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J. Biol. Chem., Vol. 278, Issue 40, 38875-38883, October 3, 2003
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¶ ¶ ¶ ¶ 
From the
Departments of Pediatrics, Ophthalmology and Pharmacology, Research Center of Hôpital Sainte-Justine, Montréal, Québec H3T 1C5, Canada, the ||Department of Anatomy and Cell Biology, Université de Sherbrooke, Sherbrooke, Québec J1H 5N4, Canada, the ¶Department of Pharmacology and Therapeutics, McGill University, Montréal, Québec H3G 1Y6, Canada, and the **Department of Medicine and Microbiology-Immunology, University of California, San Francisco, California 94143-0711
Received for publication, December 9, 2002 , and in revised form, June 18, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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B (see Ref. 12 for review). Alternatively, LPA may exert intracellular actions by generating its own formation, inferring possible active intracellular binding sites. Along this line of thought, LPA signaling at discrete subcellular domains may be provided in part by intracellular conveyors such as gelsolin and the fatty acid-binding protein (8), by uptake of extracellular LPA bound to albumin (via albondin receptor; gp60) (13), and/or to lipocalins (14).
Lastly, several established pathways of GPCR regulation and desensitization driven by extracellular agonists have also been implicated in their intracellular relocalization, which results in delayed complementary signaling cascades (15, 16). In this context, accumulating evidence suggests that nuclear translocation of peptide growth factors (e.g. angiogenin, PDGF, basic FGF, EGF, and PTH) and/or their integral membrane receptors, is mandatory for gene transcription associated with proliferating/growth events (17). Because most of these latter receptors (e.g. EGF, PDGF, and PTH receptors) co-localize within caveolae (18), we speculated that translocation to this specific organelle proceeds via receptor endocytosis through the caveolar compartment. Whether this phenomenon holds true for the transmission of LPA gene responses (1, 2) is not yet known.
The biochemical mechanisms by which GPCRs, including LPA-Rs, modulate gene transcription are complex and not fully understood (19). Some mechanisms recently uncovered to explain GPCR-mediated gene induction implicate an endocytosis-associated 
-arrestin-c-Src interaction leading to downstream activation of Ras and mitogen-activated protein kinases (MAPKs) through possibly metalloprotease-dependent transactivation of receptor tyrosine kinases (RTK) involving de novo release of their ligand (19). Herein, we postulated that endogenous LPA stimulates gene expression, specifically the pro-inflammatory genes cyclooxygenase-2 (COX-2) and inducible nitric-oxide synthase (iNOS), through a formerly undescribed mechanism, which involves the activation of nuclear LPA receptors that pre-exist at the nuclear envelope or originate from the internalization/endocytosis of plasmalemmal LPA receptors. In the present study, we focused on endothelial cells, which are known to express predominantly LPA1R (20) and consolidated our findings on LPA1R stably transfected HTC4 rat hepatoma cells (21) and rat liver tissue specimens. Our in vitro and in vivo findings support the existence of constitutive LPA1R at the cell nucleus, which upon stimulation mediates calcium transients and transcriptional signals of immediate-early response genes. Our results also suggest that (i) LPA-induced PLA2-dependent COX-2 expression is not reliant on prostaglandin, leukotriene, or epoxide production and (ii) contrary to common peptide growth hormone receptors, the sequestration and transcellular transport of LPA-R via caveolae to the nucleus is not a prerequisite for LPA-R activity on gene expression. This study unravels an as yet undescribed mechanism by which LPA modulates gene expression.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-cyclodextrin, ibuprofen (Sigma); EGTA, pertussis toxin (PTX), fura-2-AM, wortmannin, ionomycin, MK886, cytidine-5-diphosphocholine, mepacrine, tyrphostin AG 1478, PD 98059, and tunicamycin (Calbiochem); SK&F96365, CV 3988, and methylcarbamyl platelet-activating factor (C-PAF) (BIOMOL); ketoconazole (ICN Biochemicals Inc.); fluo-4-AM (Molecular Probes); PNGase F assay kit (New England BioLabs); brain microvessel endothelial growth media (BioWhittaker); Dulbecco's modified Eagle's medium (Invitrogen); [3H]oLPA (PerkinElmer Life Sciences); RNA guard RNase inhibitor (Amersham Biosciences); and recombinant human interleukin-1 (IL-1) (BIOSOURCE International). All other chemicals were analytical reagents and were purchased from Fisher Scientific (Montréal, Québec, Canada).
