help button home button Endocrine Society Endocrinology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Samih, N.
Right arrow Articles by Fayet, G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Samih, N.
Right arrow Articles by Fayet, G.
Endocrinology Vol. 141, No. 11 4146-4155
Copyright © 2000 by The Endocrine Society

ARTICLES

Glut-1 Translocation in FRTL-5 Thyroid Cells: Role of Phosphatidylinositol 3-Kinase and N-Glycosylation

Nezha Samih, Sonia Hovsepian, Azedine Aouani, Dominique Lombardo and Guy Fayet

INSERM Unité 260, Faculté de Médecine, Université de la Méditerranée,13385 Marseille Cedex 5, France

Address all correspondence and requests for reprints to: Dr. Guy Fayet, INSERM Unité 260, Faculté de Médecine, 27 Boulevard Jean Moulin, 13385 Marseille Cedex 5, France. E-mail: U260{at}medecine.univ-mrs.fr


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
It was previously demonstrated that insulin or TSH treatment of FRTL-5 cells resulted in an elevation of glucose transport and in an increase of cell surface expression of the glucose transporter Glut-1. However, the signaling mechanisms leading to the insulin or TSH-induced increase in the cell surface expression of Glut-1 were not investigated. In the present study, we demonstrated that wortmannin and LY294002, two specific inhibitors of phosphatidylinositol 3-kinase (PI3-kinase), interfere both in the signaling pathways of insulin and TSH leading to glucose consumption enhancement and Glut-1 translocation. Two hours after insulin treatment, TSH or cAMP analog (Bu)2cAMP stimulation, glucose transport was increased and most of the intracellular Glut-1 pool was translocated to plasma membranes. Wortmannin or LY294002 blocked the insulin, (Bu)2cAMP, and the TSH-induced translocation of Glut-1. Wortmannin or LY294002 alone did not alter the basal ratio between intracellular and cell surface Glut-1 molecules. These results suggest that in FRTL-5 cells wortmannin and LY294002 inhibited the insulin, (Bu)2cAMP and TSH events leading to Glut-1 translocation from an intracellular compartment to the plasma membrane. Likewise, (Bu)2cAMP effects on glucose transport and Glut-1 translocation to plasma membrane were repressed by PI3-kinase inhibitors but not by the protein kinase A (PKA) inhibitor H89. We suggest that (Bu)2cAMP stimulates Glut-1 translocation to plasma membrane through PI3-kinase-dependent and PKA-independent signaling pathways. To further elucidate mechanisms that regulate the translocation of Glut-1 to cell membrane, we extended this study to the role played by the N-glycosylation in the translocation and in the biological activity of Glut-1 in FRTL-5 cells. For this purpose we used tunicamycin, an inhibitor of the N-glycosylation. Our experiments with tunicamycin clearly showed that both the glycosylated and unglycosylated forms of the transporter reached the cell surface. Likewise, a decrease in glucose consumption (-50%) after treatment of cells with tunicamycin was accompanied by a decrease (-70% vs. control) in the membrane expression of a 50-kDa form of Glut-1 and an increase in its unglycosylated 41-kDa form. These results suggest that carbohydrate moiety is essential for the biological activity of glucose transport but is not required for the translocation of Glut-1 from the intracellular membrane pool to the plasma membrane.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
FACILITATED glucose transport across plasma membranes is mediated by a set of homologous glycoprotein molecules (Gluts) that are expressed in a tissue-specific manner (1). These carriers are energy- and sodium-independent and differ from the SGLT cotransporter family (2). Glut-1 is widely expressed in normal tissues and highly represented in erythrocytes and brain, Glut-2 in liver and pancreatic cells, Glut-4 in muscle and adipose tissues. Their expression and function are regulated by intracellular and extracellular signals such as growth factors (3), death-inducing agents (4), or extracellular stress (5). In insulin-responsive cells, insulin acutely stimulates glucose transport by translocating Glut-1 or Glut-4 from the intracellular pool to the plasma membrane (6) rather than stimulating preexist Glut-s molecules from the plasma membrane. In addition to this recruitment, insulin modifies the intrinsic activity of both Glut-1 and Glut-4 (7).

In FRTL-5 cells, a rat thyroid cell line used as a model system, it was shown previously that insulin (8) or TSH (9) exert a large stimulation of glucose transport. This transport occurs through the Glut-1 isoform. Under both basal and stimulated conditions, the stimulation of glucose consumption was explained by an increased translocation of Glut-1 toward the FRTL-5 cell surface (10, 11).

However, this translocation process has not yet been studied in this thyroid system in its mechanistic aspects. A putative candidate as a signaling intermediate between the insulin activated receptor and the translocation of Glut-1 is the enzyme phosphatidylinositol 3-kinase (PI3-kinase). Phosphatidylinositides are phosphorylated in position 3 by PI3-kinase leading to phosphatidylinositol phosphates (12). These D-3 phosphorylated inositides may be involved in the control of cell growth and metabolism (12). PI3-kinase is a heterodimer consisting in the association of 110-kDa (p110) and 85-kDa (p85) subunits. Complementary DNA cloning of these proteins revealed that p110 is a catalytic subunit (13) and p85 an adaptor subunit containing two src homologous region 2 (SH2) and one SH3 domain (14). The SH2 domains are involved in interactions with tyrosine-phosphorylated proteins (15). Insulin and IGF-1 increase PI3-kinase activity (16, 17).

To investigate the possible implication of PI3-kinase in insulin or TSH-stimulatory processes, we used wortmannin and LY294002, two specific inhibitors of this enzyme. Wortmannin, which binds to the catalytic p110 but not to the p85 regulatory subunit, blocks PI3-kinase activity both in vivo and in vitro (18, 19), and does not act on the tyrosine phosphorylation of the insulin and IGF-1 receptors and on IRS-1 (20). LY294002, another inhibitor of PI3-kinase is structurally distinct from wortmannin (21). The aim of this study being to understand events that may regulate the translocation of Glut-1, we extended this study by investigating the role of the N-glycosylation in the translocation and biological activity of glucose transporters, which is still unclear (22, 23). The intrinsic activity of Glut-1 can be modulated by changes in the glycosylation state of the protein (24, 25). Glut-1 contains a single potential N-glycosylation site at Asn45, between the membrane regions M1 and M2 (23).

Using cell surface biotinylation followed by isolation of cell surface proteins and quantitative estimation of Glut-1 sites by Western blotting, we demonstrated in this paper that the wortmannin and LY294002 inhibitory effect on glucose consumption or glucose transport is associated with an impairment of the Glut-1 translocation process. This was performed during the early phase of the stimulation of FRTL-5 cells with insulin, (Bu)2cAMP or TSH. Secondly, we also demonstrated that tunicamycin, which prevents N-linked glycosylation of Glut-1, is associated with a reduced glucose consumption but does not affect the translocation of the transporter toward the plasma membrane.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell cultures
FRTL-5 cells (Fisher rat thyroid) were grown in Coon’s modified Ham’s F12 medium containing 12.2 mM glucose, 5% FCS and the six components (6H) proposed by Ambesi-Impiombato et al. (26) i.e. 1 mU/ml TSH, 5 µg/ml insulin, 5 µg/ml transferrin, 10 nM hydrocortisone, 10 ng/ml somatostatin and 10 ng/ml glycyl-L-histidyl-L-lysine acetate. Cells were cultured in 100 mm Petri dishes or 24-well tissue culture clusters at 37 C under air-CO2 (95–5%) in a water-saturated incubator. When confluence was reached, the cells were washed twice with Ca2+ and Mg2+ -free Puck F and then maintained for 48–72 h in Coon’s modified Ham’s F12 medium supplemented with transferrin and 0.2% FCS or 0.2% BSA. The cells were then incubated for varying time periods with the additives as indicated when required. We also used in this study porcine thyroid cells from primary cultures and two porcine thyroid cell lines expressing different glucose consumption: the Porthos cells, which are permanently growing normal porcine thyroid cells deposited at the European Collection of Animal Cell Cultures (Salisbury) under the accession number 93 12 23 23 and the Athos cells (27), which are a subline deriving from the previous one characterized by a high glucose consumption (ECACC 93 12 23 25). Primary cultured cells were maintained in Click-RPMI medium (8.3 mM glucose) with 1% newborn calf serum and 5 µg/ml insulin. Porthos and Athos cells were routinely grown in the same medium with 6H and 5H (6H without TSH), respectively. Media in all cases were changed 24–48 h before initiation of studies (see Results). Cells were washed twice with Ca2+ and Mg2+-free Puck F and then maintained for varying periods with the additives as indicated.

