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• W2085125612 abstract "Syntaxin 1C is an alternative splice variant lacking the transmembrane domain of HPC-1/syntaxin 1A. We found previously that syntaxin 1C is expressed as a soluble protein in human astroglioma (T98G) cells, and syntaxin 1C expression is enhanced by stimulation with phorbol 12-myristate 13-acetate (PMA). However, the physiological function of syntaxin 1C is not known. In this study, we examined the relationship between syntaxin 1C and glucose transport. First, we discovered that glucose transporter-1 (GLUT-1) was the primary isoform in T98G cells. Second, we demonstrated that glucose uptake in T98G cells was suppressed following an increase in endogenous syntaxin 1C after stimulation with PMA, which did not alter the expression levels of other plasma membrane syntaxins. We further examined glucose uptake and intracellular localization of GLUT-1 in cells that overexpressed exogenous syntaxin 1C; glucose uptake via GLUT-1 was inhibited without affecting sodium-dependent glucose transport. The value of Vmax for the dose-dependent uptake of glucose was reduced in syntaxin 1C-expressing cells, whereas there was no change in Km. Immunofluorescence studies revealed a reduction in the amount of GLUT-1 in the plasma membrane in cells that expressed syntaxin 1C. Based on these results, we postulate that syntaxin 1C regulates glucose transport in astroglioma cells by changing the intracellular trafficking of GLUT-1. This is the first report to indicate that a syntaxin isoform that lacks a transmembrane domain can regulate the intracellular transport of a plasma membrane protein. Syntaxin 1C is an alternative splice variant lacking the transmembrane domain of HPC-1/syntaxin 1A. We found previously that syntaxin 1C is expressed as a soluble protein in human astroglioma (T98G) cells, and syntaxin 1C expression is enhanced by stimulation with phorbol 12-myristate 13-acetate (PMA). However, the physiological function of syntaxin 1C is not known. In this study, we examined the relationship between syntaxin 1C and glucose transport. First, we discovered that glucose transporter-1 (GLUT-1) was the primary isoform in T98G cells. Second, we demonstrated that glucose uptake in T98G cells was suppressed following an increase in endogenous syntaxin 1C after stimulation with PMA, which did not alter the expression levels of other plasma membrane syntaxins. We further examined glucose uptake and intracellular localization of GLUT-1 in cells that overexpressed exogenous syntaxin 1C; glucose uptake via GLUT-1 was inhibited without affecting sodium-dependent glucose transport. The value of Vmax for the dose-dependent uptake of glucose was reduced in syntaxin 1C-expressing cells, whereas there was no change in Km. Immunofluorescence studies revealed a reduction in the amount of GLUT-1 in the plasma membrane in cells that expressed syntaxin 1C. Based on these results, we postulate that syntaxin 1C regulates glucose transport in astroglioma cells by changing the intracellular trafficking of GLUT-1. This is the first report to indicate that a syntaxin isoform that lacks a transmembrane domain can regulate the intracellular transport of a plasma membrane protein. The protein machinery that regulates intracellular transport and vesicle formation, docking, and fusion has been the focus of intense research over the last few years. The SNARE 1The abbreviations used are: SNARE, soluble N-ethylmaleimide-sensitive fusion protein attachment protein receptor; 2-DG, 2-deoxyglucose; DMEM, Dulbecco's modified Eagle's medium; FCS, fetal calf serum; GLUT, glucose transporter; HA, hemagglutinin; PKC, protein kinase C; PMA, phorbol 12-myristate 13-acetate; SGLT, sodium-dependent