Antibodies and their sources are: Anti-murine iNOS monoclonal antibody, anti-CD51 monoclonal antibody, and anti-human caveolin-1 monoclonal antibody (Transduction Laboratory); anti-human COX-2 polyclonal antibody (Cayman); anti-human -actin polyclonal antibody; anti-human Von Willebrand factor polyclonal antibody (Dako, Denmark); anti-phospho-MAPK (Erk1/2) polyclonal antibody (Promega); anti-MAPK (Erk1/2) polyclonal antibody (Upstate Biotechnology); anti-phospho-Akt (Ser473) and anti-Akt polyclonal antibodies (New England BioLabs); anti-alkaline phosphatase polyclonal antibody (Abcam); anti-lamin A/C monoclonal antibody (Chemicon International); anti-cytochrome c monoclonal antibody (BD Pharmingen); horseradish peroxidase-conjugated goat anti-rabbit and anti-mouse IgG (Pierce); rabbit polyclonal anti-human LPA1R-C antibody (raised against a C-terminal epitope; consisting of amino acids 328-344) and its cognate peptide antigen (Upstate Biotechnology); and rabbit polyclonal and mouse monoclonal anti-human LPA1R-N receptor antibodies (N-terminal epitope consisting of amino acids 6-25; Dr. E. J. Goetzl, University of California, CA). The specificity of anti-LPA1R antibodies has been fully established elsewhere (22-25).
Animals—Experiments were performed on endothelial cells derived from Yorkshire piglet brain microvasculature (Fermes Ménard, Quebec, Canada) and hepatocytes from adult Sprague-Dawley male rats (Charles River, Quebec, Canada). Animal housing and experimental protocols were carried out in accordance with regulations set by the Canadian Council of Animal Care Committee and were approved by the Sainte-Justine Hospital Animal Care Committee.
Cell Culture and Fractionation—Primary endothelial cells obtained from porcine cerebral microvessels (26) and stably transfected, Geneticin-resistant LPA1-HTC4 rat hepatoma cells (21) were cultured and passaged, as previously reported. Cell lines were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 1% penicillin G, 1% streptomycin and used between passages 6-13 at 
80% confluency unless otherwise stated. Prior to stimulation, cells were serum-starved overnight. Isolation of nuclei was achieved by cell fractionation using the hypotonic/Nonidet P-40 lysis method (27). Nuclear envelopes were prepared by incubating nuclear suspensions (8 x 106 nuclei/ml) with DNase I (800 units, pancreas type II, Roche Applied Science) and RNase A (32 mg/ml, Promega) for 30 min at 37 °C (28). Supernatants of homogenized cells (in lysis buffer) were sequentially centrifuged at 10,000 x g for 15 min then 120,000 x g for 60 min to obtain mitochondrial and plasma membrane fractions, respectively. Plasma and nuclear membranes and intact nuclei were stored at -80 °C unless otherwise stated. Protein concentration was determined by Bradford protein assay using BSA as standard. The morphological integrity and purity (>98%) was routinely assessed by light microscopy after trypan blue staining and confirmed by electron microscopy (27) (Fig. 1D, inset). The plasma membrane marker 5'-nucleotidase activity (Sigma assay kit) in nuclear versus plasma membrane fraction was less than 7%.
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Isolation of Subcellular Fractions of Rat Liver—Hepatocytes were harvested following digestion of liver with collagenase (type II, 0.05%, Sigma), as described (29). Isolation of liver nuclei was carried out by ultracentrifugation through a sucrose gradient according to Nicotera et al. (30). Plasma membrane microsomes and nuclear envelopes were prepared, and morphological integrity and purity were ascertained as indicated above; 5'-nucleotidase activity endowed in the isolated nuclear fraction was less than 2% of that of plasma membrane fraction. Purity of subcellular fractions was further substantiated by means of immunological methods using the specific organelle marker antigens alkaline phosphatase and CD51 (plasma membrane), lamin A/C (nuclei), and cytochrome c (mitochondria, cytosol), as depicted below in Fig. 1A. Fig. 1B exhibits electron micrographs of highly purified nuclei (>98%) and prepared nuclear envelopes (right panel), appearing as intact spheres, following this isolation procedure.
Radioligand Binding Assays—Cells were seeded into 24-well plates (500 µl of media/well) and allowed to reach 90% confluency (70,000 cells/well) before beginning experiments. Quiescent cells were washed thrice with PBS containing BSAfaf (0.1%), soybean trypsin inhibitor (100 µg/ml), and phenylmethylsulfonyl fluoride (1 mM) then incubated in the same buffer at 4 °C for 90 min with 10 nM radioactive tracer [3H]oLPA (50 Ci/mmol). Thereafter, cells were rinsed twice with cold incubation buffer (1 ml), lysed with sodium hydroxide (0.1 N), and transferred into scintillation vials. Nonspecific binding was determined in the presence of unlabeled LPA (10 µM). Radioactivity of samples was measured with a -counter. In parallel, cells from untreated wells within the same plate were harvested with trypsin and counted thrice with a hemacytometer for cell count normalization.
Subcellular Fractions—Binding assays were conducted on either whole nuclei or nuclear envelopes originating from porcine EC and rat liver using 10 nM radioactive tracer [3H]oLPA. Thawed rat liver nuclei (5 x 106 nuclei corresponding to 250 µg of protein) and prepared envelopes (5 x 106 nuclear envelope corresponding to 100 µg of protein) were resuspended in buffer consisting of Trizma-HCl (50 mM), pH 7.4, KCl (25 mM), MgCl2 (5 mM), sucrose (0.25 mM), CuSO4 (0.5 mM), BSAfaf (0.1%); pCMVEC-isolated nuclei (0.15 x 106 nuclei/100 µg of protein) in Trizma-HCl (20 mM), pH 7.4, KCl (10 mM), MgCl2 (3 mM), phenylmethylsulfonyl fluoride (0.5 mM), BSAfaf (0.1%). The reaction was terminated by diluting samples (twice with a 40x volume excess) followed by rapid filtration on GF/C filters presoaked in incubation buffer supplemented with BSAfaf (1%). In each experiment, nonspecific binding was determined in the presence of unlabeled LPA or surrogates (10 µM). Under these experimental conditions, the level of binding was directly proportional to time exposure (equilibrium time 60 min) or the amount of proteins added (50-400 µg) (not shown).