Glucose consumption
Glucose consumption was calculated by comparing the glucose concentration in cell culture media at time 0 with the glucose concentration measured after required incubation times. Glucose was measured by the glucose oxidase method, using a glucose analyzer Hitachi 717.

Glucose uptake
FRTL-5 cells were grown to confluence in 24-well dishes. These cells were washed twice in Coon’s modified Ham’s F12 medium and maintained for 48–72 h in this medium containing 0.2% BSA. The cells were washed twice in Krebs-Ringer-HEPES (20 mM) pH 7.4, and a glucose-free incubation was done in Krebs-Ringer-HEPES. Cells were incubated with LY294002 or H89 for 30 min (Sigma, St. Louis, MO) and then treated with 1 mU/ml TSH or 1 mM (Bu)2cAMP for 2 h. During the last 10 min of TSH or (Bu)2cAMP stimulation, glucose uptake was initiated by the addition of 0.1 mM [3H] 2-deoxy-D-glucose (2-DOG) (1 µCi/ml; 21 Ci/mmol) (NEN Life Science Products, Boston, MA) and 100 µM unlabeled 2-DOG. Nonspecific uptake was determined in the presence of 20 µM cytochalasin B (9, 28).

DNA measurement
DNA was determined by the fluorometric assay according to Labarca and Païgen (29) using calf thymus DNA as standard.

Biotinylation of surface proteins
Surface biotinylation of FRTL-5 cells was adapted from Shetty et al. (30) and Fayadat et al. (31) with minor modifications. Confluent cells on 100 mm plates were preincubated 30 min with 1 µM wortmannin, 20 µM LY294002, 25 µM H89 (Sigma), or DMSO (vector). These cells were then treated for 2 h at 37 C in the same medium either with or without 5 µg/ml insulin (Ins), 1 mU/ml TSH or 1 mM (Bu)2cAMP washed twice with ice-cold PBS and incubated with 0.25 mg/ml N-hydroxysuccinimide-long-chain-biotin (NHS-LC-biotin (Pierce Chemical Co., Madison, WI) in PBS for 30 min at 4 C. The reaction was stopped by rinsing the plates three times with 15 mM glycine in ice-cold PBS. Cells were then scraped on ice in PBS and centrifuged at 200 x g for 5 min. Cell pellets were solubilized for 30 min on ice in solubilization buffer (0.15 M NaCl, 50 mM HEPES pH 7.1, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 10 mM EDTA, 2 mM PMSF, 2 µg/ml aprotinin, 0.1 mM antipain, 0.2 mM leupeptin, and 0.5 U/ml {alpha}-macroglobulin). The supernatant was separated by centrifugation for 3 min at 9,000 x g and mixed with 50 µl streptavidin-agarose beads (Sigma) that had been sedimented following a preequilibration in the solubilization buffer. The suspension was gently mixed overnight at 4 C, and beads pelleted by centrifugation. The bead pellet was washed five times with 1 ml of 0.15 M NaCl, 10 mM Tris-HCl pH 7 containing 2 mM PMSF and 0.2 mM leupeptin, and once again with PBS. The final pellet was resuspended in 120 µl of Laemmli’s buffer (32) (1.2-fold concentrated without mercaptoethanol and bromophenol blue) and incubated for 30 min at 65 C. The supernatant containing solubilized surface proteins was separated from the beads by centrifugation, collected and kept at -80 C until use. Protein content was determined by the microBCA protein assay kit (Pierce Chemical Co.) using BSA as a standard and dissolved in the same buffer as the intracellular (IC) and surface (S) proteins preparation.

PAGE and Western blot analysis
Confluent monolayers were rinsed with PBS, scraped on ice in PBS, and centrifuged at 200 x g. Cell pellets were resuspended in solubilization buffer for 30 min on ice. After centrifugation (9,000 x g, 3 min), proteins extracted (supernatant) were separated on a 8% or 10% SDS-PAGE according to Laemmli (32) and electroblotted on nitrocellulose membrane. Blots were incubated overnight at 4 C with rabbit antibodies directed against Glut-1 (Chemicon). These latter antibodies were raised against the synthetic peptide corresponding to the 13 amino acids of the C-terminal region of the rat brain Glut-1 (33). Blots were then developed by Enhanced Chemiluminescence using peroxidase-conjugated antibodies against rabbit IgG. The intensity of bands was determined by densitometric scanning (HP Scan Jet 5100 C or Quato X Finity).

Digestion of glycoproteins with glycopeptidase F (PNGase F)
Protein-samples (20–30 µg) were suspended in digestion buffer (30 µl, 2.6% Nonidet P-40, 40 mM Tris-HCl pH 8, 5 mM O-phenanthroline and 33 mM ß-mercaptoethanol) and incubated with Glycopeptidase F (0.6U) at 37 C for 18 h (34). These digested samples were resolved on SDS-PAGE and visualized by immunoblotting using anti-Glut-1 antibodies.

Statistical analysis
Data were analyzed statistically using Student’s t test and considered significant when P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Glucose consumption and Glut-1 expression in thyroid cells
To understand the possible relationship between Glut-1 expression and glucose consumption, we studied these two parameters in thyroid cells from different species. Fisher Rat Thyroid FRTL-5 cells, porcine primary cell culture, Athos and Porthos cells lines were cultured for 24 h in Click-RPMI medium containing 6H and 5% FCS. FRTL-5 cells, Athos cells and porcine primary cultured cells consumed 3–4 more glucose than Porthos cells (Fig. 1AGo). Protein extracts corresponding to 5 µg cell DNA were analyzed on SDS-PAGE and by immunoblotting (Fig. 1CGo). A 50-kDa Glut-1 protein was detected in porcine cells from primary cultures and FRTL-5 cells instead of a 54-kDa Glut-1 protein in Athos and Porthos cells. Glut-1 expression was comparable (Fig. 1BGo) in FRTL-5, Athos and primary cultured porcine thyroid cells but was 3–4 times lower in Porthos cells.