glucose transporter; Syn, syntaxin; PBS, phosphate-buffered saline; RT, reverse transcription; nt, nucleotide; CNS, central nervous system; SNAP, soluble NSF attachment protein; VAMP, vesicle-associated membrane protein. hypothesis (soluble N-ethylmaleimide-sensitive fusion protein (NSF) attachment protein receptor) constitutes a widely accepted model in which dynamic interactions among proteins within the acceptor (t-SNARE: syntaxin and SNAP-25) and donor (v-SNARE: VAMP) compartments control exocytosis (1Sollner T. Whiteheart S.W. Brunner M. Erdjument-Bromage H. Geromanos S. Tempst P. Rothman J.E. Nature. 1993; 362: 318-324Crossref PubMed Scopus (2623) Google Scholar, 2Rothman J.E. Nature. 1994; 372: 55-63Crossref PubMed Scopus (2007) Google Scholar). Recent studies have revealed that syntaxins function in a wide variety of cells and tissues, including neurons, endocrine glands, amphibian ectodermal cells, epithelial cells, cells of the immune system, platelets, and yeast (3Burgoyne R.D. Morgan A. Physiol. Rev. 2003; 83: 581-632Crossref PubMed Scopus (543) Google Scholar). Consequently, a unified role for the SNARE complex in the docking and fusion of vesicles during intracellular trafficking, as well as in nerve terminals, has been proposed. To date, 18 members of the mammalian syntaxin family have been identified, all of which localize to specific membrane compartments via a transmembrane domain at the C terminus. In contrast to the localization of syntaxins 5-18 to different intracellular compartments, such as the Golgi and post-Golgi apparatus (4Jahn R. Sudhof T.C. Annu. Rev. Biochem. 1999; 68: 863-911Crossref PubMed Scopus (1021) Google Scholar), syntaxins 1-4 are restricted predominantly to the plasma membrane, where they mediate constitutive and regulated vesicle trafficking to the cell surface (4Jahn R. Sudhof T.C. Annu. Rev. Biochem. 1999; 68: 863-911Crossref PubMed Scopus (1021) Google Scholar). All syntaxins have a coiled-coil helix domain (called H3 in syntaxin 1A) next to the transmembrane domain at the C terminus. The H3 domain is a highly conserved region that interacts with several different SNARE proteins, including SNAP-25, VAMP, and α-SNAP, and to some extent, nSec-1/Munc-18 (4Jahn R. Sudhof T.C. Annu. Rev. Biochem. 1999; 68: 863-911Crossref PubMed Scopus (1021) Google Scholar). Syntaxin 1C is an alternative splice variant of HPC-1/syntaxin 1A. Syntaxin 1A is involved in the docking of synaptic vesicles at active zones in neurons (5Inoue A. Obata K. Akagawa K. J. Biol. Chem. 1992; 267: 10613-10619Abstract Full Text PDF PubMed Google Scholar, 6Bennett M.K. Calakos N. Scheller R.H. Science. 1992; 257: 255-259Crossref PubMed Scopus (1073) Google Scholar), and is deleted hemizygously in patients with the neurodevelopmental disorder, Williams syndrome (7Jagadish M.N. Tellam J.T. Macaulay S.L. Gough K.H. James D.E. Ward C.W. Biochem. J. 1997; 321: 151-156Crossref PubMed Scopus (30) Google Scholar, 8Nakayama T. Matsuoka R. Kimura M. Hirota H. Mikoshiba K. Shimizu Y. Shimizu N. Akagawa K. Cytogenet Cell Genet. 1998; 82: 49-51Crossref PubMed Scopus (35) Google Scholar). In a previous study, we demonstrated that syntaxin 1C is expressed as a soluble protein in astroglioma cells (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). While the N-terminal domain of syntaxin 1C is the same as that of syntaxin 1A, the functionality of the H3 and transmembrane domain has been lost, caused by the generation of a novel 34-residue C-terminal domain by the insertion of a 91-bp splicing region. Several other isoforms of syntaxin that lack a transmembrane domain by alternative splicing have been identified, namely syntaxin 2D, syntaxin 3D, and syntaxin 16C (10Quinones B. Riento K. Olkkonen V.M. Hardy S. Bennett M.K. J. Cell Sci. 1999; 112: 4291-4304Crossref PubMed Google Scholar, 11Ibaraki K. Horikawa H.P. Morita T. Mori H. Sakimura K. Mishina M. Saisu H. Abe T. Biochem. Biophys. Res. Commun. 