 240 min (Fig. 2B). As a positive control, the radioactivity present in the palmitoylated caveolin-1 (Resh 1999), immunoprecipitated from DLD-1 cells, was measured (Cav-1) (Fig. 2C). A significant decrease in ERß and caveolin-1 palmitoylation occurred in DLD-1 cells pre-treated with the PAT inhibitor, 2-bromohexadecanoic acid (2-Br) (Fig. 2C). These findings indicate that ERß , like caveolin-1, undergo PAT-dependent palmitoylation.

View larger version (14K): [in this window] [in a new window]   Figure 2 ERß palmitoylation. (A) Western blot analysis of immunoprecipitated ERß in the pellet (P) or the supernatant (S). Fifty four kilodaltons of human ERß recombinant protein (5 ng, Invitrogen) was used as control. (B) Time course of [3H]-palmitate incorporation in immunoprecipitated ERß in DLD-1 cells. Data are the means of six independent experiments ± S.D. (C) [3H]-palmitate incorporation (120 min) in immuno-precipitated ERß or caveolin-1 (Cav-1) in the presence or absence of PAT inhibitor 2-Bromopalmitate (2-Br, 10 µ M). (D) Time course of [3H]-palmitate incorporation in HeLa cells transfected with ER or ERß expression vectors. ERs were immunoprecipitated and radioactivity counted. Data are the means of four independent experiments ± S.D. of duplicate analyses. For details see text.

ERß palmitoylation is negatively modulated by E2

To assess the ability of E2 to modulate ERß palmitoylation, DLD-1 cells were incubated with [³H]-palmitate for 4 h in the presence of different E2 concentrations. Physiological E2 concentration (1–10 nM) decreased the amount of [³H]-palmitate incorporated in ERß by more than half; a higher E2 concentration (i.e., 100 nM) was more efficient to decrease palmitate incorporation in ERß , whereas hormone lower concentration was ineffective (i.e., 0.1 nM; Fig. 3A). The time course of 10 nM E2 stimulation in DLD-1 cells showed that 60 min of E2 stimulation reduced the [³H]-palmitate incorporation in ERß by 38% (Fig. 3B) with an increase in the protein level (Fig. 3C). The increase in ERß levels were also detected 120 and 240 min after E2 stimulation (Fig. 3C), which is in good accordance with data reported in the literature (Chiang et al. 2000, Matthews & Gustafsson 2003 and literature therein). The kinetics of E2-induced de-palmitoylation is very similar for both ERs, as demonstrated in transfected HeLa cells (Fig. 3D).

View larger version (18K): [in this window] [in a new window]   Figure 3 E2 effect on ERß palmitoylation. (A) Dose-response curve of E2. [3H]-palmitate was incorporated in DLD-1 cells for 120 min, then different E2 concentrations were administrated for further 120 min and ERß was immunoprecipitated. Data are the means of five independent experiments ± S.D. (B) Time course of E2 stimulation. [3H]-palmitate was incorporated in DLD-1 cells for 120 min, then cells were stimulated with 10 nM E2 at different time and ERß was immunoprecipitated. Data are the means of five independent experiments ± S.D. (C) Typical Western blot of immunoprecipitated ERß from DLD-1 cells stimulated with 10 nM E2 at different time. (D) HeLa cells were transfected with human ER or human ERß expression vectors. [3H]-palmitate was incorporated in HeLa cells for 120 min, then cells were stimulated with 10 nM E2 at different time and ERß or ER were immunoprecipitated. Data are the means of four independent experiments ± S.D. For details see the text.

Besides, lipid modification, the association with membrane proteins could allow extrinsic proteins to localize at the plasma membrane. After E2 binding, ER is able to associate to adaptor and/or signaling proteins, which in turn are responsible for signaling cascade activation important for cell proliferation (Greger et al. 2006). This prompted us to evaluate the ability of ERß to interact with some of these signaling proteins, both in the absence and in the presence of 10 nM E2 for 15 min. No association between ERß and Src or MNAR was observed before or after E2 stimulation of DLD-1 cells (Fig. 4A and B, respectively). This result was followed by the inability of ERß to induce ERK or AKT phosphorylation (Fig. 4C and D, respectively). On the contrary, the association of ERß with caveolin-1 is present even in the absence of E2 (Fig. 5A and B). This association increase 60 min after E2 stimulation (Fig. 5A and B). Notably, E2-induced ERß-caveolin-1 association was completely prevented by pre-treatment with the PAT inhibitor, 2-Br (Fig. 5C). No variation in the level of caveolin-1 was present after E2 stimulation (Fig. 5B).