View larger version (24K):
[in this window]
[in a new window]
  		Figure 1. Glucose consumption and immunoblot analysis of Glut-1 in thyroid cells from two species (porcine and rat). Cultured thyroid cells from porcine and rat were maintained in their respective culture media. At confluence the cells were washed twice with Ca2+ and Mg2+-free Puck F and then treated for 24 h in Click-RPMI medium supplemented with 5% FCS and 6H. A, Glucose consumption in media from primary cultured porcine cells, porcine cell lines (Athos and Porthos) and FRTL-5 cells. Statistically significant differences between Porcine primary culture, Athos or FRTL-5 cells vs. Porthos cells (*), P < 0.05; (**), P < 0.01, (***), P < 0.001; Porcine primary culture vs. Athos cells, not significant. B, Quantification was performed by scanning the band corresponding to Glut-1. Statistically significant differences between Porcine primary culture, Athos or FRTL-5 cells vs. Porthos cells (*), P < 0.05; (**), P < 0.01, (***), P < 0.001. C, Cell lysates (5 µg DNA/lane) were separated on 8% SDS-PAGE as described in Materials and Methods. D, N-glycosidase F digestion of Glut-1 in thyroid cell extracts. Cell lysates (5–25 µg protein/lane) from primary cultured porcine cells, porcine thyroid cell lines (Athos and Porthos) and FRTL-5 cells were incubated without or with N-glycosidase F, followed by the detection of Glut-1 by immunoblotting. Each point in A and B is the mean ± SD of at least three independent experiments, C and D are representative of three experiments.

 
The above-observed difference in Glut-1 electrophoretic migration was likely due to variation in carbohydrate moieties as illustrated in Fig. 1DGo. Indeed, when cell lysates were treated with PNGase F, which cleaves N-linked oligosaccharides of glycoproteins, either the 50- and 54-kDa forms of Glut-1 were converted into the same 41-kDa product, which likely represents the unglycosylated form of the glucose transporter (35).

Effects of insulin and TSH on glucose consumption
The effects of insulin and TSH on glucose consumption in FRTL-5 cells were measured in Coon’s cell culture media containing transferrin and 0.2% FCS. After a treatment for 24 h with insulin, glucose consumption increased in a concentration-dependent manner reaching a maximal stimulation at 1 µg/ml insulin (Fig. 2AGo). Under similar conditions, TSH effect was also concentration dependent with a maximal stimulation observed at 1 mU/ml TSH (Fig. 2BGo). These results demonstrate that insulin and TSH increased the glucose consumption of FRTL-5 cells.



View larger version (17K):
[in this window]
[in a new window]
  		Figure 2. Glucose consumption in FRTL-5 cells vs. insulin (A) or TSH (B) concentrations, measured after 24 h. Confluent cells on 24-well tissue culture clusters were maintained for 48 h in Coon’s modified Ham’s F12 medium containing transferrin and 0.2% FCS and were treated then for 24 h in the presence of insulin (Ins) or TSH at the final concentrations as indicated. Glucose consumption was determined by comparing the glucose concentration in cell culture media at the beginning of the incubation with that measured at the end of the experiment. Each point is the mean ± SD. The experiments were repeated at least three times with triplicate determinations within one experiment.

 
Effects of insulin and TSH on glucose consumption, glucose transport and Glut-1 expression
To determine whether these effects were related to an increase in the expression of the Glut-1 protein, Western blot analyses were performed with FRTL-5 cell extracts after stimulation with insulin or TSH for 2 and 24 h. Figure 3AGo indicates that insulin (5 µg/ml) increased glucose consumption at 2 h (x1.66 ± 0.36 over control, P < 0.001), whereas Glut-1 protein in the whole cell homogenate did not change (Fig. 3Go, B and C). In contrast after 24 h incubation, insulin increased both glucose consumption (Fig. 3AGo) (x 3.60 ± 0.37 over control, P < 0.001) and Glut-1 expression (Fig. 3Go, B and C) (x 1.64 ± 0.01, P < 0.001). Similar results were obtained with TSH (1 mU/ml), which also increased glucose consumption and glucose transport at 2 h (x 1.81 ± 0.09, P < 0.01; x 2.8 ± 0.006, P < 0.001 over control, respectively) (Figs. 3AGo and 4AGo) without modification in Glut-1 level (Fig. 3Go, B and C). (Bu)2cAMP (1 mM) also increased glucose transport (x 2.96 ± 0.02 over control, P < 0.001, Fig. 4BGo). After 24 h, TSH increased both glucose consumption (x 3.70 ± 1.14 over control) and Glut-1 protein (x 1.91 ± 0.02, P < 0.001). Although cycloheximide alone decreased by some 21% Glut-1 expression compared with control, when cells were treated for 24 h with this inhibitor of protein synthesis either with insulin (5 µg/ml) or TSH (1 mU/ml), the cellular expression of Glut-1 was unchanged compared with control (Fig. 3DGo). This last result suggests that the increase in Glut-1 expression in FRTL-5 cells after 24 h stimulation with or without insulin or TSH (Fig. 3Go, C and D) was due to a de novo synthesis of Glut-1.



View larger version (25K):
[in this window]
[in a new window]
  		Figure 3. Effects of insulin or TSH on glucose consumption and Glut-1 expression in FRTL-5 cells vs. time. Confluent cells maintained for 48 h in Coon’s modified Ham’s F12 medium containing transferrin and 0.2% FCS were treated then for 2 and 24 h in the absence (control) or in the presence of 5 µg/ml insulin or 1 mU/ml TSH. A, Glucose consumption. Control and insulin-stimulated or TSH- stimulated states for 2 h differ significantly (*), P < 0.05, (**), P < 0.01, (***), P < 0.001; Control and insulin-stimulated or TSH-stimulated states for 24 h differ significantly (**), P < 0.01, (***), P < 0.001. B, Quantification was performed by scanning the band corresponding to Glut-1. Control and insulin-stimulated or TSH- stimulated levels for 24 h differ significantly (**), P < 0.01, (***), P < 0.001. C, Western blot analysis of Glut-1 protein with or without 5 µg/ml insulin or 1 mU/ml TSH at 2 and 24 h. D, Cells were treated with or without 10 µg/ml cycloheximide (CH) for 24 h and with or without 5 µg/ml insulin or 1 mU/ml TSH. Cell lysates (5 µg DNA/lane) were separated on 8% SDS-PAGE gel as described in Materials and Methods. Each point in A and B is the mean ± SD of at least three independent experiments, C and D are representative of three experiments.

 


View larger version (22K):
[in this window]
[in a new window]
  		Figure 4. Inhibition of glucose uptake by LY294002. Confluent cells were deprived of serum and glucose and preincubated in Krebs-Ringer-HEPES containing 0.2% BSA with the indicated concentrations of LY294002 (A), in the presence (+) or absence (-) of TSH. Statistically significant differences vs. 0 (*), P < 0.05; (**), P < 0.01, (***), P < 0.001. B, cells were incubated with LY294002 (20 µM) for 30 min, followed by incubation in the presence (+) or absence (-) of H89 (25 µM) or (Bu)2cAMP (1 mM) for 2 h. Statistically significant differences vs. (Bu)2cAMP (*), P < 0.05; (**), P < 0.01, (***), P < 0.001. Glucose uptake was determined during the last 10 min of TSH or (Bu)2cAMP treatment by the addition of 0.1 mM [3H]-2-deoxy-D-glucose. Each point represents the mean ± SD of three experiments.

