Introduction

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TXA₂ induces many cellular responses, including platelet shape change and aggregation and constriction of bronchial and vascular smooth muscle cells (Negishi et al., 1993). These actions are mediated through interaction with the shared endoperoxide prostaglandin (PG)H₂/TP (Kinsella et al., 1997), a member of the seven-transmembrane domain G-protein-coupled receptor family. The human TP is encoded by a single gene on chromosome 19p13.3 (Nusing et al., 1993). Two isoforms of the receptor exist in humans: the platelet/placental TP, TP (Hirata et al., 1991), and the endothelial isoform, TP (Raychowdhury et al., 1994, 1995). TP and TP, which arise by differential splicing in exon 3, are identical for their first 328 amino acids and differ exclusively in their carboxyl terminal tails.

Functional coupling of the TP to G_q and G₁₁ in vivo recently was confirmed by demonstration of mobilization of Ca⁺⁺_i in response to the TXA₂ mimetic U46619 and the isoprostane 8-epi-prostaglandin F₂ in mammalian cells (Kinsella et al., 1997). However, whereas the two TP isoforms show similar PLC activation, they oppositely regulate adenylyl cyclase activity, with TP activating and TP inhibiting (Hirata et al., 1996). Analysis of TP activation indicates that it is regulated in vivo both by direct phosphorylation and by regulated expression of the TP gene (Kinsella et al., 1994, 1995). Both TP isoforms are phosphorylated in vivo in response to U46619 in a time- and dose-dependent manner (Habib et al., 1997). In Chinese hamster ovary cells, prolonged exposure of TP-expressing cells to U46619 led to an apparent decrease in the number of binding sites expressed, whereas the number of TP sites increased (Yukawa et al., 1997). Furthermore, protein kinase C activation by the phorbol myristic acid inhibited intracellular Ca⁺⁺ mobilization in response to another TP agonist, I-BOP, in cells expressing TP but not TP.

Many GPCRs contain one or two potential N-linked glycosylation sites (defined by Asparagine-X-Serine/Threonine, where X represents any amino acid; Marshall, 1972). Of the fully characterized GPCRs, it is evident that N-linked glycosylation of GPCRs may confer varying functional roles depending on the receptor. For example, site-directed mutagenesis studies on the consensus glycosylation sites of the hamster beta₂-adrenergic receptor (beta₂ AR) amino (N) terminus, Asn⁶ and Asn¹⁵, revealed that although glycosylation does not affect ligand binding, it is important for normal coupling to the G_s/adenylyl cyclase system and for the correct subcellular localization of the receptor on the cell membrane (Rands et al., 1990). On the other hand, inhibition of N-linked glycosylation of the alpha₁-adrenergic receptor has no apparent effect on ligand binding or on membrane insertion of receptor (Sawutz et al., 1987). The retinal rhodopsin receptor has two highly conserved N-linked glycosylation sites. A naturally occurring mutation in one of the N-linked glycosylation consensus sites, at Thr¹⁷, is responsible for a form of the autosomal dominant disorder retinitis pigmentosa (Sung et al., 1991). N-linked glycosylation also has been reported as functionally important in the thrombin receptor. Treatment of fibroblasts with tunicamycin, a specific inhibitor of N-linked glycosylation, inhibited thrombin binding to its receptor (Frost et al., 1991) and inhibited thrombin-induced Ca⁺⁺ mobilization in human T-lymphoblastoid cells in a dose-dependent manner (Tordai et al., 1995).

In the prostanoid receptors, N-linked glycosylation also apparently plays divergent functional roles. For the EP₃ subtype of the PGE₂ receptor, site-directed mutagenesis studies on two potential N-linked glycosylation sites at Asn¹⁶ and Asn¹⁹³ indicated an essential role for glycosylation in determining both affinity and specificity for PGE₂ binding (Huang and Tai, 1995). The human PGI₂ receptor also was glycosylated (Smyth et al., 1996). The human TP isoforms have two potential N-linked glycosylation sites in their NH₂-terminal region at Asn⁴ and Asn¹⁶. These sites are conserved in the mouse, rat and bovine TP receptors (Namba et al., 1992; Kitanaka et al., 1995; Muck et al., 1998). Preparations of TPs purified from human platelets display a diffuse band of 59 kDa on denaturing polyacrylamide gels (Ushikubi et al., 1989), whereas the cDNA for the human TP and TP isoforms codes for proteins with predicted sizes of 37.4 kDa (Hirata et al., 1991) and 44 kDa (Raychowdhury et al., 1994, 1995), respectively. These size discrepancies between the purified receptor and those predicted from the cDNA sequences may be caused by glycosylation at one or both of the potential N-linked sites at Asn⁴ and Asn¹⁶. Site-directed mutagenesis led to the elucidation of the importance of particular residues within the TP, particularly in identifying those residues involved in ligand binding and/or receptor-effector coupling (Funk et al., 1993; D'Angelo et al., 1996, Chiang et al., 1996).

In this study, we sought to determine the importance of N-linked glycosylation on the signaling and surface expression of the human TP isoforms. We used both tunicamycin and endo H treatment of cells combined with site-directed mutagenesis of the Asn⁴ and Asn¹⁶ residues of the TP to explore the significance of N-linked glycosylation on TP function. Our results indicated that N-linked glycosylation of the human TP isoforms is important for ligand binding and for efficient G protein coupling, and that N-linked glycosylation, in at least one site (Asn⁴ or Asn¹⁶), is required for membrane expression.