View larger version (18K): [in this window] [in a new window]   Figure 4 ERß association to Src and MNAR and activation of ERK/MAPK and PI3K/AKT pathways in DLD-1 cells. Cells were grown in the absence (0) or stimulated for 15 min with 10 nM E2, ERß was immunoprecipitated with anti-ERß antibody followed by Western blot with anti-Src (A) or with anti-MNAR (B) and anti-ERß antibodies. Western blot analysis of ERK (C) and AKT (D) phosphorylation in DLD-1 cells were performed on unstimulated (0) and stimulated cells for 10–30 min with E2 (10 nM). Data represent a typical Western blot of three different experiments. For details see text.

View larger version (36K): [in this window] [in a new window]   Figure 5 Role of ERß palmitoylation in ERß association to caveolin-1. DLD-1 cells were stimulated with 10 nM E2 for different times then were subjected to ERß immunoprecipitation (A and A') or caveolin-1 immunoprecipitation (B and B') followed by Western blot with anti-caveolin-1 or with anti-ERß antibodies. (C and C') DLD-1 cells were pre-treated for 30 min with 10 µ M PAT inhibitor 2-Br then stimulated with 10 nM E2 for different times and subjected to ERß immunoprecipitation or caveolin-1 immunoprecipitation followed by Western blot with anti-caveolin-1 or with anti-ERß antibodies. (A, B, and C) Typical Western blot; (A', B', and C') densitometric analysis of four different experiments. Data are the mean ± S.D. For details see text.

View larger version (62K): [in this window] [in a new window]   Figure 6 (A) Immunofluorescence analysis of unstimulated (left panel), 10 nM E2 stimulated for 120 min (central panel), and pre-treated for 30 min with 10 µ M PAT inhibitor 2-Br then stimulated with 10 nM E2 for 120 min (right panel) DLD-1 cells. White arrows indicate the membrane localization of ERß . (B) Cell fractionation assay of unstimulated (1, control), 10 nM E2 stimulated for 120 min (2, + E2), and pre-treated for 30 min with 10 µ M PAT inhibitor 2-Br then stimulated with 10 nM E2 for 120 min (3, + E2 + Br) DLD-1 cells. Typical Western blot (upper panel) and densitometric analysis (bottom panel) of four different experiments. Data are the mean ± S.D. For details see text.

In DLD-1 cells, the rapid, persistent E2-induced activation of the p38/MAPK pathway is sufficient and necessary for E2-induced apoptosis (i.e., caspase-3 activation and PARP cleavage; Marino et al. 2006). We next determined whether ERß palmitoylation could have an impact on these rapid non-genomic ERß activities. E2 induced the rapid and persistent phosphorylation of p38 in DLD-1 cells (Fig. 7A). We investigated whether a physical association existed between p38 and ERß by immunoprecipitation. Under basal conditions a complex formed by the unphosphorylated form of p38 and ERß has been detected (Fig. 7B). After 10 min of E2 treatment, ERß –p38 association as well as p38 phosphorylation significantly increased (Fig. 7B). However, when the cells were pre-treated with the PAT inhibitor, 2-Br, the E2-induced activation of this signaling kinase was completely blocked even though the basal p38 levels were unaffected (Fig. 7C).

View larger version (35K): [in this window] [in a new window]   Figure 7 Role of ERß palmitoylation on p38/MAPK activation. (A and A') Time course analysis of p38/MAPK phosphorylation was performed on untreated (0) and E2-treated (10 nM) DLD-1 cells at the indicated times. The amount of protein levels were normalized by comparison with actin expression. (B and B') DLD-1 cells were stimulated with 10 nM E2 for different times then were subjected to ERß or p38 immunoprecipitation followed by Western blot with anti-p38 or anti-p38 phosphorylated (p38-P) or anti-ERß antibodies. (C and C') DLD-1 cells were pre-treated for 30 min with 10 µ M PAT inhibitor 2-Br then stimulated with 10 nM E2 for different times and subjected to ERß or p38 immunoprecipitation followed by Western blot with anti-p38 or anti-p38 phosphorylated (p38-P) or anti-ERß antibodies. (A, B, and C) Typical Western blot; (A', B', and C') densitometric analysis of four different experiments. Data are the mean ± S.D. For details see text.