 
Effect of PI3-kinase inhibitors on glucose consumption, glucose transport and Glut-1 expression
PI3-kinases have been found to regulate various steps in receptor-dependent endocytic trafficking. Therefore, we attempted to determine whether PI3-kinases were implicated in the cycling of Glut-1 expressed after stimulation with insulin or TSH and (Bu)2cAMP. The effects of wortmannin and LY294002, two PI3-kinase inhibitors, were examined. In a first series of experiments, various concentrations of wortmannin were used in combination with optimized concentration of insulin or TSH, 5 µg/ml and 1 mU/ml, respectively. We firstly recorded the glucose consumption after 24-h stimulation. As indicated in Fig. 5AGo for insulin-stimulated FRTL-5 cells and in Fig. 5BGo for TSH-stimulated cells, 1 µM wortmannin inhibited the hormonal-stimulated glucose consumption (x 0.41 ± 0.05, P < 0.001; x 0.48 ± 0.045, P < 0.001 for insulin and TSH, respectively). The effect of wortmannin appeared more specific in hormone-stimulated conditions compared with controls where wortmannin did not inhibit the basal glucose consumption. These results were confirmed using LY294002 and similar effects were obtained with 20 µM LY294002 in FRTL-5 cells stimulated by TSH or (Bu)2cAMP (x 0.41 ± 0.01, P < 0.001; x 0.43 ± 0.02, P < 0.001, respectively) (Fig. 4Go, A and B).



View larger version (18K):
[in this window]
[in a new window]
  		Figure 5. Effects of wortmannin on glucose consumption in FRTL-5 cells after 24 h. FRTL-5 cells were incubated in Coon’s modified Ham’s F12 medium containing transferrin and 0.2% FCS without [control, closed circles (A) and closed squares (B)] or with 5 µg/ml of insulin (open circles) (A) or 1mU/ml of TSH (open squares) (B) in the presence of various concentrations of wortmannin as indicated. Each point is the mean ± SD. The experiments were repeated three times with triplicate determinations within one experiment.

 
To understand the partition of Glut-1 molecules between the intracellular pool and the plasma membrane domain, we developed the biotinylation of cell surface protein. After biotinylation, proteins from the two domains were separated with streptavidin-agarose, analyzed by SDS-PAGE, and transferred to membranes (30, 31). When the blots were probed with anti Glut-1 antibody, we observed that the conjugation with biotin affected a little the electrophoretic migration of cell-surface Glut-1 (52–50 kDa) compared with nonbiotinylated Glut-1 from the intracellular pool (Fig. 6Go). We next examined the effect of wortmannin on Glut-1 expression and its partition i.e. the intracellular pool (IC) vs. cell surface (S) of FRTL-5 cells treated either with wortmannin (1 µM), insulin (5 µg/ml) or insulin + wortmannin after 2 h stimulation (Fig. 6AGo). Clearly, data indicated that insulin promoted the translocation of Glut-1 from the intracellular pool to cell surface (lane 6 compared with 5). Insulin increased Glut-1 level at the surface (x 2.39 ± 0.16, P < 0.001, lane 6 compared with 2). Under identical condition TSH (1 mU/ml) or (Bu)2cAMP (1 mM) have similar effects (Fig. 6Go, B and C) and induced the translocation of Glut-1 from the intracellular pool to the cell surface (Fig. 6BGo, lane 6 compared with 5; Fig. 6CGo, lane 7 compared with 8 and lane 3 compared with 4). TSH also increased Glut-1 level at the cell surface compared with control (x 2.76 ± 0.06, P < 0.001, lane 6 compared with 2). Indeed, under stimulation conditions most of the expressed Glut-1 was translocated toward the cell surface. Wortmannin or LY294002 alone have no significant effects compared with controls (lane 3 compared with 1, and lane 4 compared with 2; lane 12 compared with 10, and lane 11 compared with 9, respectively). However, used in conjunction with insulin or TSH, wortmannin inhibited the hormone-induced translocation of Glut-1 toward plasma membranes (x 0.3 ± 0.06, P < 0.001; x 0.5 ± 0.01, P < 0.001 for insulin and TSH, respectively, Fig. 6AGo and 6BGo, lane 6 compared with 8). Similar results were obtained with LY294002 in the presence of TSH (x 0.48 ± 0.01, P < 0.001, Fig. 6CGo, lane 5 compared with 7). Like TSH, (Bu)2cAMP increased Glut-1 level at cell surface (x 2.16 ± 0.03, P < 0.001, Fig. 6CGo, lane 3 compared with 11) and LY294002 inhibited the (Bu)2cAMP-induced translocation of Glut-1 toward cell surface (x 0.47 ± 0.02, P < 0.001, lane 3 compared with 1). These data allowed us to conclude that the increase in glucose consumption observed after 2 h insulin or TSH stimulations might be explained by the translocation of Glut-1 from the intracellular pool to the plasma membranes. Wortmannin and LY294002 inhibited this process and as a consequence decreased the stimulation of glucose consumption or glucose transport by hormones.



View larger version (42K):
[in this window]
[in a new window]
  		Figure 6. Intracellular and surface Glut-1 content in FRTL-5 cell isolated after biotinylation of cell surface proteins. FRTL-5 cells preincubated in Coon’s modified Ham’s F12 medium containing only either transferrin and 0.2% FCS or 0.2% BSA in the absence or in the presence 1 µM wortmannin (A and B), 20 µM LY294002 (C) or 25 µM H89 (D) for 30 min, were treated either without (control) or with 5 µg/ml insulin (A), 1 mU/ml TSH (B and C) or 1 mM (Bu)2cAMP (C and D) for 2 h, followed by incubation with NHS-LC-biotin (0.25 mg/ml). Following homogenization, extracts were incubated with streptavidin-agarose beads. After removal of the supernatant and washing, cell surface proteins were solubilized in Laemmli’s buffer. Proteins of the supernatant (10 µg) following reaction with streptavidin-agarose beads [Intracellular (IC), Fig. 6Go, A and B, lanes 1, 3, 5, 7; Fig. 6CGo, lanes 2, 4, 6, 8, 10, 12 and Fig. 6DGo, lanes 2, 4, 6, 8), and surface proteins (10 µg) (Surface (S), Fig. 6Go, A and B, lanes 2, 4, 6, 8; Fig. 6CGo, lanes 1, 3, 5, 7, 9, 11 and Fig. 6DGo, lanes 1, 3, 5, 7] were separated on 10% SDS-PAGE and analyzed by Western blotting using specific antibodies to Glut-1. Each point represents the mean ± SD. The data were representative of three to five independent experiments.

 
Effect of PKA inhibitor on glucose transport and Glut-1 expression
Together, these results suggest that PI3-kinase dependent signaling pathway is required for insulin-, TSH-, or (Bu)2cAMP-stimulated translocation of Glut-1. The question now is to determine how does (Bu)2cAMP stimulated the translocation of Glut-1. Recently, it has been demonstrated that (Bu)2cAMP, which activates principally and directly protein kinase A (PKA), also activates others targets, such as ion channels and Rap-specific guanine nucleotide exchange factors (36, 37). Therefore, we next examined whether (Bu)2cAMP effects on the translocation of Glut-1 were PKA dependent. Treatment of FRTL-5 cells with the PKA inhibitor H89 did not reduce (Bu)2cAMP-stimulated glucose transport (Fig. 4BGo). Similarly, H89 did not reduce (Bu)2cAMP-stimulated Glut-1 translocation to the cell surface (Fig. 6DGo, lane 1 compared with 3). These results suggest that (Bu)2cAMP exerts stimulatory effects on glucose transport and Glut-1 translocation to the cell surface, effects that are PKA independent but PI3-kinase dependent.