	Materials and Methods

Top Abstract Introduction Materials & Methods Results Discussion References

Materials. The following chemicals were obtained from Cayman Chemical Company (Ann Arbor, MI): 5-Heptenoic acid, 7-[6-(3-hydroxy-1-octenyl)-2-oxabicyclo [2,2,1] hept-5-yl] [1R-[1,4,5(z), 6(1E,3S*)]-9,11-dideoxy-9,11-methanoepoxy prostaglandin F₂ (U46619); 5-Heptenoic acid, 7-[3-[[Z-[phenylamino) carbonyl] hydrazino] methyl] -7- oxabicyclo [2.2.1] hept -2-yl] ,[1S-[1,2(Z),3,4]] (SQ29,548), thrombin, G418, 1-[2-(5-carboxyoxazol-2-yl)-6-aminobenzofuran-2 oxy}-2-(2'-amino-5'-methylphenoxy)-ethane-N, N,N'N'-tetraacetic acid, pentaacetoxymethyl ester] (FURA2/AM), D-myo-inositol 1,4,5 triphosphate, 3-deoxy-hexa sodium salt (stable analog of IP₃) and ionomycin. [³H]SQ29,548 (50.4Ci/mmol) and ¹²⁵I-labeled goat anti-rabbit IgG (7.61 µCi/µg; NEX 167) were obtained from DuPont NEN (Boston, MA). Gq/G11 (C19) specific antibody was obtained from Santa Cruz Laboratories (Santa Cruz, CA). [³H] cAMP (15-30 Ci/mM) was obtained from American Radiolabeled Chemicals Inc. (St. Louis, MO). Duralon-UV nylon membranes were obtained from Stratagene (LaJolla, CA). Tunicamycin was purchased from Sigma Chemical Inc. (St. Louis, MO); Ultraspec RNA isolation system from Biotecx (Houston, TX). Endo--N-acetylglucosaminidase H, recombinant enzyme, was obtained from Boehringer Mannheim (Basel, Switzerland).

Plasmid construction and site-directed mutagenesis. The plasmids pCMV5 and pCMVTXR, containing the full-length cDNA (nucleotides 4 to +1035) for the human platelet/placental TP, as an EcoR1-Hind111 insert in pCMV5, have been described previously (Kinsella et al., 1994).

Mutator oligonucleotide 1. Mutation N4-Q4. 5' AAC AGG GCC CCA GGG AAC TGC CCT GGG GCC ACA TG 3' (Complementary to coding region).

Mutator oligonucleotide 2. Mutation N16-Q16. 5' TCC CTG GGG CCC TGT TTC CGG CCC ACA CAG ATT ACC CTG GAG 3' where the nucleotides carrying the mutated sequences are underlined.

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Cell culture and transfections. HEL 92.1.3 cells and HEK 293 cells were obtained from the American Type Culture Collection (Manassas, VA). HEL cells were routinely grown in RPMI 1640 medium, 10% FBS. HEK 293 cells were grown in minimal essential medium containing 10% FBS. For transfection studies, HEK 293 cells were plated in 100-mm culture dishes, approximately 48 h before transfection at a density of 1.8 to 2 × 10⁶ cells/dish. Cells were transfected with 10 µg of pADVA and 25 µg of pCMV5 or the pCMV-based vectors with the calcium phosphate/DNA coprecipitation procedure (Kinsella et al., 1997). For transient transfection, cells were harvested 48 h after transfection. To create stable cell lines expressing TP or TP, HEK 293 cells were transfected with 10 µg Sca1-linearized pADVA + 25 µg Pvu1-linearized pcDNA-based vectors with the calcium phosphate/DNA coprecipitation procedure. Forty-eight hours after transfection, G418 (0.8 mg/ml) was added; individual G418-resistant colonies were selected after approximately 21 days and clonal cell lines were expanded.

Radioligand binding studies. Transfected HEK 293 or HEL cells were harvested by centrifugation at 500 × g at 4°C for 5 min, were washed three times in Dulbecco's phosphate-buffered saline and were resuspended in modified Ca⁺⁺/Mg⁺⁺-free HBSSHB buffer, containing 10 mM HEPES, pH7.67, 0.1% bovine serum albumin. Alternatively, to fractionate the cells into their soluble (S100) or membrane (P100) components, washed cells were resuspended and homogenized in HED buffer (20 mM HEPES, pH 7.67, 1 mM EGTA, 0.5 mM dithiothreitol) supplemented with 1 mM phenylmethylsulfonyl fluoride, 10 µM indomethacin. The homogenates were centrifuged at 100,000 × g for 30 min at 4°C, and the membrane fractions (P100) were resuspended in HEDG buffer (20 mM HEPES, pH 7.67, 1 mM EGTA, 0.5 mM dithiothreitol, 100 mM NaCl, 10% glycerol) supplemented with 1 mM phenylmethylsulfonyl fluoride, 10 µM indomethacin. Protein determinations were carried out according to the Bradford assay (Kinsella et al., 1997). For ligand binding studies, protein concentrations in the membrane, fractions or whole cell fractions, were diluted to 1 mg/ml in HEDG or HBSSHB buffer, respectively. Radioligand binding assays were carried out in the presence of the TP antagonist [³H]SQ29,548 (50.4 Ci/mmol) at 30°C for 30 min in 100-µl reactions (containing 100 µg protein/assay, unless otherwise specified) in the presence of 0 to 40 nM [³H]SQ29,548 for Scatchard analyses or in the presence of 20 nM [³H]SQ29,548 for saturation radioligand binding experiments. For competition binding studies, radioligand binding of [³H]SQ29,548 (20 nM, 50.4 Ci/mmol) was carried out in the presence of the following competitor ligands: SQ29,548 (10⁹-10⁶ M); U46619 (10⁸-10⁵ M). In all cases, nonspecific binding was determined in the presence of excess nonlabeled SQ29,548 (10 µM). Reactions were terminated by the addition of 4 ml of ice-cold 10 mM Tris-HCl, pH 7.4, followed by filtration through Whatman GF/C glass filters, and subsequent washing of the filters three times with 10 mM Tris-HCl, pH 7.4, followed by liquid scintillation counting of the filters in 5 ml of scintillation fluid. Radioligand binding data was analyzed with the INPLOT 4 computer program (GraphPad Software Inc., San Diego, CA) to determine the K_d and B_max values.