1995; 211: 997-1005Crossref PubMed Scopus (46) Google Scholar, 12Simonsen A. Bremnes B. Ronning E. Aasland R. Stenmark H. Eur. J. Cell Biol. 1998; 75: 223-231Crossref PubMed Scopus (100) Google Scholar), but the function of syntaxin isoforms that lack a transmembrane domain is unknown. Facilitative glucose transporters (GLUTs) are proteins that regulate the entry of glucose into cells and maintain cell metabolism and homeostasis throughout the periphery and brain (13Vannucci S.J. Maher F. Simpson I.A. Glia. 1997; 21: 2-21Crossref PubMed Scopus (529) Google Scholar). There are at least six different GLUT genes with differential tissue distributions, subcellular localizations, and kinetics for glucose uptake (14Mueckler M. Eur. J. Biochem. 1994; 219: 713-725Crossref PubMed Scopus (953) Google Scholar). In the brain, there are two GLUT isoforms, namely GLUT-1 and GLUT-3. GLUT-1 appears during early embryogenesis and is required for cell metabolism and homeostasis in glial cells (13Vannucci S.J. Maher F. Simpson I.A. Glia. 1997; 21: 2-21Crossref PubMed Scopus (529) Google Scholar, 15Olson A.L. Pessin J.E. Annu. Rev. Nutr. 1996; 16: 235-256Crossref PubMed Scopus (388) Google Scholar). GLUT-3 is found primarily in neurons (15Olson A.L. Pessin J.E. Annu. Rev. Nutr. 1996; 16: 235-256Crossref PubMed Scopus (388) Google Scholar). GLUT-4 is expressed only in muscle and fat cells, where it resides in an intracellular compartment under basal conditions and is translocated to the cell surface after stimulation with insulin (15Olson A.L. Pessin J.E. Annu. Rev. Nutr. 1996; 16: 235-256Crossref PubMed Scopus (388) Google Scholar). Recently, it became clear that syntaxin 4 and several SNARE-related molecules participate in the translocation of GLUT-4 to the plasma membrane (16Thurmond D.C. Kanzaki M. Khan A.H. Pessin J.E. Mol. Cell. Biol. 2000; 20: 379-388Crossref PubMed Scopus (87) Google Scholar, 17Foster L.J. Klip A. Am. J. Physiol. Cell Physiol. 2000; 279: C877-C890Crossref PubMed Google Scholar). In addition, recent studies have revealed that glucose transport is regulated though several signal transduction pathways, including those that involve mitogen-activated protein kinase, phosphatidylinositol 3-kinase, and protein kinase C (PKC) (18Fujishiro M. Gotoh Y. Katagiri H. Sakoda H. Ogihara T. Anai M. Onishi Y. Ono H. Funaki M. Inukai K. Fukushima Y. Kikuchi M. Oka Y. Asano T. J. Biol. Chem. 2001; 276: 19800-19806Abstract Full Text Full Text PDF PubMed Scopus (109) Google Scholar, 19Martin S.S. Haruta T. Morris A.J. Klippel A. Williams L.T. Olefsky J.M. J. Biol. Chem. 1996; 271: 17605-17608Abstract Full Text Full Text PDF PubMed Scopus (215) Google Scholar, 20Tsuru M. Katagiri H. Asano T. Yamada T. Ohno S. Ogihara T. Oka Y. Am. J. Physiol. Endocrinol. Metab. 2002; 283: E338-E345Crossref PubMed Scopus (39) Google Scholar). We showed previously that astroglioma cells express syntaxin 1C but not syntaxin 1A, and that the expression of syntaxin 1C protein is up-regulated via a PKC signaling pathway by stimulating cells with phorbol 12-myristate 13-acetate (PMA) (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). In the present study, we used a human astroglioma cell line that expresses syntaxin 1C to determine whether syntaxin 1C is involved in glucose transport. We found that the induction of endogenous syntaxin 1C expression by PMA caused a reduction in GLUT-1 in the plasma membrane and suppressed glucose uptake. Expression of exogenous syntaxin 1C in T98G cells had the same effects. These results suggest that