Effect of tunicamycin on glucose consumption and Glut-1 expression in FRTL-5 cells
We further attempted to examine the role of the N-glycosylation in both activity (glucose consumption) and translocation toward cell surface of Glut-1. For this purpose, cells were incubated for 18 h in media without or with (10 µg/ml) tunicamycin. At the end of the incubation, time media and corresponding cell layers were collected. Glucose in cell media and Glut-1 protein in whole cell homogenates were evaluated per microgram of cell DNA as indicated in Fig. 7Go, A and B. Cell surface proteins were biotinylated and separated from cell extracts. Intracellular (IC) and surface (S) Glut-1 were analyzed by immublotting (Fig. 7CGo).



View larger version (28K):
[in this window]
[in a new window]
  		Figure 7. Effects of tunicamycin (Tu) on the activity and the cell surface delivery of Glut-1 in FRTL-5 cells. Cells were treated for 18 h in Coon’s medium supplemented with 6H and 5% FCS with or without 10 µg/ml of Tu. A, Effect of the inhibition of N-glycosylation on glucose consumption. Each point represents the mean ± SD. Statistically significant differences between control and tunicamycin (***), P < 0.001. B, Western blot analysis of total Glut-1 in FRTL-5 cells. Samples were analyzed by SDS-PAGE. C, Intracellular (IC) and cell surface (S) Glut-1 were then separated by the cell surface biotinylation method and analyzed by SDS-PAGE. D. Quantification by scanning the bands corresponding to Glut-1. Numbers represent the respective apparent molecular mass of the nonglycosylated (41 kDa) or glycosylated (50 kDa) forms of Glut-1 at the intracellular (IC) or plasma membrane (S) levels. The data were representative of three independent experiments.

 
In the presence of tunicamycin, glucose consumption decreased by about 50% compared with control value (Fig. 7AGo) without any loss of cell viability (data not shown). In the whole homogenate, some 70% of Glut-1 expressed in the presence of tunicamycin were not glycosylated and migrated as a 41-kDa protein (see Fig. 7Go, B and C and D), whereas the remaining fraction of Glut-1 that is still glycosylated migrated as a 50-kDa protein (Fig. 7Go, B and C and D). Interestingly, these two forms reached the cell surface and 70% of Glut-1 linked to the plasma membrane was not glycosylated (Fig. 7Go, C and D). Given the fact that glucose transport declines up to 50% of its initial value, these results strongly suggest that N-glycans are essential for Glut-1 activity but are not required for the intracellular transport of Glut-1 to the plasma membrane.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the thyroid, the presence of glucose transporter Glut-1 has been demonstrated in FRTL-5 cells (10) and then in human tissue. For the first time in this study, we demonstrated the presence of Glut-1 isoform in porcine thyroid cells from primary culture and from Athos and Porthos cell lines. In FRTL-5 cells Glut-1 is insulin, TSH and IGF-1 sensitive (10). Under hormonal stimulation, Glut-1 messenger RNA was increased 5- to 8-fold over control (38). We demonstrate here that most of the early stimulation by insulin or TSH is mediated by an increase in the translocation of Glut-1 to the plasma membrane, whereas in chronic treatment the enhanced amount of Glut-1 is due to an increase in total cellular Glut-1. However, the exact mechanism determining the fate of Glut-1, its translocation from the intracellular postGolgi vesicles up to the plasma membrane as well as the molecular events occurring concomitantly, are still undetermined.

A series of recent data indicates that PI3-kinase is involved in insulin-induced Glut-4 and Glut-1 translocation (6, 19, 39, 40, 41). We demonstrated here that the translocation of Glut-1 in the rat thyroid FRTL-5 cells is inhibited by wortmannin and LY294002, two specific PI3-kinase inhibitors. We first confirmed, that insulin or TSH increased glucose consumption and expression of Glut-1 protein in agreement with Filetti et al. (9) and Russo et al. (10). To identify the molecular mechanisms, we employed cell surface biotinylation followed by isolation of surface proteins and quantitative Glut-1 Western blotting. This accurate cell surface biotinylation technique allowed us to appreciate the cell surface expression of Glut-1 and to compare this expression with that of the intracellular compartment.

Increased glucose consumption in this system was significant at 2 h exposure to TSH (+45%) or insulin (+40%) despite the absence of variation in Glut-1 expression. This was explained by an increase in the membrane-bound Glut-1 and a decrease in the intracellular Glut-1. These observations were identical in insulin- or TSH-stimulated cells. Similar findings have also been reported in studies in 3T3-L1 adipocytes (6), in CHO cells (42) and in thyroid cells (10). Consequently, a short-term stimulation resulted in an increase in glucose consumption and in the translocation of Glut-1 to plasma membrane. After 24 h stimulation, glucose consumption was significantly increased both by insulin (+72%) and TSH (+73%) and Glut-1 expression also increased in parallel. But this was likely due to an increase of Glut-1 expression resulting from a de novo synthesis as shown by the experiments performed in the presence of cycloheximide. The increased amount of total cellular Glut-1 in 24 h and in the absence of hormone and cycloheximide (control conditions) could be due to the low intracellular accumulation of newly synthesized Glut-1 constituting the intracellular pool. Furthermore, it seems that Glut-1 in this pool is not yet functional (glycosylation, folding, degradation, and intracellular trafficking). These results suggest that, in FRTL-5 cells, insulin and TSH stimulate glucose transport either by recruitment of Glut-1 from an intracellular pool to plasma membrane or by enhancing the newly synthesized Glut-1 molecules through insulin or TSH signals. The increase in Glut-1 protein and glucose transport were previously correlated with the increase in Glut-1 messenger RNA TSH- or insulin-stimulated (10, 38). According to the high insulin concentrations used, an effect via the IGF-1 receptor cannot be excluded more especially as Russo et al. (10) demonstrated that IGF-1 was able to increase glucose transport via Glut-1 translocation in FRTL-5 cells.

Consequently, to gain understanding on the molecular mechanism driving Glut-1 translocation, we selected the 2 h incubation experiments to avoid interference with protein synthesis. Concerning the mechanism of Glut-1 translocation, two pathways have been proposed; the Ras-activated MAP kinase and the PI3-kinase pathway. Although a change in glucose transport occurs in response to elevation of Ras (39, 43) and MAP kinase expression (44), these effects may be due to an increased transcription (39, 43) and/or translocation of Glut-1. The possibility that glucose transport activation occurs through coupling of PI3-kinase activity to IRS-1 has been described (19, 40). PI3-kinase that participates in the regulation of cell growth (12, 45, 46) is activated by several tyrosine kinase receptors including the insulin receptor. Many studies demonstrate that the inhibition of PI3-kinase by wortmannin or LY294002 leads to a blockade in insulin-stimulated glucose transport (6, 19, 40, 41). Wortmannin impairs the translocation of Glut-1 in CHO cells and in 3T3-L1 cells without interfering directly with the glucose transporters (6). In FRTL-5 cells, we demonstrate that wortmannin under our conditions induces a decrease in glucose consumption both in TSH-stimulated and in insulin-stimulated FRTL-5 cells. Wortmannin and LY294002 also affect TSH, (Bu)2cAMP and insulin effects on Glut-1 translocation to plasma membranes. In conclusion, our data suggest that the PI3-kinase inhibitors, wortmannin and LY294002, block insulin TSH and (Bu)2cAMP effects at an early step in the thyroid system. In contrast, H89 does not affect TSH and (Bu)2cAMP effects on Glut-1 translocation to cell surface. This difference in actions of inhibitors of PKA and PI3-kinase on glucose transport and Glut-1 translocation to plasma membrane suggests a divergence in (Bu)2cAMP effects through PKA-dependent pathways and PI3-kinase-dependent pathways on glucose transport and Glut-1 translocation to plasma membrane. This hypothesis is strengthened by the fact that PI3-kinase is known to be involved in other membrane trafficking events, especially because it has been demonstrated that the VPS 34 protein (an analog of PI3-kinase in the yeast) operates in conjunction with a protein kinase in controlling membrane trafficking events (47, 48).