Endo H studies. HEK1 and HEK3 stable cells were plated at a density of 2 × 10⁶ cells per 100-mm cell culture dish in 8 ml minimal essential medium, 10% FBS. After 24 h, an aliquot (4 µl; 4 mU) of Endo H was added directly to each dish or, as controls, enzyme that had been heat inactivated by boiling for 10 min. Cells were incubated for an additional 24 h at 37°C. Thereafter, cells were harvested by scraping, and membranes were prepared and assayed by radioligand [³H]SQ29,548 binding (20 nM, 50.4 Ci/mmol) as described previously. Data are presented as picomole radioligand bound per milligram of cell protein ± S.E., (i.e. pmol/mg ± S.E.) and are the mean values of four to six experiments.

Northern Blot analysis. Total RNA was isolated from transfected HEK 293 cells with the Ultraspec RNA isolation procedure as recommended by the manufacturers. RNA (10 µg/lane) was analyzed by electrophoresis on 1.1% agarose/formaldehyde/formamide gels followed by transfer and cross-linking to Duralon-UV nylon membranes essentially as described by Sambrook et al. (1989). Northern Blots were screened with radiolabeled RNA riboprobes complementary to the human TP receptor mRNA 3' coding region (nucleotides 474-1032); the probes were prepared by the in vitro transcription of the Not1 linearized plasmid pGEM4TXR with T7 RNA polymerase in the presence of [-³²P]CTP (800 Ci/mmol, 10 mCi/ml), essentially as described (Melton et al., 1984). Filters were hybridized at 70°C in 5× SSC, 50% formamide, 5× Denhardt's solution, 0.2% SDS, 100 µg/ml denatured sonicated salmon sperm DNA for 12 to 16 h. Filters were washed in 0.1× SSC, 0.1% SDS at 70°C for 20 min; incubated with RNase A (1 µg/ml) in 2 × SSC at room temperature for 10 min and washed again at 70°C in 0.1× SSC, 0.1% SDS for 20 min.

Measurement of intracellular Ca⁺⁺ mobilization. Ca⁺⁺_i measurements in either transfected HEK 293 cells or in HEL cells were made by monitoring the intensity of FURA2 fluorescence. For the transfected cells, 48 h after transfection the HEK 293 cells were washed twice in phosphate-buffered saline, resuspended in HBSSHB buffer at 10⁷ cells/ml and incubated in the dark with 5 µM FURA2/AM for 45 min at 37°C. Subsequently the cells were collected by centrifugation (900 × g, 5 min), were washed once in an equal volume of HBSSHB, and were finally resuspended in HBSSHB buffer at 10⁷ cells/ml and kept at room temperature in the dark until use. For each measurement of Ca⁺⁺_i, aliquots of HEK 293 cells were diluted to 0.825 × 10⁶ cells/ml in HBSSHB buffer containing 1 mM CaCl₂.

Measurement of cAMP. Transfected cells were washed three times in ice-cold phosphate-buffered saline; cells (approximately 1-2 × 10⁶ cells) were resuspended in 200 µl HBS (140 mM NaCl, 4.7 mM KCl, 2.2 mM CaCl₂, 1.2 mM KH₂PO₄, 11 mM glucose, 15 mM Hepes-NaOH, pH 7.4) containing 1 mM 3-isobutyl-1-methylxanthine and preincubated at 37°C for 10 min. Thereafter, cells were stimulated in the presence of 1 µM U46619 (from a 5 µM U46619 stock, 50 µl) or in the presence of HBS (50 µl) at 37°C for 10 min. Reactions were terminated by heat inactivation (100°C, 5 min) and the level of cAMP produced was quantified by radioimmunoassay with the cAMP-binding protein from bovine adrenal medulla, essentially as described by Farndale et al., (1992). Protein determinations were carried out with the Bradford assay (Bradford, 1976). Levels of cAMP produced by U46619-stimulated cells relative to basal stimulation, in the presence of HBS, were expressed in picomoles cAMP per milligram cell protein ± S.E.M. (pmol/mg ± S.E.M.) and as fold stimulation to basal (fold increase ± S.E.M.). The data presented are representative of four independent experiments.

Data analyses. Radioligand binding data were analyzed with the INPLOT 4 computer program (GraphPad Software Inc.) to determine the K_d and B_max values. Statistical analyses were carried out with the unpaired Student's t test with the Statworks Analysis Package.