Western Blot Analysis of LPA1R—Equal amounts (25 µg) of plasma membrane (PM), whole nuclei (WN), and derived nuclear envelope (NE) protein from pCMVEC, HTC4, and hepatocyte cells were solubilized in Laemmli buffer, separated by 12% reducing SDS-PAGE, and electro-blotted onto PVDF membranes. Membranes were then blocked with 5% nonfat dry milk in Tris-buffered saline-Tween 20 (20 mM Tris-HCl, 150 mM NaCl, and 0.1% Tween 20) (TBS-T) and subsequently incubated overnight at 4 °C either with a monoclonal anti-LPA1R-N (1:100), a polyclonal anti-LPA1R-N (1:2000), or a polyclonal anti-LPA1R-C (1: 2000). Thereafter, membranes were washed with TBS-T and incubated for 1 h at RT with secondary antibodies (monoclonal, 1:2500; polyclonal, 1:5000) conjugated to horseradish peroxidase. Finally, membranes were thoroughly washed and developed using an enhanced plus ECL system (PerkinElmer Life Sciences). In some experiments, the effects of the N-glycosylation inhibitor, tunicamycin, and the endoglycosidase PNG-ase F were evaluated on cultured pCMVECs and an enriched plasmalemmal fraction of rat liver, respectively. These experiments were based on the presence of putative glycosylation consensus sequences (NX(S/T)) residing in rat (27NES29; 35NES37) and human (27NES29; 35NRS37) LPA1R orthologues (31, 32). For this purpose, pCMVEC were treated or not with tunicamycin (0.05, 0.5, or 1.0 µg/ml) for 24 h at 37 °C. Cells were then washed twice with cold PBS, lysed in 50 mM Tris-HCl, pH 7.4, 1% Nonidet P-40, 0.25% sodium deoxycholate, 150 mM NaCl, 2 mM Na3VO4, 1 mM NaF, complete protease mixture inhibitor, and solubilized for 1 h at 4 °C using a rotary shaker. Cellular extracts were centrifuged (21,000 x g for 10 min at 4 °C), and the resulting supernatant was collected and proteins were measured. For deglycosylation experiments, 50 µg of rat liver PM was denatured and incubated or not with peptide N-glycosidase F (PNGase F) (500,000 units/ml; 5 µl) at 37 °C for 5 h according to the manufacturer's instructions. Samples (10-50 µg of proteins) were resolved by SDS-PAGE (9-12%), transferred onto PVDF membranes, and blotted with a polyclonal anti-LPA1R-N antibody (1:4000).
Electron and Cryomicroscopic Immunohistochemistry of LPA1R— Male Sprague-Dawley rats (250-300 g) were used. Rat liver tissue sectioning and pre-embedding immunogold staining was done as described in detail previously (33). Vibratome sections (50 µm) of liver were incubated with the primary antibody (a rabbit anti-LPA1R-C antibody) (1:50) overnight at 4 °C followed by another overnight incubation with goat anti-rabbit gold (10 nm)-conjugated IgG (1:50) (British Biocell International). In control sections, the primary antibody was either removed or pre-absorbed with its cognate peptide (10-fold excess by weight, Upstate Biotechnology). Thereafter, specimens were postfixed in 1% osmium tetroxide, subsequently dehydrated in graded ethanol, and embedded in Epon according to standard technique. Ultrathin sections were cut using a Reichert Ultracut ultramicrotome, mounted on Formvar-coated copper grids, stained with uranyl acetate and lead nitrate, and examined with a transmission electron microscope (Philips 410LS, Netherlands).
Electron Cryomicroscopy—All procedures were based on previously reported methods (34). Rat liver sections were cut at -100 °C in an FC4D cryochamber using a Reichert-Jung ultramicrotome and transferred to Formvar-coated copper grids. For immunolabeling assays, sections were incubated for 30 min at RT with rabbit polyclonal anti-human LPA1R-N or -C antibody (1:10) followed by incubation with a goat anti-rabbit gold (10 nm)-conjugated IgG (1:20) (Sigma Chemical Co.). Negative controls were analyzed by omitting primary antibodies. Frozen sections were contrasted and embedded as described (35) and viewed through of a Philips 400T electron microscope.