View larger version (35K): [in this window] [in a new window]   Figure 8 Role of ERß palmitoylation on p38-dependent pro-apoptotic cascade activation. (A and A') Western blot analysis of caspase-3 and PARP (panel b) activation were performed on unstimulated or 24 h E2-treated (10 nM) DLD1 cells. When indicated 5 µ M p38/MAPK cascade inhibitor SB 203 580 (SB) or 10 µ M PAT inhibitor 2-Br was added 30 min before E2 administration. The amount of protein levels were normalized by comparison with actin expression. (A and B) Typical Western blot; (A' and B') densitometric analysis of five different experiments. Data are the mean ± S.D. For details see text.

We previously demonstrated that ER palmitoylation is required for E2-induced gene transcription (Acconcia et al. 2005b). Since, we wanted to compare the ER and ERß transcriptional activity in the same cellular context, the ERs devoid HeLa cells were used as experimental model. HeLa cells were co-transfected with the ERE containing pC3 promoter and ERß or ER expression vectors. As expected, in HeLa cells transfected with ER , the E2 treatment induced a three- and a two-fold increase of pC3 and pD1 promoter activities, respectively (Fig. 9). On the other hand, in ERß-transfected HeLa cells E2 induced a twofold increase only on the pC3 promoter activity (Fig. 9). Notably, the pre-treatment of ER or ERß-transfected HeLa cells with the PAT inhibitor 2-Br reduced the E2-inducible pC3 promoter activity by 30% without affecting the basal pC3 promoter activity (Fig. 9). As expected, the ER palmitoylation has a powerful effect on the E2-induced cyclin D1 promoter activity, which is totally impaired by 2-Br treatment, whereas ERß was unable to mediate cyclin D1 promoter activity, both in the presence or absence of 2-Br.

View larger version (31K): [in this window] [in a new window]   Figure 9 Role of ERs palmitoylation on the E2-induced genomic activity in HeLa cells. Cells were co-transfected with human ERß or human ER expression vectors together with pC3-luciferase (pC3) or pD1-luciferase (pD1) constructs and pre-treated 30 min with 10 µ M PAT inhibitor 2-Br before E2 administration (10 nM for 6 h). The data are the mean values ± S.D. of five different experiments. P < 0.001 was calculated with Bonferroni’s test: a, significantly different from control value (open bar); b, significantly different from E2 stimulated samples; and c, significantly different from ER transfected HeLa cells.

	Discussion

Top Abstract Introduction Materials and methods Results Discussion References

Then, which are the molecular mechanisms underlying the above opposite effects? Like most other members of the nuclear receptor family, ERs have a modular architecture of four interacting domains: the N-terminal A/B domain, the C or DNA binding domain (DBD), the D or hinge domain and the C-terminal E/F or LBD (Marino et al. 2005, Ascenzi et al. 2006). The homology in the DBD is 97% (Ascenzi et al. 2006), which suggests that ER and ERß are capable of regulating gene transcription through a classical mechanism involving the consensus ERE element, but ERß seems to be a weaker transactivator (Cowley & Parker 1999).

To date, only a limited number of genes have been shown to be regulated by this classical mode of action. E2 also transcriptionally regulates target genes via ERs though a non-ERE mode of action. These effects are mediated through promoter elements that bind various transcription factors (e.g., AP-1-binding sites and Sp1-binding sites; Webb et al. 1998, Porter et al. 1997). At these sites, ER and ERß could have opposite transcriptional effects in some circumstances. In fact, ERß is not able to potentiate an AP-1-containing reporter in the presence of E2 in a tissue-specific manner (Paech et al. 1997). In addition, E2 interaction with ERß does not result in the formation of a transcriptionally active complex at a promoter containing Sp1 elements (Saville et al. 2000). More significant differences are reported between ERß and ER actions with respect to their ability to activate rapid E2-induced signals. The activation of ERK/MAPK, PI3K/AKT, and PKC, rapidly generated after E2 binding to ER in different cell lines, are all defined as necessary and sufficient for E2-induced G1 to S phase progression, to increase survival pathways (e.g., Bcl-2), and to regulate the transcription of AP-1- and Sp1-dependent genes important for cell cycle modulation (e.g., cyclin D1). Interestingly, the transcriptional activity of the E2–ER complex could be inhibited by pre-treating cells with the ERK inhibitors PD98059 and U0126 (Levin 2005, Marino et al. 2005).