	Results

Top Abstract Introduction Materials & Methods Results Discussion References

Effect of inhibition of N-linked glycosylation on TP expression. HEL cells were incubated in the presence of tunicamycin, a direct inhibitor of N-linked glycosylation, to investigate initially whether inhibition of N-linked glycosylation had any effect on TP expression. Tunicamycin has been shown to inhibit the incorporation of oligosaccharides into proteins with little or no effect on de novo protein synthesis itself and to exhibit low general toxicity in the effective concentration range of 1 to 20 µg/ml (Struck and Lennarz, 1977). Thus, a time course assay (0-48 h) was used initially to examine the effect of tunicamycin on TP expression used at nontoxic concentrations of 2 µg/ml. When used at these concentrations, tunicamycin was confirmed to be noncytotoxic to both HEL and HEK 293 cells with the Trypan Blue dye-exclusion assay. TP expression was monitored by radioligand binding assays with the radiolabeled TP antagonist [³H]SQ29,548. The platelet-like HEL cells are known to express TPs (Kinsella et al., 1994); with a reverse transcriptase polymerase chain reaction approach, we have confirmed that HEL cells express both the TP and TP isoforms (Miggin and B.T. Kinsella, unpublished). Control HEL cells, not treated with tunicamycin, displayed specific binding of [³H]SQ29,548 of 19.9 ± 5.9 fmol/mg cell protein. In the presence of tunicamycin, TP expression was reduced to levels of 45 ± 3.2% relative to the nontreated cells after 12 h incubation (fig. 1), indicative of a role of N-linked glycosylation in TP expression or function.

View larger version (14K): [in this window] [in a new window]   Fig. 1.   Effect of tunicamycin on TP expression in HEL cells. HEL cells grown in RPMI, 10% FBS (0.5 × 106 cells/ml) were incubated in the presence of tunicamycin (2 µg/ml) for 0 to 48 h. At the indicated times, aliquots were removed and cells were assayed for TP expression in the presence of [3H]SQ29,548 (50.4Ci/mmol, 20 nM) as described under "Materials and Methods." Results are expressed as a percentage TP expression relative to the control HEL cells that had not been incubated with tunicamycin ± standard error (% expression ± S.E.); n = 4). TP expression in the control HEL cells was 19.9 ± 5.90 fmol/mg cell protein.

View larger version (16K): [in this window] [in a new window]   Fig. 2.   Effect of tunicamycin on TP and TP expression in HEK 293 cells. HEK1 or HEK3 cells stably transfected with the cDNA for TP (Kd for [3H]SQ29,548 = 9.43 ± 0.92 nM, Bmax = 3.02 ± 0.43 pmol/mg) or TP (Kd for [3H]SQ29,548 = 8.44 ± 1.44 nM, Bmax = 3.24 ± 0.33 pmol/mg, n = 6), respectively, were incubated in the presence of tunicamycin (2 µg/ml) for 0 to 48 h. At the indicated times, cells were harvested and assayed for TP () or TP () expression in the presence of [3H] SQ29,548 (50.4Ci/mmol, 20 nM). In each case, results are expressed as percentage TP expression relative to the control cells that had not been incubated with tunicamycin (percentage expression ± standard error of mean = % expression ± S.E.M.; n  3). Levels of TP in control HEK1 cells were 3.29 ± 0.30 pmol/mg cell protein and for TP expression in control HEK3 cells were 2.88 ± 0.32 pmol/mg cell protein.

Effect of inhibition of N-linked glycosylation on intracellular Ca⁺⁺ mobilization. Functional coupling of the TP or TP isoforms to PLC activation was assessed by monitoring mobilization of Ca⁺⁺_i in FURA2/AM-loaded HEK1 cells and HEK3 cells, respectively, in response to the TXA₂ mimetic U46619. It has been previously reported that for efficient TP coupling to Ca⁺⁺_i mobilization in HEK 293 cells, it was necessary to coexpress a member of the G_q (G_q or G₁₁) family (Kinsella et al., 1997). Thus, HEK1 or HEK3 cells were transiently cotransfected with the cDNA for G₁₁; where indicated, tunicamycin (2 µg/ml) was added directly to the cells 2 to 44 h after transfection. Thereafter, tunicamycin-treated or nontreated cells were harvested 48 h after transfection, and Ca⁺⁺_i mobilization, was measured in FURA2/AM-loaded cells by spectrofluorometry in response to U46619 (2 µM). Positive overexpression of G₁₁ in HEK1 and HEK3 cells was confirmed by Western Blot analyses (data not shown). Both HEK1- and HEK3-transfected cells displayed a rapid transient rise in Ca⁺⁺_i mobilization in response to U46619 stimulation (fig. 3 A and B). However, incubation of HEK1 or HEK3 cells with tunicamycin resulted in time-dependent reductions in Ca⁺⁺_i mobilization, respectively. Treatment of HEK1 cells with tunicamycin for 48 h reduced Ca⁺⁺_i mobilization from 96.9 ± 16.3 nM to 15.7 ± 7.09 nM, whereas treatment of HEK3 cells with tunicamycin for 48 h reduced Ca⁺⁺_i mobilization from 94.1 ± 21.1 nM to 11.7 ± 4.99 nM. Tunicamycin treatment of HEK1 or HEK3 cells had no effect on Ca⁺⁺_i mobilization in response to the IP₃-stable analog or in response to ionomycin, indicating that tunicamycin treatment, per se, does not interfere with Ca⁺⁺_i mobilization (data not shown). Moreover, tunicamycin was confirmed to be noncytotoxic to HEK 293 cells with the Trypan Blue dye-exclusion assay. Thus, the effects of tunicamycin on TP expression/TP ligand binding and on Ca⁺⁺_i mobilization in response to U46619 are evidence that N-linked glycosylation of TP and TP is functionally important.