Indirect Immunofluorescence Staining—Isolated nuclei from pCMVECs were resuspended in blocking buffer constituted of 10 mM Tris-HCl, 20 mM NaCl, 3 mM MgCl2, 300 mM sucrose, complete protease mixture inhibitor, 5% horse serum, and 5% fetal calf serum. Double staining was achieved by incubating the nuclear suspension (1 h at RT) with a polyclonal rabbit anti-LPA1R-N (1:50) and a monoclonal mouse anti-caveolin-1 antibody (1:25). The nuclear suspension was diluted in a 5-fold excess of blocking buffer then centrifuged (800 x g, 10 min) and resuspended in similar buffer. The nuclei were subsequently incubated 60 min with a goat anti-rabbit AlexaFluor 488-conjugated IgG (2 µg/ml) and a chicken anti-mouse AlexaFluor 647-conjugated IgG (10 µg/ml) (Molecular Probes) and washed as indicated above. The nuclear suspension was then placed on poly-L-ornithine-treated glass coverslips (25 mm) and examined with a laser-scanning confocal microscope. Samples were imaged using a 60x differential interference contrast oil immersion objective lens on a Nikon TE300 microscope with a Bio-Rad Radiance 2000 confocal accessory. The two images were collected by using the same Z values and were merged using a Silicone graphic (SGI) software. Nuclear staining was realized at the end of the experiment using the DNA dye Syto-11 (100 nM) (Molecular Probes). For negative controls the primary antibodies were omitted.
Immunoprecipitation of LPA1R and Western Blotting—Cellular and nuclear extracts (500 µg) of pCMVEC were lysed, immunoprecipitated by an anti-LPA1R-N polyclonal antibody (4 µg/ml), separated by SDS-PAGE (12%) and transferred onto nitrocellulose membrane. Immunoblotting was performed with either an anti-LPA1R-N polyclonal antibody (serving as positive control) (1:2000), anti-clathrin (1:1000), or caveolin-1 (1:1000) monoclonal antibody. Cellular and nuclear extracts of EC were subjected to extraction in Triton X-100 followed by sucrose density gradient centrifugation to isolate caveolae (35). Caveolae-enriched fractions (low density and Triton X-100-insoluble materials) were verified by SDS-PAGE and probed for caveolin-1. Equal amounts of proteins from caveolar and non-caveolar membranes were then immunoblotted for LPA1R as indicated above.
Western Blot of COX-2 and iNOS Proteins—pCMVEC were seeded into 10-cm dishes, rendered quiescent with starving medium, and treated for 6 h with either the vehicle (bidistilled and deionized water), sLPA (1-10 µM) or human IL-1 (10 ng/ml). Cells were then washed twice with cold PBS (5 ml) and harvested in the same buffer by gentle scraping. Sedimented cells were then lysed by brief sonification in buffer consisting of Tris-HCl (10 mM), pH 7.4, EDTA (5 mM), and complete protease mixture inhibitor and then centrifuged at 200 x g for 10 min at 4 °C. The supernatant was collected and submitted to an ultracentrifugation (100,000 x g for 60 min at 4 °C). Resulting crude membrane and cytosolic extracts were concentrated (Centricon Plus 20, PL-10, Millipore) and analyzed for protein content. Denatured membrane proteins (30 µg) were resolved by SDS-PAGE on 7.5% gel and transferred onto PVDF membranes, which were subsequently cut and probed with either an anti-human Von Willebrand factor polyclonal antibody (1:200), an anti-COX-2 polyclonal antibody (1:1000), or an anti--actin polyclonal antibody (1:500). Denatured cytosolic proteins (30 µg) were processed in a similar way and immunoprobed using a mouse anti-iNOS monoclonal antibody (1:200). Relative quantities of proteins were expressed in terms of a densitometric ratio to -actin. Densitometric analysis was carried out by integrating the intensity of all pixels in a representative area, excluding background, using Image-Pro+ software (Version 4.1; Media Cybernetic, Silver Spring, MD).
Gene Induction and Semi-quantification of iNOS and COX-2 mRNA— pCMVEC seeded into 10-cm dishes were treated for 2 h at 37 °C with either the vehicle (sterile bidistilled and deionized water), sLPA (1-10 µM), or human IL-1 (10 ng/ml) in the presence or absence of caveolae-disrupting agents filipin (5 µg/ml) and methyl--cyclodextrin (5 mM), non-selective phospholipase A2 inhibitors quinacrine (20 µM), and cytidine-5-diphosphocholine (2 µM), 5-lipoxygenase inhibitor MK 886 (20 µM), epoxygenase cytochrome P-450 inhibitor ketoconazole (30 µM), non-selective cyclooxygenase inhibitor ibuprofen (20 µM), PAF receptor antagonist CV 3988 (1 µM), MEK inhibitor PD 98059 (10 µM) or RTK inhibitor tyrphostin AG 1478 (250 nM) (27, 36-39). Inhibitors were incubated 1 h prior to LPA stimulation. Cells were then washed twice with cold PBS (5 ml), collected, and frozen in liquid nitrogen. Total cellular RNA was isolated using the RNeasy Protect Mini Kit (Qiagen).