the physiological function of syntaxin 1C in astroglioma cells is the regulation of intracellular trafficking of GLUT-1. Reagents—All tissue culture reagents were purchased from Invitrogen, Life Technologies (Carlsbad, CA) with the exception of fetal calf serum (FCS), which was purchased from Sigma. Human insulin was purchased from Roche Applied Science (Basel, Switzerland). Acrylamide/bis-acrylamide was obtained from WAKO Chemical (Osaka, Japan). All other reagents were purchased from Sigma or Calbiochem (San Diego, CA), unless otherwise noted. DNA Cloning—Total cellular RNA was extracted using the QIA-Amp RNA extraction kit from Qiagen (Valencia, CA), according to the manufacturer's protocol. The reverse transcription (RT)-PCR was carried out using an RNA PCR kit (Takara, Tokyo, Japan), according to the manufacturer's protocol. To clone the coding region of human syntaxins, we designed oligonucleotide primers based on the sequence of human (h) syntaxin 1A and syntaxin 1C (DDBJ accession nos. D37932 and AB086954M, respectively), and syntaxin 4 (GenBank™ accession no. NM004604). The primers were used in RT-PCR cloning, using mRNA that was isolated from the human brain library (BIO101) and T98G human astroglioma cells, as described below. RT was performed using the oligo(dT) primer and AMV reverse transcriptase (Takara), according to the manufacturer's instructions. The full-length human syntaxin cDNAs were inserted into the BamHI/EcoRI site of a pcDNA3 expression vector (Invitrogen, Life Technologies), or cloned into the BamHI site of a pcDNA3 expression vector as an N-terminal 5× HA-tagged version. The subcloned syntaxins were confirmed by using an ABI 377 sequencer (Applied Biosystems, Foster City, CA). Semiquantitative PCR—GLUT gene expression was quantified according to the method of Schreiber et al. (21Schreiber J. Enderich J. Sock E. Schmidt C. Richter-Landsberg C. Wegner M. J. Biol. Chem. 1997; 272: 32286-32293Abstract Full Text Full Text PDF PubMed Scopus (53) Google Scholar). The cDNA template (5 ng) that was synthesized from total RNA in cells was used for semiquantitative PCR, with primer pairs that were specific for hGLUT-1, hGLUT-2, hGLUT-3, and hGLUT-4 (GenBank™ accession nos. K03195, J03810, M20681, and M20747, respectively). Human small intestine cDNA was a kind gift from Dr. Yoshikatsu Kanai (Kyorin University, Japan). The primer pairs were as follows: hGLUT-1 sense, 906-932 nt; hGLUT-1 antisense, 1519-1491 nt; hGLUT-2 sense, 2223-2243 nt; hGLUT-2 antisense, 2626-2607 nt; hGLUT-3 sense, 884-912 nt; hGLUT-3 antisense, 1375-1349 nt; hGLUT-4 sense, 1485-1513 nt; and hGLUT-4 antisense, 2076-2048 nt. As a control, we used a pair of primers for β-actin (Maxim Biotech, South San Francisco, CA) that amplifies a 540-bp DNA segment. The primers for β-actin span at least one intron, and contamination of RNA samples by genomic DNA can be detected according to the size of the amplified product (1116 bp for genomic DNA). PCR was linear up to 30 cycles for each pair of primers (data not shown). The intensity of the SYBR-green (Molecular Probes, Eugene, OR) signals from scanned images of the gels was measured using NIH Image (rsb.info.nih.gov/nih-image/). Northern Blot Analysis—Northern blot analysis was carried out, according to the method of Nagamatsu et al. (22Nagamatsu S. Nakamichi Y. Inoue N. Inoue M. Nishino H. Sawa H. Biochem. J. 1996; 319: 477-482Crossref PubMed Scopus (41) Google Scholar). Total RNA (20 μg) isolated from native and transfected (see below) T98G cells was separated by electrophoresis in 1.0% formaldehyde-agarose denaturing gels. The EcoRI-digested 600-bp fragments of the GLUT-1 and GLUT-3 cDNA were labeled with 32P by random priming. The GLUT-1 and GLUT-3 cDNAs were a kind