Insulin acts through an activation of PI3-kinase and our results obtained with wortmannin are in this case easy to understand. However, an activation of PI3-kinase by TSH has never been described so far. We hypothesized that a common intermediate could exist at the intersection between the two intracellular cascades either at the level of, or in upstream position, to PI3-kinase. This hypothesis strengthened, by works of Takahashi et al. (49, 50) and Ito et al. (51) in FRTL-5 cells and hepatocytes. These authors demonstrated that cAMP-dependent and insulin-dependent signals converge into a common pathway at the level of tyrosine phosphorylation. Secondly, since the signaling via transmembrane receptors including the receptor of TSH lead to a massive increase of PtdIns (3.4.5)P3 produced by a trimeric G protein-coupled ({alpha} and ß{gamma} subunits) sensitive to a novel PI3-kinase, named PI3-kinase {gamma}. This PI3-kinase {gamma} has been described recently as a dual kinase capable of phosphorylating lipids and proteins. Both activities are inhibited by wortmannin (52, 53). The implication of this PI3-kinase {gamma}, which is activated by a trimeric G protein-coupled({alpha} and ß{gamma} subunits) coupled to seven transmembrane receptors and inhibited by wortmannin, could explain the TSH effect on Glut-1 translocation via the PI3-kinase. Recently, the works of Cass et al. (54) and Medina et al. (46) in rat thyroid cells from WRT and FRTL-5 lines, respectively, demonstrated that PKA-dependent and PI3-kinase-dependent signals pathways contribute to cAMP-stimulated proliferation. These authors also demonstrated that cAMP effects on membrane ruffling and Akt (a serine/threonine-specific protein kinase that mediates many of the effects of PI3-kinase) are PKA independent but PI3-kinase dependent, while those on p70s6k (another member of the ribosomal S6 kinase family that is involved in cell proliferation) require PKA but not PI3-kinase activity (46, 54).

Because the targeting and activity of many glycoproteins can be affected by their glycosylation, we next tempted to clarify the relationship between N-glycosylation of Glut-1 and glucose consumption. The oligosaccharide structures of many glycoproteins play an important role in recognition of determinants, folding of proteins and biological activities (55). Likewise, these parameters are of the highest importance for facilitated diffusion. We demonstrated an association between the N-linked glycosylation of Glut-1 and glucose consumption in FRTL-5 cells. Tunicamycin-induced inhibition of glucose consumption correlated with marked changes in the electrophoretic mobility of Glut-1 protein. Several studies have been performed to elucidate the functional role N-glycosylation of Glut-1 protein. These studies have provided conflicting results and the biological role of N-glycosylation of Glut-1 protein remains unclear. Asano et al. (23), reported a decrease in affinity for glucose and in fine a decrease in transport activity after mutagenesis of the N-linked glycosylation site and expression in CHO cells. Haspel et al. (22) demonstrated rather an increase in glucose transport activity after depletion of aspargine-linked oligosaccharide in murine fibroblasts deprived of glucose. A decrease in Vmax for glucose uptake without a significant change in the Km had previously been reported in 3T3 cells after inhibition of glycosylation with tunicamycin (24). Feugeas et al. (56) described a loss of transport activity after N-glycopeptidase F treatment of Glut-1 in human erythrocytes. In our system, such treatment of membranes with N-glycopeptidase F resulted in a deglycosylation of Glut-1 leading to a 41-kDa protein representing the molecular mass of the core protein. These experiments allowed us to detect the unglycosylated and the glycosylated forms of Glut-1 in whole cells.

Our experiments with tunicamycin clearly showed that the inhibition of N-glycosylation did not affect the intracellular targeting of Glut-1. This is in the contrast with the behavior of some other integral membrane proteins. Using the biotinylation technique, we demonstrate that both the glycosylated and unglycosylated forms of the transporter reach the cell surface. The biological activity of nonglycosylated Glut-1 is still unknown, but according to Fig. 6AGo glucose consumption is decreased by 50%, whereas only 30% of the glycosylated Glut-1 reaches the cell surface. However, 70% of nonglycosylated Glut-1 reaches the cell surface too. This suggests that the unglycosylated Glut-1 transporter is still active but less efficient for glucose transport. This hypothesis corroborates data of Ahmed and Berridge (25) showing that the affinity for glucose of the unglycosylated Glut-1 expressed by leukemic cells is 2-fold decreased (Km x 2). Therefore, the N-glycosylation of Glut-1 plays a crucial role in maintaining a fully active structure of the transporter but does not interfere with its intracellular targeting. The existence of an other unsuspected pathway for glucose entry unsensitive to cytochalasin B and to some other intracellular traffic inhibitors (57) could also explain the discrepancy observed between the activity of the nonglycosylated Glut-1 (which could be inactive) compared with the glycosylated one.


   Acknowledgments
 
We thank Prof. H. Rochat (C.H.U. Timone, Marseille) for glucose measurements performed with the analyzer Hitachi 717 and Dr. J.-L. Franc (INSERM Unité 38, Marseille) for his encouragement and helpful discussions during this study.