iNOS gene expression was also studied in a cell-free nuclear system. For this purpose, isolated rat liver nuclei (106 nuclei/assay; 250 µg of protein) were placed in buffer of the following composition: Tris-HCl (10 mM), pH 7.5, KCl (10 mM), MgCl2 (3 mM), CaCl2 (100 nM), ATP, UTP, GTP, and CTP (500 µM), and Superase RNase inhibitor (10 units, Ambion, total reaction medium volume of 60 µl). Nuclei were incubated at 37 °C for 60 min with sterile water vehicle or oLPA (1 µM) in the presence or absence of Ca2+ chelator EGTA (100 µM), pre-activated PTX (20 µg/ml, 60-min preincubation), or phosphoinositide 3-kinase/Akt (PI 3-kinase) inhibitor wortmannin (100 nM) (27). Total RNA from purified nuclei was isolated by the standard guanidine isothiocyanate method (40). iNOS and COX-2 mRNA were quantified by reverse transcriptasepolymerase chain reaction (RT-PCR) method as described elsewhere (29, 41, 42), with slight modifications. Briefly, 1 µg of total RNA was transcribed with Moloney murine leukemia virus reverse transcriptase (Invitrogen) and the cDNAs (50 ng) were amplified following a 35-cycle protocol (94 °C for 30 s, 58.5 °C for 35 s, and 68 °C for 1 min), using Platinum Pfx DNA polymerase (Invitrogen) and specific primers for both rat and pig iNOS, COX-2, and -actin genes. The sequences of pig COX-2 primers (sense: 5'-ATGATGTACGCCACAATCTGG; antisense: 5'-GTTGAAAAGCAGCTCTGGGTC) and pig -actin primers (sense: 5'-TCCCTGGAGAAGAGCTACGA; antisense: 5'-ATCTGCTGGAAGGTGGACAG) are as shown. The primers used for rat iNOS (41), COX-2 and -actin (29), and pig iNOS (42) were as described. PCR products were quantified using an Agilent 2100 bioanalyzer in conjunction with DNA 7500 Labchip kits and by electrophoresis on precast 1% agarose gels stained with SYBR® Gold nucleic acid stain (Agilent Technologies). Gels were imaged and analyzed with a fluorescence gel imaging system. Alternatively, the PCR products were 32P-labeled, separated on a 6% polyacrylamide gel, and quantified with a PhosphorImager (Amersham Biosciences).
Measurement of Ca2+ Signals by LPA in Isolated Nuclei—Nuclear Ca2+ signals were measured by fura-2-AM technique as essentially described (32) with minor modifications. Briefly, isolated nuclei (1-5 x 106 nuclei/ml) placed in incubation buffer (Hepes (25 mM), pH 7.2, KCl (125 mM), K2HPO4 (2 mM), MgCl2 (4 mM), CaCl2 (400 nM)), with an osmolarity adjusted with glucose to 300 mosM, were loaded in the dark with fura-2-AM (7 µM) (45 min at 4 °C). Fura-2-loaded nuclei were then diluted (1:10), centrifuged at 600 x g for 10 min, and resuspended with the incubation buffer. Leakage of fura-2 from nuclei was minimal (<2%) during the time frame of experiments as reported (31). The [Ca2+]nuclear was determined in 1 ml of fura-2-loaded nuclei (5 x 106 nuclei/ml) at 37 °C under constant stirring using the ratio of excitation wavelengths 350/380 nm with emission set at 500 nm. Nuclear calcium signals were measured by means of a spectrofluorometer (model LS50, PerkinElmer Life Sciences, UK). Calibration of maximal (Rmax) fluorescence signals was determined by sequential addition of ionomycin (5 mM) and calcium (1 mM), whereas that of minimal (Rmin) fluorescence signals was obtained with Triton X-100 (1%) followed by EGTA (25 mM). The [Ca2+]n was calculated according to Grynkiewicz et al. (43) by the equation: [Ca2+]n = Kd (224 nM) [(R - Rmin)/(Rmax - R)] (Sf2/Sb2), where Kd is the dissociation constant (224 nM for fura-2) and Sf2/Sb2 is the ratio of fluorescent intensity at 380-nm wavelength. In another set of experiments, confocal microscopy was used to monitor movements of nuclear Ca2+ induced by LPA (1 pM to 1 µM) in a single nucleus. For this purpose, nuclear suspension was loaded with fluo-4-AM (30 µM) at 25 °C for 45 min then diluted in 20x excess volume, spun down, and resuspended in the above-mentioned buffer. Loaded nuclei (in 500 µl) were settled on glass coverslips and examined with a Molecular Dynamics (Sunnyvale, CA) Multi Probe 2001 confocal argon laser-scanning system as described previously (44). Rapid linescan imaging was used to monitor temporal oscillations of intranuclear calcium. Stimulated nuclei were recorded at a rate of 320 ms per scan for a total of 250 frames. Each frame consisted of 32 lines/scan (512 pixels) and a pixel size of 0.17 µm.
Western Blot of MAP Kinases and Akt Activation—Hepatocyte-derived nuclei (100 µg of protein) were treated or not with oLPA (1-10 µM) for 0-30 min in the above-mentioned buffer for calcium signal assay. In concomitant experiments, nuclear suspensions were pretreated with PI 3-kinase/Akt inhibitor wortmannin (100 nM) for 15 min at 37 °C and processed as above. Freezing samples in liquid nitrogen terminated the reaction. Proteins were resolved by SDS-PAGE on 9% gel, transferred onto PVDF membranes, and then probed with Erk1/2, phospho-Erk (1/2), Akt, and phospho-Akt antibodies (diluted 1:1000) according to manufacturer's instructions. Autoradiograms were scanned and analyzed by densitometry (ImagePro 4+ software).