gift from Dr. Shinya Nagamatsu (Kyorin University, Japan). The intensity of the autoradiographic signals was measured directly from digital images (Bas 2000, Fuji, Tokyo, Japan). Cell Culture and Transfection—Two human astroglioma cell lines, T98G and U87MG, were provided by Dr. Hiroki Sawa (Kyorin University, Japan). Cells were grown on 90-mm in diameter plastic dishes in Dulbecco's modified Eagle's medium (DMEM), supplemented with 10% (v/v) FCS, penicillin (100 μg/ml), and streptomycin (100 μg/ml). For drug stimulation, cells were treated for 3-48 h with 1-10 μm PMA (Sigma), 10 μm 4α-PMA (Sigma), or 10 μm forskolin (RBI). T98G and U87MG cells were trypsinized, washed twice with phosphate-buffered saline (PBS), and resuspended in 0.3 ml of serum-free DMEM on ice. Approximately 1 × 106 T98G cells were transfected with 150 μg/ml recombinant syntaxins by electroporation (Bio-Rad gene pulsar, 0.75 kV/cm field strength, 960 microfarad capacitance). The cells were then cultured in DMEM (that included 800 μg/ml neomycin) for 2 weeks, and the transfected cells were cloned as a single colony. Immunoblot Analysis—Equal amounts of protein from each sample were separated on 12% SDS-polyacrylamide gels as described previously (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). Following antibodies were used as a primary antibody: a monoclonal antibody (14D8) that had been raised against the N-terminal of syntaxin 1A (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar), a polyclonal anti-syntaxin 1C antibody (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar), a polyclonal anti-syntaxin 2 antibody (Stressgen Biotech, Victoria, BC, Canada), a polyclonal anti-syntaxin 3 antibody (Sigma), a monoclonal anti-syntaxin 4 antibody (BD-Transduction Laboratory, San Jose, CA), anti-GLUT-1 antiserum, anti-GLUT-3 antiserum (Chemicon, Temecula, CA), or anti-HA monoclonal antibody (3F10, Roche Applied Science). After washing, the membranes were incubated with horseradish peroxidase-conjugated anti-mouse IgG, anti-rabbit IgG (Cappel, Irvine, CA), or anti-rat IgG (Jackson Laboratories, Bar Harbor, ME). For GLUT immunoblotting, a total cell membrane preparation was made, as described previously (23Fladeby C. Bjonness B. Serck-Hanssen G. J. Cell. Physiol. 1996; 169: 242-247Crossref PubMed Scopus (17) Google Scholar). The cell surface biotinylation assay for GLUT-1 was carried out, according to the method of McMahon et al. (24McMahon R.J. Hwang J.B. Frost S.C. Biochem. Biophys. Res. Commun. 2000; 273: 859-864Crossref PubMed Scopus (8) Google Scholar). Quantitative analysis of the syntaxin immunoblots was carried out as described previously (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). After drug application, cells were treated with 10% trichloroacetic acid. After centrifugation, the precipitate was solubilized in 8 m urea, 1% SDS, 10 mm Tris-Cl (pH 7.5). The protein concentration was measured by using a DC-protein assay (Bio-Rad). The intensities of the immunoblotted signals were measured using NIH Image and normalized to that with anti-α-tubulin IgG (DM1A) (Sigma). Immunocytofluorescence—Immunostaining was carried out essentially as described previously (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). Briefly, to study GLUT-1 localization in cells treated with PMA or transfected with HA-tagged syntaxin, cells were fixed and permeabilized with acetone/methanol (1:1). After treatment with a blocking solution, the cells were then incubated with a monoclonal antibody (14D8) (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar) or an anti-HA monoclonal antibody (3F10) and an anti-GLUT-1 polyclonal antibody. After another wash with PBS, the cells were exposed to either anti-mouse IgG, anti-rat IgG