View larger version (25K): [in this window] [in a new window]   Fig. 3.   Effect of tunicamycin on intracellular Ca++ mobilization in HEK 293 cells stably expressing either TP and TP. HEK1 cells (panel A) or HEK3 cells (panel B) stably transfected with TP or TP, respectively, were transiently cotransfected with the cDNA for G with the Ca++ phosphate DNA coprecipitation procedure. Tunicamycin (2 µg/ml) was added to duplicate transfected cultures 2 h after transfection and cells were incubated in the presence (+) or absence () of tunicamycin for an additional 48 h. Alternatively, cells were incubated in the presence of tunicamycin for 0, 4 or 24 h before harvesting (Insets, panels A and B). Harvested cells then were preloaded with FURA2/AM before stimulation with the TP mimetic U46619 (2 µM) at the times indicated by the arrows. FURA2/AM fluorescence was recorded on a Perkin-Elmer LS50-B spectrofluorometer at excitation wavelengths of 340 nM and 380 nM and emission wavelengths of 510 nM. The ratio of fluorescence at 340 nM to fluorescence at 380 nM is a measure of Ca++ mobilization assuming a Kd of 225 nM Ca++ for FURA2/AM. The profiles presented are representative of at least three independent experiments. Data were calculated as changes in intracellular Ca++ mobilized ([Ca++]i ± S.E.M.); average resting or basal [Ca++]i ranged from 80 to 120 nM. Levels of intracellular Ca++ mobilized ([Ca++]i) were: (A) HEK1 cells, () tunicamycin, 96.9 ± 16.3 nM (n = 11); HEK1 cells, (+) tunicamycin, 15.7 ± 7.09 nM; (B) HEK3 cells, () tunicamycin, 94.1 ± 21.1 nM (n = 9); HEK3 cells, (+) tunicamycin, 11.7 ± 4.99 nM.

Site-directed mutagenesis of N-linked glycosylation sites of TP. To establish definitively whether one or both putative N-linked glycosylation sites are functionally required, site-directed mutagenesis was carried out on the TP isoform whereby the pivotal asparagine (N) residues were mutated to glutamine (Q) residues at amino acid residues N4, N16 or both to generate the mutated receptors TP^N4-Q4, TP^N16-Q16, TP^{N4,N16-Q4,Q16}, respectively. HEK 293 cells were transiently transfected with the cDNAs for the wild-type and mutant TP receptors and the level of TP expression was assessed by saturation radioligand ([³H]SQ29,548) binding. Levels of TP expression for the wild-type TP were 2.43 ± 0.32 pmol/mg cell protein. However in the mutant receptors, specific [³H]SQ29,548 binding was reduced to 47.6 ± 3.2% (TP^N4-Q4), 58.4 ± 7.3% (TP^N16-Q16) and 8.3 ± 1.0% (TP^{N4,N16-Q4,Q16}) relative to the wild-type receptor (100%). To address whether this reduction in binding exhibited by the mutant receptors was caused by a change in the affinity (K_d) or B_max for [³H]SQ29,548, radioligand binding isotherms were carried out and the results were analyzed by Scatchard analysis (table 1 and fig. 4). Consistent with the previous results, although there were no observed differences in the affinity (K_d) of each of the mutant receptors for [³H]SQ29,548, a substantial reduction occurred in maximal binding, with the double mutant receptor (TP^{N4,N16-Q4,Q16}) only retaining approximately 8% binding relative to the nonmutant receptor (table 1). In general, HEK 293 cells transfected with the control vector (pCMV5) had between 2 and 5% radioligand ([³H]SQ29,548) binding, representing approximately 48 to 120 fmol [³H]SQ29,548/mg protein, relative to that expressed by cells transfected with the wild-type TP receptor. Thus, in the double mutant, a considerable portion (48-120 fmol [³H]SQ29,548/mg protein) of radioligand binding associated with this receptor may be caused by endogenous receptor expressed in HEK 293 cells.

View larger version (16K): [in this window] [in a new window]   Fig. 4.   Radioligand Binding. HEK 293 cells were transiently transfected with the plasmids coding for the wild-type TP or mutant TPN4-Q4, TPN16-Q16, TPN4,N16-Q4,Q16 receptors as indicated. Radioligand binding assays on whole cells (75 µg/assay) were carried out in the presence of the TP antagonist [3H]SQ29,548 (50.4Ci/mmol, 0-40 nM). A typical radioligand binding isotherm (panel A) and Scatchard plot (panel B) for each receptor type is presented. Radioligand binding data were analyzed with the INPLOT 4 computer program (GraphPad Software Inc.) to determine the Kd and Bmax values.