Statistical Analysis—Data were analyzed by one-way analysis of variance factoring for treatments. Comparison among means was performed by Tukey-Kramer method. Statistical significance was set at p < 0.05. Data are presented as mean ± S.E.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Western Blot Analysis of LPA1R—Immunoprobing of subcellular fractions from native and recombinant cell systems with an anti-LPA1R-N polyclonal antibody revealed a prominent immunoreactive band at 49 kDa (Fig. 2A), in agreement with Zheng et al. (25). This is suggestive of a receptor with putative carbohydrate adducts, at the nucleus and plasma membrane of the cell types investigated. Similar results were obtained with the anti-LPA1R-C polyclonal antibody. The presence of glycan moieties comprised in LPA1R was established by assessing the electromobility shift of LPA1R protein to the predicted theoretical molecular mass (38 kDa) (31) following the treatment of rat liver cell membrane with PNGase F (Fig. 2B), or pCMVEC with tunicamycin (Fig. 2C); these oligosaccharide moieties account for 37% of the overall LPA1R molecular weight. In pCMVEC, two glycoforms (47 and 49 kDa) of LPA1R were detected by performing a low concentration SDS-PAGE (9%) (Fig. 2C). These results conformed to the quasi-universal post-translational modifications occurring in the GPCR family (45), including that of lysophospholipid receptor system (46).
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In Situ Detection of Perinuclear and Intranuclear LPA1R by Electron Microscopy—The intracellular localization of LPA1R was ascertained in vivo by high resolution immunogold electron microscopy and cryomicroscopy on rat liver tissue sections. LPA1Rs were identified at the plasma membrane of hepatocytes (Fig. 3A) and endothelial cells (Fig. 3B), as expected, and notably intracellularly, including the perinuclear and nuclear regions (Fig. 3B) mostly confined to euchromatin structures; these nuclear associated receptors account for 7-15% of overall cellular immunoreactivity. Negative controls revealed no specific labeling when the primary antibody was pre-absorbed with its cognate peptide (Fig. 3C) or was omitted (Fig. 3D). Ultrastructural localization of immunoreactive LPA1R in rat liver sections was further substantiated by electron cryomicroscopy using polyclonal rabbit anti-LPA1R-C (Fig. 3E) and anti-LPA1R-N antibodies (Fig. 3F), which target two distinct epitopes. In both cases, a broad and similar distribution pattern for LPA1R was revealed namely at the hepatic plasmalemma (not shown), the endoplasmic reticulum, the perinuclear envelope (arrowheads), as well as inside the nucleus (Fig. 3, E and F). Sporadic cytoplasmic detection of LPA1R was also observed at higher magnification, presumptively in vesicular compartments (not shown).
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Co-localization of LPA1R and Caveolin-1 in pCMVEC-isolated Nuclei—Many endocytic and transcytotic transport processes involve subcellular redistribution of transmembrane proteins, including possibly GPCRs, which are mainly mediated via either the clathrin or caveolae membrane system (16, 18). Therefore, we sought to investigate which of these endocytic systems may be monopolized for the nuclear translocation of cell surface LPA1R, if indeed it is occurring. Immunoprecipitation experiments demonstrated that LPA1R is dually retrieved in cell membrane clathrin-coated pits and caveolae while being only present in the caveolar compartment in the nucleus (Fig. 4A). Further support for the nuclear co-localization of LPA1R with caveolae in pCMVEC was obtained from Western blot experiments depicting the presence of LPA1R primarily in cholesterol-rich detergent-resistant caveolar domains of purified nuclei (Fig. 4B) and from immunofluorescence studies describing the double labeling of LPA1R and caveolin-1 in more than 90% of nuclear organelles examined. LPA1R co-localized with caveolin-1, as anticipated not exclusively.