Received March 9, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Mueckler M 1994 Facilitative glucose transporters. Eur J Biochem 219:713–725[Medline]
  2. Turk E, Kerner CJ, Lostao MP, Wright EM 1996 Membrane topology of the human Na+/glucose cotransporter SGLT-1. J Biol Chem 271:1925–1934[Abstract/Free Full Text]
  3. Birnbaum MJ, Haspel HC, Rosen OM 1986 Cloning and characterization of a cDNA encoding the rat brain glucose-transporter protein. Proc Natl Acad Sci USA 83:5784–5788[Abstract/Free Full Text]
  4. Berridge MV, Tan AS, McCoy KD, Kansara M, Rudert F 1996 CD95 (Fas/Apo-1)-induced apoptosis results in loss of glucose transporter function. J Immunol 156:4092–4099[Abstract]
  5. Wertheimer E, Sasson S, Cerasi E, Ben-Neriah Y 1991 The ubiquitous glucose transporter GLUT-1 belongs to the glucose-regulated protein family of stress-inducible proteins. Proc Natl Acad Sci USA 88:2525–2529[Abstract/Free Full Text]
  6. Clarke JF, Young PW, Yonezawa K, Kasuga M, Holman GD 1994 Inhibition of the translocation of GLUT1 and GLUT4 in 3T3–L1 cells by the phosphatidylinositol 3-kinase inhibitor, wortmannin. Biochem J 300:631–635
  7. Whitesell RR, Abumrad NA 1985 Increased affinity predominates in insulin stimulation of glucose transport in the adipocyte. J Biol Chem 260:2894–2899[Abstract/Free Full Text]
  8. Trischitta V, Damante G, Foti D, Filetti S 1987 Insulin binding and biological activities in the FRTL-5 rat thyroid cell line. Metabolism 36:379–383[CrossRef][Medline]
  9. Filetti S, Damante G, Foti D 1987 Thyrotropin stimulates glucose transport in cultured rat thyroid cells. Endocrinology 120:2576–2581[Abstract]
  10. Russo D, Damante G, Foti D, Costante G, Filetti S 1994 Different molecular mechanisms are involved in the multihormonal control of glucose transport in FRTL5 rat thyroid cells. J Endocrinol Invest 17:323–327[Medline]
  11. Samih N, Aouani A, Hovsépian S, Amphoux T, Fayet G The glucose transporters Glut-1 expressed in thyroid cells from different species or physiopathological origin differ mainly by their oligosaccaride moieties. Program of the 26th Annual Meeting of the European Thyroid Association, Milan, 1999, p 44 (Abstract)
  12. Hawkins PT, Jackson TR, Stephens LR 1992 Platelet-derived growth factor stimulates synthesis of PtdIns(3,4,5)P3 by activating a PtdIns(4,5)P2 3-OH kinase. Nature 358:157–159[CrossRef][Medline]
  13. Hiles ID, Otsu M, Volinia S, Fry MJ, Gout I, Dhand R, Panayotou G, Ruiz-Larrea F, Thompson A, Totty NF, Hsuan JJ, Courtneidge SA, Parker PJ, Waterfield MD 1992 Phosphatidylinositol 3-kinase: structure and expression of the 110 kd catalytic subunit. Cell 70:419–429[CrossRef][Medline]
  14. Escobedo JA, Navankasattusas S, Kavanaugh WM, Milfay D, Fried VA, Williams LT 1991 cDNA cloning of a novel 85 kd protein that has SH2 domains and regulates binding of PI3-kinase to the PDGF beta-receptor. Cell 65:75–82[CrossRef][Medline]
  15. Koch CA, Anderson D, Moran MF, Ellis C, Pawson T 1991 SH2 and SH3 domains: elements that control interactions of cytoplasmic signaling proteins. Science 252:668–674[Abstract/Free Full Text]
  16. Ruderman NB, Kapeller R, White MF, Cantley LC 1990 Activation of phosphatidylinositol 3-kinase by insulin. Proc Natl Acad Sci USA 87:1411–1415[Abstract/Free Full Text]
  17. Yamamoto K, Altschuler D, Wood E, Horlick K, Jacobs S, Lapetina EG 1992 Association of phosphorylated insulin-like growth factor-I receptor with the SH2 domains of phosphatidylinositol 3-kinase p85. J Biol Chem 267:11337–11343[Abstract/Free Full Text]
  18. Yano H, Nakanishi S, Kimura K, Hanai N, Saitoh Y, Fukui Y, Nonomura Y, Matsuda Y 1993 Inhibition of histamine secretion by wortmannin through the blockade of phosphatidylinositol 3-kinase in RBL-2H3 cells. J Biol Chem 268:25846–25856[Abstract/Free Full Text]
  19. Okada T, Kawano Y, Sakakibara T, Hazeki O, Ui M 1994 Essential role of phosphatidylinositol 3-kinase in insulin-induced glucose transport and antilipolysis in rat adipocytes. Studies with a selective inhibitor wortmannin. J Biol Chem 269:3568–3573[Abstract/Free Full Text]
  20. Kotani K, Yonezawa K, Hara K, Ueda H, Kitamura Y, Sakaue H, Ando A, Chavanieu A, Calas B, Grigorescu F, Nishiyama M, Waterfield MD, Kasuga M 1994 Involvement of phosphoinositide 3-kinase in insulin- or IGF-1-induced membrane ruffling. EMBO J 13:2313–2321[Medline]
  21. Vlahos CJ, Matter WF, Hui KY, Brown RF 1994 A specific inhibitor of phosphatidylinositol 3-kinase, 2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one (LY294002). J Biol Chem 269:5241–5248[Abstract/Free Full Text]
  22. Haspel HC, Revillame J, Rosen OM 1988 Structure, biosynthesis, and function of the hexose transporter in Chinese hamster ovary cells deficient in N-acetyl-glucosaminyl-transferase 1 activity. J Cell Physiol 136:361–366[CrossRef][Medline]
  23. Asano T, Katagiri H, Takata K, Lin JL, Ishihara H, Inukai K, Tsukuda K, Kikuchi M, Hirano H, Yazaki Y 1991 The role of N-glycosylation of GLUT1 for glucose transport activity. J Biol Chem 266:24632–24636[Abstract/Free Full Text]
  24. Kitagawa T, Tsuruhara Y, Hayashi M, Endo T, Stanbridge EJ 1995 A tumor-associated glycosylation change in the glucose transporter GLUT1 controlled by tumor suppressor function in human cell hybrids. J Cell Sci 108:3735–3743[Abstract]
  25. Ahmed N, Berridge MV 1999 N-glycosylation of glucose transporter-1 (Glut-1) is associated with increased transporter affinity for glucose in human leukemic cells. Leuk Res 23:395–401[CrossRef][Medline]
  26. Ambesi-Impiombiato FS, Parks LAM, Coon HG 1980 Culture of hormone-dependent functional epithelial cells from rat thyroids. Proc Natl Acad Sci USA 77:3455–3459[Abstract/Free Full Text]
  27. Hovsépian S, Fayet G 1993 In vitro reorganization of porcine thyroid cells from the ATHOS line into follicle-like structures. Cell Mol Biol 39:427–433
  28. Cheatham B, Vlahos CJ, Cheatham L, Wang L, Blenis J, Kahn CR 1994 Phosphatidylinositol 3-kinase activation is required for insulin stimulation of pp 70 s6 kinase, DNA synthesis, and glucose transporter translocation. Mol Cell Biol 14:4902–4911[Abstract/Free Full Text]
  29. Labarca C, Païgen K 1980 A simple, rapid, and sensitive DNA assay procedure. Anal Biochem 102:344–352[CrossRef][Medline]
  30. Shetty M, Loeb JN, Vikstrom K, Ismail-Beigi F 1993 Rapid activation of GLUT-1 glucose transporter following inhibition of oxidative phosphorylation in clone 9 cells. J Biol Chem 268:17225–17232[Abstract/Free Full Text]
  31. Fayadat L, Niccoli-Sire P, Lanet J, Franc JL 1998 Human thyroperoxidase is largely retained and rapidly degraded in the endoplasmic reticulum. Its N-glycans are required for folding and intracellular trafficking. Endocrinology 139:4277–4285[Abstract/Free Full Text]
  32. Laemmli UK 1970 Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685[CrossRef][Medline]
  33. Haspel HC, Rosenfeld MG, Rosen OM 1988 Characterization of antisera to a synthetic carboxyl-terminal peptide of the glucose transporter protein. J Biol Chem 263:398–403[Abstract/Free Full Text]
  34. Tarentino AL, Gomez CM, Plummer TH 1985 Deglycosylation of asparagine-linked glycans by peptide: N-glycosidase F. Biochemistry 34:4665–4671
  35. Mueckler M, Caruso C, Baldwin SA, Panico M, Blench I, Morris HR, Allard WJ, Lienhard GE, Lodish HF 1985 Sequence and structure of a human transporter. Science 229:941–945[Abstract/Free Full Text]
  36. de Rooij J, Zwartkruis FT, Verheijen MHG, Cook RH, Nijman SMB, Wittinghofer A, Bos JL 1998 Epac is a Rap1 guanine-nucleotide-exchange factor directly activated by cyclic AMP. Nature 396:474–477[CrossRef][Medline]
  37. Kawasaki H, Springett GM, Mochizuki N, Toki S, Nakaya M, Matsuda M, Housman DE, Graybiel AM 1998 A family of cAMP-binding proteins that directly activate Rap1. Science 282:2275–2279[Abstract/Free Full Text]
  38. Hosaka Y, Tawata M, Kurihara A, Ohtaka M, Endo T, Onaya T 1992 The regulation of two distinct glucose transporter (GLUT1 and GLUT4) gene expressions in cultured rat thyroid cells by thyrotropin. Endocrinology 131:159–165[Abstract]
  39. Hiraki Y, Rosen OM, Birnbaum MJ 1988 Growth factors rapidly induce expression of the glucose transporter gene. J Biol Chem 263:13655–13662[Abstract/Free Full Text]
  40. Kanai F, Ito K, Todaka M, Hayashi H, Kamohara S, Ishii K, Okada T, Hazeki O, Ui M, Ebina Y 1993 Insulin-stimulated GLUT4 translocation is relevant to the phosphorylation of IRS-1 and the activity of PI3-kinase. Biochem Biophys Res Commun 195:762–768[CrossRef][Medline]
  41. Cheatham B, Vlahos CJ, Cheatham L, Wang L, Blenis J, Kahn CR 1994 Phosphatidylinositol 3-kinase activation is required for insulin stimulation of pp70 S6 kinase, DNA synthesis, and glucose transporter translocation. Mol Cell Biol 14:4902–4911
  42. Hara K, Yonezawa K, Sakaue H, Ando A, Kotani K, Kitamura T, Kitamura Y, Ueda H, Stephens L, Jackson TR, Hawkins PT, Dhand R, Clark AE, Holman GD, Waterfield MD, Kasuga M 1994 1-Phosphatidylinositol 3-kinase activity is required for insulin-stimulated glucose transport but not for RAS activation in CHO cells. Proc Natl Acad Sci USA 91:7415–7419[Abstract/Free Full Text]
  43. Rollins BJ, Morrison ED, Usher P, Flier JS 1988 Platelet-derived growth factor regulates glucose transporter expression. J Biol Chem 263:16523–16526[Abstract/Free Full Text]
  44. Merrall NW, Plevin RJ, Stokoe D, Cohen P, Nebreda AR, Gould GW 1993 Mitogen-activated protein kinase (MAP kinase), MAP kinase kinase and c-Mos stimulate glucose transport in Xenopus oocytes. Biochem J 295:351–355
  45. Cantley LC, Auger KR, Carpenter C, Duckworth B, Graziani A, Kapeller R, Soltoff S 1991 Published erratum appears in Cell; 65: following 914 oncogenes and signal transduction. Cell 64:281–302[CrossRef][Medline]
  46. Medina DL, Toro MJ, Santisteban P 2000 Somatostatin interferes with thyrotropin-induced G1-S transition mediated by cAMP-dependent protein kinase and phosphatidylinositol 3-kinase. J Biol Chem 275:15549–15556[Abstract/Free Full Text]
  47. Schu PV, Takegawa K, Fry MJ, Stack JH, Waterfield MD, Emr SD 1993 Phosphatidylinositol 3-kinase encoded by yeast VPS34 gene essential for protein sorting. Science 260:88–91[Abstract/Free Full Text]
  48. Stack JH, Herman PK, Schu PV, Emr SD Rollins BJ, Morrison ED, Stiles CD 1993 A membrane-associated complex containing the Vps15 protein kinase and the Vps34 PI 3-kinase is essential for protein sorting to the yeast lysosome-like vacuole. EMBO J 12:2195–2204[Medline]
  49. Takahashi SI, Conti M, Van Wyk JJ 1990 Thyrotropin potentiation of insulin-like growth factor-I dependent deoxribonucleic acid synthesis in FRTL-5 cells: mediation by an autocrine amplification factor(s). Endocrinology 126:736–745[Abstract]
  50. Takahashi SI, Conti M, Prokop C, Van Wyk JJ, Earp HS 1991 Thyrotropin and insulin-like growth factor I regulation of tyrosine phosphorylation in FRTL-5 cells. Interaction between cAMP-dependent and growth factor-dependent signal transduction. J Biol Chem 266:7834–7841[Abstract/Free Full Text]
  51. Ito Y, Uchijima Y, Ariga M, Seki T, Takenaka A, Hakuno F, Takahashi SI, Ariga T, Noguchi T 1997 Interaction between cAMP-dependent signal pathways in tyrosine phosphorylation in primary cultures of rat hepatocytes. Biochem J 324:379–388
  52. Stoyanova S, Bulgarelli-Leva G, Kirsch C, Hanck T, Klinger R, Wetzker R, Wymann MP 1997 Lipid kinase and protein kinase activities of G-protein-coupled phosphoinositide 3-kinase {gamma}: structure-activity analysis and interactions with wortmannin. Biochem J 324:489–495
  53. Stoyanov B, Volinia S, Hanck T, Rubio I, Loubtchenkov M, Malek D, Stoyanova S, Vanhaesebroeck B, Dhand R, Nürnberg B, Gierschik P, Secdorf K, Hsuan JJ, Waterfield MD, Wetzker R 1995 Cloning and characterization of a G protein-activated human phosphoinositide-3 kinase. Science 269:690–693[Abstract/Free Full Text]
  54. Cass LA, Summers SA, Prendergast GV, Backer JM, Birnbaum MJ, Meinkoth JL 1999 Protein kinase A-dependent and -independent signaling pathways contribute to cyclic AMP-stimulated proliferation. Mol Cell Biol 19:5882–5891[Abstract/Free Full Text]
  55. Rademacher TW, Parekh RB, Dwek RA 1988 Glycobiology. Annu Rev Biochem 57:785–838[CrossRef][Medline]
  56. Feugeas JP, Neel D, Pavia AA, Laham A, Goussault Y, Derappe C 1990 Glycosylation of the human erythrocyte glucose transporter is essential for glucose transport activity. Biochim Biophys Acta 1030:60–64[Medline]
  57. Guillam MT, Burcelin R, Thorens B 1998 Normal hepatic glucose production in the absence of GLUT2 reveals an alternative pathway for glucose release from hepatocytes. Proc Natl Acad Sci USA 95:12317–12321[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 JNMHome page
D. Blaser, S. Maschauer, T. Kuwert, and O. Prante
In Vitro Studies on the Signal Transduction of Thyroidal Uptake of 18F-FDG and 131I-Iodide
J. Nucl. Med., August 1, 2006; 47(8): 1382 - 1388.
[Abstract] [Full Text] [PDF]