View larger version (30K): [in this window] [in a new window]   Fig. 5.   Subcellular distribution of TP receptors in transfected HEK 293 cells. HEK 293 cells were transiently transfected with the cDNAs for TP (wild-type), TPN4-Q4 (N4-Q4), TPN16-Q16 (N16-Q16), TPN4,N16-Q4,Q16 (N4,N16-Q4,Q16); cells were harvested 48 h after transfection. Aliquots (50-100 µg/assay) of the P100 () or S100 () fractions were assayed for TP expression by saturation radioligand binding with [3H]SQ29,548 (50.4Ci/mmol, 20 nM) as described under "Materials and Methods"; from these data, values for Total (; Total = P100 + S100) expression were calculated. Results are expressed as: (A) [3H]SQ29,548 bound (pmol/mg ± S.E.M.) or (B) percentage TP expression in the individual P100 or S100 fractions relative to total expression (% expression ± S.E.M.). The data presented are the mean values of three independent experiments. Actual levels of total TP expression, taken as 100% expression in the case of each receptor type, were: TP (wild-type) 2.90 ± 0.43 pmol/mg; TPN4-Q4, 1.38 ± 0.20 pmol/mg; TPN16-Q16, 1.70 ± 0.26 pmol/mg; TPN4,N16-Q4,Q16, 0.24 ± 0.03 pmol/mg. Levels of TP expression in the respective P100 and S100 fractions were: TP (Wild type) 2.45 ± 0.15 pmol/mg and 0.44 ± 0.15 pmol/mg; TPN4-Q4, 1.08 ± 0.08 pmol/mg and 0.31 ± 0.09 pmol/mg; TPN16-Q16, 1.38 ± 0.07 pmol/mg and 0.31 ± 0.06 pmol/mg; TPN4,N16-Q4,, 0.14 ± 0.02 pmol/mg and 0.10 ± 0.02 pmol/mg.

Northern Blot analysis. Northern Blot analysis was carried out to address whether the substantial reduction in [³H]SQ29,548 binding by the mutant TP receptors could be accounted for by reductions in gene expression (fig. 6). RNA was isolated from HEK 293 cells transiently transfected with wild-type or mutant TP receptors, or with the vector, pCMV5, and Northern Blots were probed with a ³²P-radiolabeled antisense RNA transcript based on the 3' coding region of the TP mRNA. Equal loading of RNA (fig. 6A) and equal intensity of the hybridization signal by cells transfected with either wild-type TP or mutant TPs (TP^N4-Q4, TP^N16-Q16, TP^{N4,N16-Q4,Q16}) but not with the vector pCMV5 (fig. 6B) was confirmed by densitometric scanning of the gel and autoradiogram, respectively.

View larger version (73K): [in this window] [in a new window]   Fig. 6.   Northern Blot analysis of transfected HEK 293 cells. Total RNA isolated from HEK 293 cells transfected with pCMVTXR (lane 1), pCMVTXRN4-Q4 (lane 2), pCMVTXRN16-Q16 (lane 3), pCMVTXRN4,N16-Q4,Q16 (lane 4), pCMV5 (lane 5) were analyzed by formaldehyde/formamide/agarose gel electrophoresis (panel A) followed by Northern Blot analysis (panel B). Equal aliquots (10 µg) of total RNA were analyzed for each sample. The Northern Blot (panel B) was screened vs. a 32P-radiolabeled antisense riboprobe based on human TP, as outlined under "Materials and Methods." The position of the 18S and 28S ribosomal RNAs are indicated on the left and right of panels A and B, respectively.

Analyses of intracellular signaling. To address whether the observed differences in radioligand ([³H]SQ29,548) binding by the mutant TP receptors was confined to a reduced ability to bind the antagonist SQ29,548 exclusively or whether there was also a reduced ability to interact with other TP ligands, competition binding profiles were performed. Competition binding studies of [³H]SQ29,548 were carried out in the presence of U46619 and SQ29,548 on HEK 293 cells transfected with TP wild-type or mutant receptors. For the competing ligand U46619, ligand binding by the wild-type TP and each of the mutant receptors (TP^N4-Q4, TP^N16-Q16, TP^{N4,N16-Q4,Q16}) were not significantly different with typical K_i values between 0.24 and 0.77 µM U46619 (table 2). Similarly, no significant differences were obtained with SQ29,548 as competing ligand by the wild-type or mutant TP receptors with typical K_i values ranging between 0.76 and 1.60 × 10⁸ M SQ29,548 (table 2). The K_i values reported here compare favorably with those reported previously by Habib et al. (1997) and Muck et al. (1998).

View this table: [in this window] [in a new window]   TABLE 2 Competition binding profiles performed to determine observed differences in radioligand [3H]SQ29,548 binding by the mutant TP receptors

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View larger version (21K): [in this window] [in a new window]   Fig. 7.   Effect of site-directed mutagenesis of N-linked glycosylation sites of TP on intracellular Ca++ mobilization in HEK 293 cells. HEK 293 cells were transiently cotransfected with the cDNA coding the G with TP (Wild Type), TPN4-Q4 (N4-Q4), TPN16-Q16 (N16-Q16), TPN4,N16-Q4,Q16 (N4,N16-Q4,Q16) or with the control vector pCMV5. Cells were harvested 48 h after transfection, were preloaded with FURA2/AM and stimulated with U46619 (2 µM) at the times indicated by the arrows. FURA2/AM fluorescence was recorded on a Perkin-Elmer LS50-B spectrofluorometer at excitation wavelengths of 340 nM and 380 nM and emission wavelengths of 510 nM. The ratio of fluorescence at 340 nM to fluorescence at 380 nM is a measure of Ca++ mobilization assuming a Kd of 225 nM Ca++ for FURA2/AM. The profiles presented are representative of at least three independent experiments. Data were calculated as changes in intracellular Ca++ mobilized ( [Ca++]i ± S.E.M.); average resting or basal [Ca++]i were in the range of 80 to 120 nM. Levels of intracellular Ca++ mobilized ( [Ca++]i) were: TP (wild-type), 61.6 ± 16.1 nM; TPN4-Q4 (N4-Q4), 48.4 ± 12.9 nM; TPN16-Q16 (N16-Q16), 54.7 ± 17.5 nM; TPN4,N16-Q4,Q16 (N4,N16-Q4,Q16), 26.0 ± 7.01 nM; pCMV5, 10.1 ± 5.29 nM.