coupled to Cy-3, or anti-rabbit IgG coupled to fluorescein isothiocyanate. The immunostained cells were examined using a confocal scanning laser microscope (Zeiss LSM 410, Jena, Germany) that was equipped with a triple band-pass filter set. Cell Growth and Cell Cycle Analysis—T98G cells that had been starved of serum for 24 h were stimulated with DMEM containing 10% FCS for 0-96 h. Control and syntaxin-expressing cells were seeded in 90-mm in diameter culture dishes (5 × 104 cells/dish). The number of living cells was counted up to 96 h following DMEM/FCS treatment. The growth rate in the logarithmic growth phase (48-96 h) was calculated for each cell line. The cell cycle analysis was carried out as reported previously (25Smit M.J. Verzijl D. Iyengar R. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 15084-15089Crossref PubMed Scopus (24) Google Scholar). Growth of T98G cells that had been plated at a density of 2 × 105 cells per 90-mm in diameter culture dish was arrested by removing serum for 24 h. The cells were restimulated for 40 h with medium containing 10% serum. Thereafter, the cells (1 × 106 cells/ml) treated with 0.5 mg/ml RNase (Nippon Gene, Tokyo, Japan) were analyzed immediately after propidium iodide (Sigma) staining using fluorescence-activated cell sorting (FACS, BD Biosciences Epics Profile II) with argon laser excitation (488 nm) and a 588-nm (FL2) emission filter. At least 10,000 cells were collected for each sample (excluding the gated cells). The percentage of cells in the G0/G1, S, and G2/M phase was estimated from FL2-height histograms using ModFit (Verity Software, Topsham, ME). Measurement of Glucose Uptake—Glucose transport was assayed by measuring the uptake of 2-deoxy-[3H]glucose (2-DG), essentially as described previously (22Nagamatsu S. Nakamichi Y. Inoue N. Inoue M. Nishino H. Sawa H. Biochem. J. 1996; 319: 477-482Crossref PubMed Scopus (41) Google Scholar), but with slight modifications. The uptake assay was carried out 48-72 h after cell passaging. Cells (1 × l05) in Hanks' balanced salt solution (HBSS), containing 0.03 g/100 ml bovine serum albumin, 136.9 mm NaCl, 5.6 mm KC1, 0.34 mm NaHPO4, 0.44 mm KH2PO4, 1.27 mm CaCl2, and 4.20 mm NaHCO3,20mm HEPES, pH 7.4, were incubated on 12-well multiplates at 37 °C, for 30 min. Glucose uptake was initiated by adding 0.5 μCi of 2-deoxy-d-[1,2-3H(N)]glucose (2-[3H]DG; PerkinElmer Life Sciences) to 0.5 ml of HBSS buffer, in the presence of 0.1 mm 2-deoxy-d-glucose, in 35-mm in diameter wells. After 13 min at room temperature, uptake was terminated by rapid washing with 1 ml of ice-cold PBS. The uptake of 2-DG was linear between 0 and 20 min of incubation (data not shown). For the kinetic analysis, we used 0.1-100.0 mm 2-DG (0.0064-6.4 μm 2-[3H]DG). The cells were solubilized in 1% SDS, and the amount of radioactivity was measured. Protein content was measured using the DC protein assay from Bio-Rad. The difference in the amount of uptake in the presence and absence of 0.5 mm cytochalasin B (a transport inhibitor) was calculated; this represented glucose transporter-dependent activity. In each experiment, glucose uptake was assayed in triplicate. We studied the kinetics of glucose uptake using different concentrations of d-glucose, as described previously (26Muona P. Sollberg S. Peltonen J. Uitto J. Diabetes. 