Limited, conflicting data are reported for ERß-mediated rapid signals. A subpopulation of ERß transfected in CHO cells is capable of activating IP₃ production, ERK/MAPK and c-Jun kinase phosphorylation (Razandi et al. 1999). Geraldes and coworkers reported that E2 reduces ERK/MAPK activity through ERß stimulation in porcine smooth muscle cells (Geraldes et al. 2003). Moreover, contradictory evidence on the ability of ERß to activate or inactivate Src and p38 kinases have also been reported (Castoria et al. 2001, Kousteni et al. 2001, Geraldes et al. 2003, Mori-Abe et al. 2003). We recently reported the ERß –E2 complex ability to activate p38/MAPK in the ER-devoid HeLa cells transiently transfected with the ERß expression vector (Acconcia et al. 2005b). The discrepancies reported in these studies could be due to the different cellular models utilized, in which ERß is over-expressed, or both ER and ERß are co-expressed, or different ERß splicing forms could even be present. This further enhances the complexity in the spectrum of potential cellular responses to E2.

Here, using colon cancer cells which contain only one ERß isoform, we demonstrated that ERß is localized to the plasma membrane and originate rapid signal transduction cascades important for anti-proliferative effects of E2. In fact, we prove that ERß undergoes PAT-dependent palmitoylation even if this isoform is a poor substrate for PAT as compared to ER (Fig. 2). In addition, the localization to the membrane is dependent on ERß palmitoylation since the PAT inhibitor impairs ERß from localizing at the membrane and from interacting with caveolin-1. Similar to that reported for ER is the time- and concentration-dependent negative regulation of ERß palmitoylation exerted by E2 (Fig. 3).

Palmitoylation function must be considered more than a simple membrane association of otherwise soluble proteins. In fact, the palmitoylation status of several proteins has also been linked to their activation and their movement within membrane subdomains. As an example, de-palmitoylation of eNOS increases in response to treatment of cells with bradykinin (Robinson et al. 1995) and de-palmitoylation has a subtle effect on membrane distribution of G-proteins affecting their partitioning within membrane subdomains (Smotrys & Linder 2004). Thus, palmitate addition is a dynamic modification that is continually turning over on cellular proteins. ERß and ER do not contain a trans-membrane domain (Zhang et al. 2004) or consensus sequences for miristoylation or prenylation (Acconcia et al. 2003), thus their ability to associate with scaffolding or/and signaling proteins at the plasma membrane seems principally due to palmitoylation (Acconcia et al. 2005b, Levin 2005). In the resting state, ERß is localized mainly in the cytosol and nucleus of DLD-1 cells and only a little amount of receptor is tethered with caveolin-1 (Fig. 5). After E2 stimulation, ERß undergoes de-palmitoylation, which increases receptor–caveolin-1 association (Fig. 5) and, thus, its presence at the plasma membrane (Fig. 6). E2 stimulation of ER-containing cells decreases receptor palmitoylation with a kinetics similar to that reported for ERß (Fig. 3). The E2-dependent de-palmitoylation decreases ER –caveolin-1 association allowing ER association with adaptors and/or signaling proteins (e.g., MNAR, Src, tyrosine kinase receptors), which in turn give rise to rapid signaling cascades (e.g., MAPK and PI3K; Levin 2005, Marino et al. 2005, Song et al. 2005, Leclercq et al. 2006). This does not occur in the presence of ERß . It has been reported that intact A/B domain and tyrosine 537 in E domain of ER are both required for receptor interaction with Src in the MNAR–ER –Src complex and the in vitro association between ERß and MNAR has been reported (Barletta et al. 2004, Greger et al. 2006). Although ERß possess a tyrosine residue at 488, which could be subjected to phosphorylation, the ER and ERß A/B domain differ in both length and amino acid sequence, exhibiting a low amino acid identity (Ascenzi et al. 2006). Moreover, present data indicate that no association between ERß and MNAR and Src was present in DLD-1 cells before and after E2 stimulation (Fig. 4). On the other hand, E2 increased ERß level (Chiang et al. 2000, Matthews & Gustafsson 2003 and present data) and its association with caveolin-1 (Fig. 5). As a whole, these data raise the intriguing possibility that the short A/B domain of ERß could facilitate the E2-induced association between ERß and caveolin-1, impairing its association with MNAR and Src. As a consequence, ERK and AKT activation does not occur. Experiments in our laboratory are in progress to better define this pathway.