	Discussion

Top Abstract Introduction Materials & Methods Results Discussion References

In this study, we investigated the potential role of asparagine N-linked glycosylation on the subcellular localization, ligand binding properties, G-protein coupling and second messenger signaling of the human TP isoforms. N-linked glycosylation is important in the correct functioning and subcellular distribution of many, but not all, membrane-associated proteins such as the acetylcholine (Giovanelli et al, 1991), transferrin (Ralton et al, 1989) and IgD (Swenson et al, 1993) receptors. When considering this variation in functional responses, it was of interest to determine the significance, if any existed, of alteration in the glycosylation status of both the TP and TP receptor subtypes.

Several genetic bleeding disorders have been described in humans in which the individual's platelets were unresponsive to TXA₂. The molecular basis of one of these dominantly inherited bleeding disorders recently was reported to be caused by a single amino acid substitution (Arg⁶⁰-Leu⁶⁰) in the first cytoplasmic loop of the TP (Hirata et al., 1994). The mutant TP receptor, when expressed in Chinese hamster ovary cells, showed decreased agonist-induced second messenger signaling despite its normal ligand binding affinities. Thus, in this study, a series of experiments were undertaken to investigate possible loss of TP receptor function either because of inhibition of glycosylation in vivo with tunicamycin or transfection of mammalian cells with site-directed mutants of the TP receptor in which either one or both consensus N-linked glycosylation sites (Hirata et al., 1991., Raychowdhury et al., 1994, 1995) have been destroyed. The antibiotic tunicamycin is a highly selective inhibitor of N-linked but not O-linked glycosylation, blocking the enzymatic transfer of N-acetylglucosamine 1-phosphate to dolichol-mono-phosphate within 1 h after treatment, thereby preventing the N-linked glycosylation of various proteins (Tkacz and Lampen, 1975; Elbein,1987). However, it has little or no effect on de novo protein synthesis itself and exhibits low general toxicity in the effective concentration range of 0.1 to 20 µg/ml (Struck and Lennarz, 1977). Calnexin and calreticulin are molecular chaperones that anchor proteins in the endoplasmic reticulum until they have folded correctly by recognizing their carbohydrate portions (Wada et al., 1997; Gahmberg and Tolvanen, 1996). Tunicamycin treatment blocks calnexin and calreticulin interactions with their targets. This may result in incorrectly folded polypeptides, increased protein degradation and mislocalization of newly synthesized proteins.

When HEL cells were treated with tunicamycin, we observed a time-dependent reduction in TP receptor expression, measured in radioligand binding assays, with [³H]SQ29,548, to yield a level of binding of 45% relative to control, nontreated cells after 12 h incubation. We have confirmed that HEL cells express the mRNAs coding for both the TP and TP isoforms (S.M. Miggin and B.T. Kinsella., unpublished). To address whether this effect may be isoform specific, stable HEK 293 cells, which exclusively express either the TP or TP isoform, were generated. Treatment of these stable cell lines with tunicamycin resulted in a similar time-dependent decline in TP and TP expression. After a 4-h treatment of HEK1 and HEK3 cells with tunicamycin, TP and TP expression was reduced to 78% and 80%, respectively, relative to the corresponding nontreated control cells. However, on prolonged exposure (24-48 h), the HEK3 cells were significantly more sensitive to tunicamycin treatment than HEK1 cells. The physiologic significance of this is unclear but may reflect differences in TP and TP protein turnover rates or indicate additional functional differences between the two receptor isoforms, such as intracellular trafficking or interaction with endoplasmic reticulum chaperones such as calnexin and calreticulin (Wada et al., 1997). Tunicamycin treatment of the HEK1 and HEK3 cells cotransfected with G₁₁ also resulted in substantial reduction in Ca⁺⁺_i mobilization in FURA2/AM-loaded cells in response to the TXA₂ mimetic U46619, but had no effect on Ca⁺⁺_i mobilization in response to ionomycin or a stable analog of IP₃. Thus, inhibition of TP and TP glycosylation with tunicamycin resulted in reduced ligand binding and reduced Ca⁺⁺_i mobilization. However, as previously stated, these data do not fully exclude other effects such as alteration in receptor folding or secondary effects on other proteins. The glycosylation of membrane proteins generally is believed to help in protein folding and correct assembly in the endoplasmic reticulum and migration to the plasma membrane (Gahmberg and Tolvanen, 1996).