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Role of Caveolae and Phospholipase A2-derived Products in LPA-induced COX-2 and iNOS Gene and Protein Expression— The induction of immediate-early response genes COX-2 and iNOS by LPA has recently been reported for various cell types. For instance, LPA-induced transcriptional up-regulation of COX-2 enzyme has been shown in renal mesangial cells (47), fibroblasts (48), and astrocytes (49), whereas iNOS has been induced in cardiomyocytes (50). We extended these observations by showing that sLPA (as well as oLPA) increases, in a concentration-dependent manner, COX-2 and iNOS protein expression in brain vascular endothelia (Fig. 5A), whereas the classic endothelial cell antigen von Willebrand factor was unaffected. The concentration-dependent effects of LPA on COX-2 gene expression were further substantiated by semi-quantitative RT-PCR (Fig. 5B). Based on the abovementioned results demonstrating co-localization of LPA1R with caveolin-1 more so in nuclear than plasma microsomal fractions (Fig. 4, A-C), we sought to explore whether caveolae was instrumental in LPA1R-mediated COX-2 expression by facilitating its transport to the cell nucleus. The caveolae-disrupting agents, filipin and methyl--cyclodextrin, both used at effective inhibitory concentrations (37, 38), failed to modify LPA activity. Similarly, inhibition of clathrin-dependent endocytosis by means of sucrose hypertonic solution (15) did not affect COX-2 expression (not shown). Conversely, LPA-induced COX-2 gene transcription was entirely blocked by the PLA2 inhibitors mepacrine and cytidine-5'-diphosphocholine suggesting a role for endogenous arachidonic acid and/or LPA in this phenomenon; partial impairment (50%) was also observed with RTK inhibitor tyrphostin AG 1478 and MEK inhibitor PD 98059 (Fig. 5B). The participation of cyclooxygenase-, lipoxygenase-, and epoxygenase-derived products of arachidonic acid and PAF was ruled out based on the inefficiency of corresponding specific enzyme inhibitors to block LPA responses (Fig. 5B). Thus, it appears that LPA signaling in inflammatory gene expression proceeds, independent of caveolae, via PLA2-triggered intracellular messengers, possibly LPA per se, acting conceivably on perinuclear/nuclear receptor sites (see Figs. 1, 2, 3). To corroborate this premise, we examined whether a direct stimulation of isolated nuclei with LPA would reproduce or mimic gene responses evoked by extracellular LPA. Results from these series of experiments are presented in the following section.
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Signaling Effectors of LPA-stimulated Calcium Transients and iNOS Expression in Highly Purified Rat Liver Nuclei—It is now becoming clear that the nucleus possesses autonomous calcium signaling machinery (e.g. ion channels, protein kinases, and phospholipases) where the nuclear envelope serves as the nuclear calcium pool (12). Furthermore, nuclear calcium is instrumental in regulating many processes, including gene transcription (12). We therefore examined whether changes in nuclear Ca2+ can be elicited by LPA1R stimulation by means of spectrofluorometry and live imaging confocal microscopy using different fluorescent Ca2+-sensitive probes (fura-2-AM and fluo-4-AM). Incubation of isolated hepatic nuclei with oLPA induced a concentration-dependent increase in nuclear Ca2+ levels (Fig. 6A); indistinguishable responses were seen with LPA analogues (1 µM) comprising distinct fatty acid species (sLPA and pLPA). Similar responses were observed in LPA1RHTC4 transformed hepatic cell-derived nuclei. Moreover, the Ca2+ chelator EGTA and putative receptor-operated Ca2+ channel blocker SK&F96365 prevented LPA-induced Ca2+ transients, as seen with PTX suggestive of Gi/o coupling, whereas the Ca2+ ATPase pump inhibitor thapsigargin was inoperative (Fig. 6, A and B). Rapid nuclear calcium mobilization in response to LPA was visualized in a single nucleus by confocal microscopy (Fig. 6B). Live scan imaging clearly illustrated a progressional transient Ca2+ movement across the nucleus (Fig. 6B, third panel). Notably, these discrete nucleoplasmic Ca2+ responses evoked by LPA were concentration-dependent (with an apparent affinity in the nanomolar range) (Fig. 6C), reversible and seemingly desensitizable (Fig. 6D) suggestive of a receptor-operated mechanism. Alternatively, this latter fading of organelle responsiveness upon repeated challenges of high concentration of LPA (1 µM) might reflect a re-localization and sequestration of calcium in unvisualized nuclear foci.
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Direct stimulation of nuclei with oLPA (as well as sLPA) also provoked dose-dependent, calcium- and PTX-sensitive iNOS gene induction (Fig. 7), consistent with results from calcium mobilization assays (see Fig. 6A). Along with the presence of nuclear Akt (Fig. 7A, inset), LPA effects were also associated with phosphorylation (and resultant activation) of this kinase (Fig. 7B), which in turn participated in iNOS gene induction. Accordingly, this latter effect was abolished by PI 3-kinase-activated protein kinase Akt inhibitor wortmannin (Fig. 7, A and B). In similar experimental conditions, LPA failed to induce phosphorylation of nuclear Erk1/2 kinases (not shown).
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Substantial evidence supports the presence of cell surface GPCRs, mostly for peptide ligands (e.g. angiotensin II, somatostatin, substance P, and neurotensin) in perinuclear/nuclear domains of various cells following their stimulation with extra-cellular agonists (12). In most cases, however, neither the mechanism of GPCR intracellular distribution nor the functional consequence of receptor internalization and nuclear translocation have been explicitly established, with the exception perhaps of angiotensin II (12). The presence of putative nuclear localization signal sequences comprised within C-terminal segments of GPCRs is believed, although equivocally, to pertain to their nuclear sequestration via a Ran-GTP/importin mechanism as suggested for angiotensin II AT1 receptors (12). However, human and rodent LPA1Rs do not bear such amino acid sequences implying that other nuclear import mechanisms may prevail, possibly brought into play following receptor biogenesis and/or endocytosis. In this context, we showed that in pCMVEC membrane, LPA1Rs are partitioned in both clathrin and caveolar endocytic microdomains (Fig. 4); the former domain usually being ascribed for receptor recycling or degradation (57), whereas the latter is thought to serve as a protein carrier responsible for intracellular redistribution of various signaling components (18). Notably, other transmembrane receptors such as 2 adrenergic (58), endothelin ETA (59), and interferon 
receptors (60), have been reported to be dually compartmentalized into caveolae and clathrin-coated pits in cell membranes. On the other hand, our results do not support a nuclear translocation of cell surface LPA1R-evoking inflammatory (COX-2) gene expression mediated through endocytic caveolar pathway (Fig. 5B), despite LPA1-R and caveolin-1 co-localization at the nucleus (Fig. 4, A-C). Rather, our data suggest a major role for released PLA2 products in LPA induced-COX-2 expression (Fig. 5B); these PLA2 products are unlikely to derive from cyclooxygenase, lipoxygenase, or cytochrome P450 enzymatic systems, but possibly LPA. Interestingly, a similar PLA2-dependent mechanism for 
2B-adrenergic receptor-mediated mitogen-activated protein kinases (MAPKs) activation has been recently documented (36).