1992; 41: 1587-1596Crossref PubMed Scopus (42) Google Scholar). Briefly, confluent cultures of T98G cells were incubated with either 5.5 or 25.0 mm d-glucose for 7 days. The culture medium was changed daily to maintain a relatively constant concentration of glucose. To study basal glucose uptake via sodium-dependent glucose transporters (SGLTs), cells were treated with Na+-free HBSS buffer containing 0.03 g/100 ml bovine serum albumin, 138 mm N-methyl-d-(-)glucamine (NMDG), 5.6 mm KC1, 0.34 mm KHPO4, 0.44 mm KH2PO4, 1.27 mm CaCl2, and 20 mm HEPES (pH 7.4). Statistical Analysis—Data are expressed as mean ± S.E. and were analyzed using one-way analysis of variance. A p value of ≤ 0.05 was considered to be statistically significant. Measurement of 2-DG Uptake via GLUTs and SGLTs in T98G Cells—To examine the relationship between syntaxin 1C expression and glucose transport, we first determined whether there was glucose uptake via GLUT in T98G cells. The amount of 2-DG uptake via GLUTs and SGLTs was measured in the presence of cytocharasin B (an antagonist of GLUTs) and in Na+-free medium. GLUT activity accounted for ∼80% of total 2-DG uptake in T98G cells. By contrast, SGLT activity accounted for less than 20% of total 2-DG uptake (data not shown). Uptake of 2-DG was linear for up to 20 min of incubation (data not shown). These observations indicate that GLUTs are expressed in T98G cells. Identification of GLUT Isoform in T98G Cells—There are several reports that the GLUT isoforms GLUT-1 and GLUT-3 are the main components of several types of glioma (22Nagamatsu S. Nakamichi Y. Inoue N. Inoue M. Nishino H. Sawa H. Biochem. J. 1996; 319: 477-482Crossref PubMed Scopus (41) Google Scholar, 27Nagamatsu S. Sawa H. Wakizaka A. Hoshino T. J. Neurochem. 1993; 61: 2048-2053Crossref PubMed Scopus (79) Google Scholar, 28Boado R.J. Black K.L. Pardridge W.M. Brain Res. Mol. Brain Res. 1994; 27: 51-57Crossref PubMed Scopus (141) Google Scholar). However, whether GLUTs are expressed in T98G cells has not been determined. To determine which GLUT isoform(s) is expressed in T98G cells, we investigated the kinetics of 2-DG uptake. As shown in Fig. 1A, the Km of 2-DG uptake in T98G cells was 2.42 ± 0.12 mM. GLUT-1, GLUT-3, and GLUT-4 are high affinity glucose transporters, whereas GLUT-2 is a low affinity transporter (29Walmsley A.R. Barrett M.P. Bringaud F. Gould G.W. Trends Biochem. Sci. 1998; 23: 476-481Abstract Full Text Full Text PDF PubMed Scopus (99) Google Scholar). Because the value of Km in the present study is far smaller than that of GLUT-2 (Km: 20-40 mM), it is likely that the GLUT isoform functioning in T98G cells is not GLUT-2, but rather a high affinity transporter, i.e. GLUT-1, GLUT-3, or GLUT-4. As shown in Fig. 1B, semiquantitative RT-PCR revealed that GLUT-1 and GLUT-3 were expressed in T98G cells; GLUT-2 and GLUT-4 expression was undetectable except under saturated PCR conditions (data not shown). To examine the expression levels of endogenous GLUT-1 and GLUT-3, we studied expression of the mRNA of these GLUT isoforms in T98G cells by Northern blot analysis. As shown in Fig. 1C, GLUT-1 mRNA was more abundant than that of GLUT-3. We also confirmed expression of GLUT-1 protein by immunoblotting (see Fig. 3C) and localization in plasma membrane in T98G cells by cell surface biotinylation assay (data not shown). It has been reported that glucose uptake via GLUT-1 in cells cultured in low glucose medium is higher than in the presence of high concentrations of glucose (26Muona P. Sollberg S. Peltonen J. Uitto J. Diabetes. 1992; 41: 1587-1596Crossref PubMed Scopus (42) Google Scholar). We investigated glucose uptake in T98G cells that were cultured with different concentrations of glucose. As expected, 2-DG uptake was ∼1.5 times greater in low-glucose medium (5.5 mM glucose), compared with high glucose medium (25 mM glucose) (Fig. 1D). Another property of GLUT-4 is that it translocates to the plasma membrane in cells that have been stimulated with insulin, which results in an increase in glucose uptake (16Thurmond D.C. Kanzaki M. Khan A.H. Pessin J.E. Mol. Cell. Biol. 2000; 20: 379-388Crossref PubMed Scopus (87) Google