The significance of the N-linked glycosylation sites at positions 4 and 16 of TP was confirmed by site-directed mutagenesis of either one or both asparagine (N) residues of these sites to glutamine (Q) residues in the TP isoform. After transient transfection of the wild-type or mutant receptors into HEK 293 cells, the level of TP expression was determined by radioligand binding studies. Specific [³H]SQ29,548 binding was reduced to 47% (TP^N4-Q4), 58% (TP^N16-Q16) and 8.3% (TP^{N4,N16-Q4,Q16}) relative to wild-type TP receptor. Scatchard analysis indicated that whereas there was no significant change in the affinity (K_d) of either mutant receptor for SQ29,548, there were substantial reductions in maximal binding (B_max) with that of double mutant receptor (TP^{N4,N16-Q4,Q16}) being reduced by 92% relative to the wild type TP. Furthermore, comparison of [³H]SQ29,548 binding by either U46619 or SQ29,548 failed to demonstrate any difference in K_i values for these ligands between the wild-type or mutant forms of TP, confirming that glycosylation per se does not have a role in determining TP ligand affinity or specificity.

The observed reduction in SQ29,548 binding for the mutant TPs could be caused by altered TP expression or by altered subcellular localization of mutated receptors between the membrane and soluble fractions of the cell relative to that of the wild-type TP. Currently, antibodies to the human TP isoforms are not available to this laboratory, which precludes us from directly determining actual TP protein expression levels or the relative cell surface expression of the wild-type and mutant receptors. However, Northern Blot analysis revealed that TP gene expression was equal among all the TP receptors transiently expressed in HEK 293 cells, which indicates that differential gene expression also may not account for the observed reduction in SQ29,548 binding displayed by the mutant receptors (TP^N4-Q4, TP^N16-Q16, TP^{N4,N16-Q4,Q16}) relative to the wild-type TP. Fractionation of transfected cells into their membrane (P100) and soluble (S100) fractions revealed similar corresponding reductions in total, membrane and cytosolic expression for each of the mutants relative to the wild-type receptor. Such a reduction in membrane expression is not, on its own, sufficient to account for the observed reduction in SQ29,548 binding in the double mutant receptor (92% reduction relative to the wild-type TP) which suggests that glycosylation may be required for ligand binding. Endo H treatment of HEK1 or HEK3 cells, stably expressing either TP or TP, respectively, resulted in approximately 50% reductions in radioligand binding, by either receptor isoform, confirming a direct role for N-linked glycosylation in ligand ([³H]SQ29,548) binding by the TPs. In the EP₃ isoform of the PGE₂ receptor, another prostanoid receptor, N-linked glycosylation has been reported to be important not only in determining the affinity, but also the specificity of the EP₃ receptor for its ligands (Huang and Tai, 1995). In general, the putative seventh transmembrane domain forms a critical portion of the ligand binding pocket for G-protein-coupled receptors (Baldwin, 1994) and is highly related among the whole prostanoid receptor family (Ushikubi et al., 1995). Previous site-directed mutagenesis studies confirmed that amino acid residues within the putative seventh transmembrane domain of the TP are critical for ligand binding (Funk et al., 1993). In addition, D'Angelo et al. (1996) reported that conservation of cysteine (C) residues (C105 and C183) located within the first and second extracellular regions, respectively, are essential for ligand binding; whereas C103, also located within the first extracellular loop, is required for optimal ligand binding affinity, capacity and cell signaling by TP. Thus, from our current studies, it is evident that conservation of N-linked glycosylation sites at Asn⁴ and Asn¹⁶, located within the extracellular amino-terminal region of TP, may be required for optimal ligand binding.

The TP functionally couples in vivo to members of the G_q (G_q and G₁₁) family of heterotrimeric G proteins mediating activation of the beta isoforms of PLC and leading to release of Ca⁺⁺ from intracellular stores (Kinsella et al., 1997). For some GPCRs, N-linked glycosylation has been reported to affect G-protein coupling and subsequent second messenger generation (Frost et al., 1991.; Rands et al., 1990). Tunicamycin treatment of HEK1 or HEK3 cells, transiently cotransfected with G₁₁, greatly reduced U46619-induced Ca⁺⁺ mobilization in these cells compared with the untreated control cells. In confirmation experiments, U46619-mediated Ca⁺⁺ mobilization in HEK 293 cells transiently cotransfected with either the wild-type (TP) or mutant receptors, and G₁₁ were reduced to 78.5% (TP^N4-Q4), 88.8% (TP^N16-Q16) or 42.2% (TP^{N4,N16-Q4,Q16}) relative to that of the wild-type receptor. Similarly, we also assessed the possible consequences of site-directed mutagenesis of the N-linked glycosylation sites of the TP on its ability to couple to adenylyl cyclase via Gs (Hirata et al., 1996). As observed with mobilization of intracellular Ca⁺⁺, adenylyl cyclase activity was modestly reduced in cells expressing the single mutants (88.1% for TP^N4-Q4 or 75.2% for TP^N16-Q16) and reduced to 39.6% in cells expressing the double mutant (TP^N4,N16-) compared to the wild-type TP. Thus, whereas the fully deglycosylated TP mutant has a 92% reduction in relative expression, as determined by saturation radioligand binding studies, the nonglycosylated receptor (TP^N4,N16-) does retain some functional activity and signal transducing properties, as was indicated by both the Ca⁺⁺ mobilization studies and cAMP assays. This may indicate that the overall structure, folding or topography of the fully nonglycosylated receptor maintains some ligand binding properties but retains a substantial ability to couple to G_q and G_s with concomitant activation of the respective signal transduction pathways.