A novel pathway has recently been reported to explain LPA-evoked signaling of cellular proliferation, which involves MAPK activation through RTK autophosphorylation (e.g. EGF and PDGF receptors) (19, 61). The presumption of auto-induced generation of LPA in extracellular LPA-evoked COX-2 expression is strengthened from experiments describing the limited effectiveness of the RTK-specific inhibitor tyrphostin and MEK kinase inhibitor PD 98059 (Fig. 5B) compared with PLA2 inhibitors. This may suggest that LPA-induced COX-2 up-regulation could proceed via a dual signaling system involving both PLA2 and RTK activities, functioning at least in part independently. Alternatively, affecting the intracellular redox state could stimulate LPA induced-COX-2 gene transactivation. Of relevance, hydrogen peroxide is known to stimulate PLA2 (62) and phospholipase D (63) enzymes, and peroxide concentration is increased in response to exogenous LPA (64). These phospholipases, which also catalyze LPA formation, can be detected in their active state at the nuclear membrane (8-12, 65). Taken together, these observations infer the nucleus as a putative organelle in LPA formation at the vicinity of its cognate peri-nuclear receptors involved in gene transcription.
The nuclear network ensures the commutation and coordination of numerous biochemical signals that are critical in gene expression. In this study, we have begun to address the molecular mechanisms underlying LPA1R-mediated inflammatory gene expression. Our results suggest that perinuclear LPA1Rs regulate iNOS gene expression through Gi/o proteins, calcium channels, and PI 3-kinase/Akt activation in accordance with the increasing ordnance of intrinsic downstream effectors found in the nucleus (10, 12, 66). The lack of potent and selective LPA receptor antagonists currently precludes unequivocal establishment of a sole dependence of LPA nuclear signaling on LPA1Rs and leaves open the possibility that other promiscuous LPA receptors participate meaningfully. Nonetheless, the inference of a predominant role for nuclear LPA1R-transducing transcriptional events stems from our observations. The presence of high levels of immunoreactive LPA1R on plasma and nuclear membranes of hepatic cells (Figs. 2 and 3) and the PTX sensitivity of LPA-evoked calcium and gene responses (Figs. 6A and 7A), independently of the degree of fatty acid saturation of LPA, all reflect common characteristic features of this LPA-R type (1, 4). Overall, the demonstration of intracellular receptor sites for LPA provides an alternative concept to explain the duality in responses to GPCR ligands (e.g. PTH and sphingosine 1-phosphate) in activation of their intracellular versus extracellular targets (12, 67, 52).
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FOOTNOTES |
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A recipient of a Canada Research Chair.
A recipient of a fellowship award from the CIHR and is currently on professorship staff at the University of Sherbrooke. To whom correspondence should be addressed: Dept. of Pharmacology, Université de Sherbrooke, 3001, 12th North Ave., Fleurimont, Québec J1H 5N4, Canada. Tel.: 819-564-5341; Fax: 819-564-5400; E-mail: Fernand.Gobeil{at}USherbrooke.ca.
1 The abbreviations used are: LPA, lysophosphatidic acid; GPCR, G protein-coupled receptor; MAPK, mitogen-activated protein kinase; Erk, extracellular signal-regulated kinase; MEK, MAPK/Erk kinase; PNGase F, peptide N-glycosidase F; PDGF, platelet-derived growth factor; FGF, fibroblast growth factor; EGF, epidermal growth factor; PTH, parathyroid hormone; R, receptor; RTK, receptor tyrosine kinase; PVDF, polyvinylidene difluoride; COX-2, cyclooxygenase-2; iNOS, inducible nitric-oxide synthase; PTX, pertussis toxin; PAF, platelet-activating factor; IL-1, interleukin-1; RT, room temperature; RT-PCR, reverse transcription-PCR; BSA, bovine serum albumin; PBS, phosphate-buffered saline; pCMVEC, porcine cerebral microvascular endothelial cell; PM, plasma membrane; WN, whole nuclei; NE, nuclear envelope; BSAfaf, bovine serum albumin fatty acid-free; oLPA, oleoyl-lysophosphatidic acid; sLPA, stearoyl-lysophosphatidic acid; pLPA, palmitoyl-lysophosphatidic acid.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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