ERK/MAPK as well as PI3K/AKT cascades cooperate in ER–E2 induced cell proliferation and cell survival, enhancing the expression of the anti-apoptotic protein (Bcl-2) and promoting the G1/S transition via the enhancement of cyclin D1 expression (Marino et al. 2002, 2003). In addition, the E2–ER complex rapidly increased p38/MAPK phosphorylation but the contemporary increase of Bcl-2 levels, mediated by ERK/MAPK and PI3K/AKT pathways, impairs the prolonged p38 activation and the downstream effects of this kinase (Acconcia et al. 2005b). On the contrary, the rapid increase of p38 phosphorylation induced by the E2–ERß complex is not modulated by ERK/MAPK and PI3K/AKT pathways, thus a more prolonged p38 phosphorylation occurs (Acconcia et al. 2005b; Fig. 6).

The ERß palmitoylation is important for E2-induced cell functions. In fact, ERß–p38 association and E2-induced prolonged activation of this kinase is prevented by the PAT inhibitor 2-Br (Fig. 6). Moreover, ERß palmitoylation is necessary for the p38-dependent activation of downstream pro-apoptotic cascade, which involves the caspase-3 activation and PARP cleavage (Fig. 7). To evaluate the impact of palmitoylation on E2-transcriptional effects, we compared ERß and ER in a cellular context, which contained the same co-activators. As expected, ERß is a weaker transactivator than ER. The palmitoylation of ERß scarcely influences ERß genomic activities (i.e., ERE-containing gene promoter transcription). Similar results were obtained in ER-containing cells, indicating that the rapid palmitoylation-dependent signal transduction pathways are important for the complete transcriptional activity of ERs. This could be due to the ability of such cytosolic signals to modulate the co-activator recruitment and the chromatin activation status (Smith & O’Malley 2004). ERß and ER have opposite effects in mediating the E2-induced transcription of cyclin D1 (i.e., non-ERE-containing gene promoter). As expected, ERß was unable to induce cyclin D1 promoter transcription, whereas a critical requirement of ER palmitoylation for cyclin D1 promoter transcription was present.

In conclusion, present findings indicate that palmitoylation localizes ERß at the plasma membrane, and this even if only in little quantity, directs several E2 effects, which allow the protective effect of this hormone in colon cancer. These data, showing the molecular mechanism, which rapidly follows E2 entry in ERß-containing cells, further sustain the tumor suppressor function played by this receptor isoform. Although ER is palmitoylated, the outcome effects in cell physiology are opposite to that reported for ERß. Thus, the expression of each ER isoform and/or their co-expression in the cells could account for the different E2-dependent modulation of cell proliferation reported.

	Acknowledgements

 
The authors wish to thank Prof. Paolo Ascenzi (Department of Biology, University ‘Roma Tre’, V.le G Marconi, 446, I-00146 Rome, Italy) for helpful and critical discussions. The generous gift of DLD-1 cells from Dr Aldo Cavallini (Biochemistry Laboratory, I R C C S ‘de Bellis’, V della Resistenza, Castellana Grotte, I-70013 Bari, Italy) and of human ERß expression vector from Prof. Masami Muramatsu (Research Center for Genomic Medicine, Saitama Medical School, 38 Morohongo, Moroyama, Iruma-gun, Saitama 350-0495, Japan) are gratefully acknowledged. The authors express gratitude to Dr Fulvio Florenzano (IRCCS, ‘Fondazione S Lucia’, V Ardeatina, Rome, Italy) for helpful advice in performing confocal microscopic analysis. The editorial assistance of Mr Peter De Muro is also acknowledged. This work was supported by grants from MIUR (COFIN-PRIN 2004) to M M. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.

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