Scholar). Consequently, we tested whether the amount of 2-DG uptake in T98G cells would increase after stimulation with insulin; this was not the case (Fig. 1D). The results shown in Fig. 1, B and D suggest that there is no functional GLUT-4 in T98G cells. The aforementioned results demonstrate that the major isoform of GLUT in T98G cells is GLUT-1. Activation of Endogenous Syntaxin 1C by PMA Suppresses Translocation of GLUT-1 to the Plasma Membrane in T98G Cells—Our previous study revealed that T98G astroglioma cells express syntaxin 1C, but not syntaxin 1A, and that syntaxin 1C expression can be activated by PMA (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar). In the present study, we investigated whether a change in the level of syntaxin 1C expression might affect glucose transport. Fig. 2, A and B shows the change in syntaxin 1C expression and 2-DG uptake in T98G cells that were treated with either PMA, forskolin, or 4α-PMA (a nonfunctional analog of PMA). Uptake of 2-DG in PMA-treated cells was reduced by ∼85%, compared with control cells (Fig. 2B), whereas 2-DG uptake was not affected by either forskolin or 4α-PMA (Fig. 2B). None of the aforementioned treatments had any effect on the uptake of 2-DG via SGLTs (data not shown). In addition, an analysis of the time course of glucose uptake (Fig. 2, C and D) revealed that 2-DG uptake in T98G cells was reduced as the PMA-induced level of syntaxin 1C expression increased. Because syntaxin 1A mRNA is not found in astroglioma cells, irrespective of whether cells are treated with PMA (9Nakayama T. Mikoshiba K. Yamamori T. Akagawa K. FEBS Lett. 2002; 536: 209-214Crossref Scopus (6) Google Scholar), the observed change in glucose uptake in PMA-treated T98G cells was not caused by the actions of syntaxin 1A. To determine whether the reduction in glucose uptake was caused by the presence of syntaxin 1C, we examined the expression of syntaxin 2, syntaxin 3, and syntaxin 4 in the plasma membrane. In contrast to the expression of syntaxin 1C, which is increased in a dose-dependent manner by PMA treatment (7.37 ± 1.34-fold increase in response to 10 μm PMA), the expression of syntaxin 2, syntaxin 3, and syntaxin 4 was not affected by PMA (Fig. 3A). Furthermore, PMA had no effect on the expression of GLUT-1 and GLUT-3 mRNA and protein in T98G cells (Fig. 3, B and C). These results suggest that the treatment of T98G cells with PMA did not alter the level of expression of GLUT-1, GLUT-3, or syntaxins other than syntaxin 1C. We also analyzed the effect of PMA using immunofluorescence and found that in cells in which the expression of endogenous syntaxin 1C had been enhanced by PMA, GLUT-1 expression in the plasma membrane decreased, whereas expression in the intracellular fraction increased (Fig. 4, B and F). By contrast, neither forskolin (Fig. 4, D and H) nor 4α-PMA (Fig. 4, C and G) had any effect on GLUT-1 expression. Glucose Uptake via GLUT-1 in T98G Cells Transfected with Syntaxin 1A, Syntaxin 1C, or Syntaxin 4—To determine whether syntaxin 1C expression affects glucose transport in astroglioma cells, we introduced exogenous syntaxin 1A, syntaxin 1C, or syntaxin 4 tagged with HA into at" @default.
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• W2085125612 date "2004-05-01" @default.
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• W2085125612 title "Activation of Syntaxin 1C, an Alternative Splice Variant of HPC-1/Syntaxin 1A, by Phorbol 12-Myristate 13-Acetate (PMA) Suppresses Glucose Transport into Astroglioma Cells via the Glucose Transporter-1 (GLUT-1)" @default.
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• W2085125612 doi "https://doi.org/10.1074/jbc.m314297200" @default.
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