Home page
 J. Clin. Endocrinol. Metab.Home page
L. Hooft, A. A. M. van der Veldt, P. J. van Diest, O. S. Hoekstra, J. Berkhof, G. J. J. Teule, and C. F. M. Molthoff
[18F]Fluorodeoxyglucose Uptake in Recurrent Thyroid Cancer Is Related to Hexokinase I Expression in the Primary Tumor
J. Clin. Endocrinol. Metab., January 1, 2005; 90(1): 328 - 334.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Endocrinol.Home page
J. S. Richards, S. C. Sharma, A. E. Falender, and Y. H. Lo
Expression of FKHR, FKHRL1, and AFX Genes in the Rodent Ovary: Evidence for Regulation by IGF-I, Estrogen, and the Gonadotropins
Mol. Endocrinol., March 1, 2002; 16(3): 580 - 599.
[Abstract] [Full Text] [PDF]

Home page
 EndocrinologyHome page
G. Napolitano, I. Bucci, C. Giuliani, C. Massafra, C. Di Petta, E. Devangelio, D. S. Singer, F. Monaco, and L. D. Kohn
High Glucose Levels Increase Major Histocompatibility Complex Class I Gene Expression in Thyroid Cells and Amplify Interferon-{gamma} Action
Endocrinology, March 1, 2002; 143(3): 1008 - 1017.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Purchase Article
Right arrow View Shopping Cart
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow