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Endocrinology, doi:10.1210/en.2007-0985
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Endocrinology Vol. 149, No. 3 917-924
Copyright © 2008 by The Endocrine Society

Mitogen-Stimulated and Rapamycin-Sensitive Glucose Transporter 12 Targeting and Functional Glucose Transport in Renal Epithelial Cells

Amy L. Wilson-O’Brien, Carrie L. DeHaan and Suzanne Rogers

The University of Melbourne, Department of Medicine (A.L.W.-O., C.L.D., S.R.), St. Vincent’s Hospital, Melbourne, Fitzroy, Victoria 3065, Australia; and The University of Melbourne, Department of Genetics (A.L.W.-O.), Parkville, Victoria 3052, Australia

Address all correspondence and requests for reprints to: Dr. Suzanne Rogers, The University of Melbourne, Department of Medicine, St. Vincent’s Hospital, Melbourne, Fitzroy, Victoria 3065, Australia. E-mail: s.rogers{at}medicine.unimelb.edu.au.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We hypothesized that glucose transporter 12 (GLUT12) is involved in regulation of glucose flux in distal renal tubules in response to elevated glucose. We used the Madin-Darby canine kidney polarized epithelial cell model and neutralizing antibodies to analyze GLUT12 targeting and directional GLUT12-mediated glucose transport. At physiological glucose concentrations, GLUT12 was localized to a perinuclear position. High glucose and serum treatment resulted in GLUT12 localization to the apical membrane. This mitogen-stimulated targeting of GLUT12 was inhibited by rapamycin, the specific inhibitor of mammalian target of rapamycin (mTOR). The functional role of GLUT12 was also examined. We constructed a GLUT12 cDNA containing a c-Myc epitope tag in the fifth exofacial loop. Assays of glucose transport at the apical membrane were performed using Transwell filters. By comparing transport assays in the presence of neutralizing anti-c-Myc monoclonal antibody, we specifically measured GLUT12-mediated glucose transport at the apical surface. GLUT12-mediated glucose transport was mitogen dependent and rapamycin sensitive. Our results implicate mTOR signaling in a novel pathway of glucose transporter protein targeting and glucose transport. Activity of the mTOR pathway has been associated with diabetic kidney disease. Our results provide evidence for a link between GLUT12 protein trafficking, glucose transport and signaling molecules central to the control of metabolic disease processes.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
FACILITATED TRANSPORT OF glucose across the plasma membrane of mammalian cells occurs via glucose transporter (GLUT) proteins. This process is essential both to provide cells with nutrients and energy and for the maintenance of whole-body glucose homeostasis (1). To achieve this critical balance of changing glucose flux, the rate of glucose transport into cells is regulated at a number of levels. Members of the GLUT family exhibit distinct glucose transport kinetic capabilities and tissue-specific patterns of expression that reflect energy requirements and metabolism (2). Variable response to changing glucose conditions is also achieved by transcriptional and posttranscriptional regulation of GLUT protein levels and activity (3). A further level of control is provided through regulated subcellular localization of GLUT proteins in response to growth factors and environmental conditions. The prime example of GLUT protein trafficking is the acute insulin-regulated localization of GLUT4. GLUT4 is expressed in muscle and fat, residing in distinct intracellular compartments. When blood glucose and insulin levels rise, GLUT4 undergoes rapid insulin-regulated movement to the cell surface, where it facilitates glucose transport into muscle and fat cells under a concentration gradient. The complex cellular mechanisms that elicit GLUT4 insulin-responsive trafficking via the insulin receptor include phosphatidylinositol 3-kinase (PI3K)-dependent and -independent signaling cascades and cellular structural elements (4, 5).

In metabolic diseased states such as type 2 diabetes, insulin resistance exists. Insulin resistance occurs in part from defective GLUT4 trafficking, and as a result, blood glucose can rise to pathological levels. Chronic hyperglycemia eventually leads to tissue damage and serious diabetic complications, including diabetic nephropathy (6, 7, 8). Hyperglycemia and glomerular filtration are compensated by increased glucose reabsorption in the renal tubules, forming part of the chain of events that leads to kidney damage (9). In diabetic, hyperglycemic rats, this increased glucose transport has been shown to be associated with both increased expression and altered localization of GLUT proteins (10, 11, 12). We have previously characterized the renal expression of the facilitative glucose transporter GLUT12. In the fetal rat, GLUT12 protein expression has been demonstrated in a wide range of tissues, including the distal tubules and collecting ducts of the kidney (13). We have also shown that GLUT12 is expressed in the distal tubules and collecting ducts of the adult rat kidney and that expression is elevated in an animal model of diabetic nephropathy (14). In this study GLUT12 was localized to a perinuclear region and at the apical membrane of tubular epithelial cells.

Our findings led us to hypothesize that GLUT12 is involved in the regulation of glucose flux in distal portions of the kidney. Second, we hypothesized that GLUT12 is targeted to the apical membrane in response to elevated glucose. In the present study, we have used the Madin-Darby canine kidney (MDCK) polarized epithelial cell model of renal ductal tubules to examine the functional role of GLUT12 in renal glucose flux. This model allows us to analyze directional glucose transport and to localize GLUT proteins in a spatial manner. We have demonstrated apical targeting and GLUT12-mediated glucose transport in response to elevated glucose. Targeting of GLUT12 and glucose transport was mitogen dependent and sensitive to rapamycin. Our results implicate the involvement of the mammalian target of rapamycin (mTOR) signaling pathway in a novel pathway of GLUT protein targeting. Activity of the mTOR pathway has been associated with kidney disease (15). Our results provide evidence for a link between GLUT12 protein targeting, glucose transport, and signaling molecules central to the control of metabolic disease processes.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Antibodies and chemicals
Rabbit polyclonal anti-GLUT12 has been previously described (16). Anti-zonula occludens-1 (anti-ZO-1) was obtained from Zymed Laboratories (San Francisco, CA), and anti-GLUT1 and anti-GLUT3 antibodies were purchased from Abcam (Cambridge, MA). Anti-c-Myc and anti-hemagglutinin (anti-HA) antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-pan-actin antibody was obtained from Neomarkers (Fremont, CA). Fluorescein isothiocyanate-conjugated (Dako, Carpinteria, CA) or Alexa-568-conjugated (Molecular Probes, Eugene, OR) secondary antibodies were used for immunofluorescence studies and horseradish peroxidase-conjugated (Dako) secondary antibodies for immunoblotting. All antibody dilutions were in 5% fetal bovine serum (FBS)/PBS for immunofluorescence work and 2% skim milk powder/Tris-buffered saline for immunoblotting. All chemicals were purchased from Sigma Chemical Co. (St. Louis, MO) unless otherwise specified.

Cells
Cell lines were maintained in DMEM, 25 mM glucose (Life Technologies, Inc., Grand Island, NY) supplemented with 5% FBS (JRH Bioscience, Lenexa, KS) at 37 C in a humidified chamber unless otherwise specified. The human packaging cell line Phoenix gag-pol (http://www.stanford.edu/group/nolan/retroviral_systems/phx.html) was received from Dr. L. Corcoran (The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia, with authorization by Prof. G. Nolan, School of Medicine, Stanford University, Stanford, CA). MDCK cells were obtained from the Department of Genetics (University of Melbourne, Parkville, Australia). MDCK cells were grown for 3–4 d to a confluent monolayer, which indicated polarization. Stably transfected cell lines were maintained in DMEM/5% FBS with 3 µg/ml puromycin.

Plasmid constructs
The DsRED-GLUT12 construct was engineered by amplifying GLUT12 cDNA with primers to introduce 5' EcoRI and 3' BamHI sites that eliminated the 5' and 3' untranslated regions and the stop codon. The PCR product was ligated into pDsRED2-N1 (Clontech, Mountain View, CA) in frame so that the DsRED protein would be expressed as a GLUT12 C-terminal fusion protein. Integrity of the construct was confirmed by bidirectional sequencing. The human GLUT3 cDNA (a kind gift from Prof. G. Bell, University of Chicago, Chicago, IL) was subcloned into the pBABE-puro retroviral expression vector (a kind gift from Prof. D. James, Garvan Institute, Sydney, Australia) using the SalI restriction site to produce BABE-GLUT3. The BABE-GLUT12myc clone was engineered using standard molecular biology subcloning techniques so that it contained 10 amino acids of the c-Myc epitope (5'-GAG CAG AAG CTG ATC AGC GAG GAG GAC CTG-3') in the fifth exofacial loop of GLUT12. The pHEF-VSVG construct (17) was a kind gift from Dr. A. Poumbourios (Burnet Institute, Melbourne, Australia).

Transient transfections
MDCK cells were seeded onto glass coverslips 24 h before being transfected. Cells were then transfected with DsRED-GLUT12 using FuGENE HD (Roche Diagnostics, Mannheim, Germany) in Opti-MEM (Life Technologies) according to manufacturer’s instructions. Once polarized, cells were treated and fixed as described.

Retroviral transfections
BABE-GLUT3 and BABE-GLUT12myc were each cotransfected along with pHEF-VSVG into Phoenix gag-pol cells with FuGENE 6 (Roche) using a 9:1 FuGENE 6/DNA ratio in Opti-MEM. Supernatant containing the viral particles was collected at 72, 96, and 102 h after transfection and snap frozen. Supernatant (450 µl) was then used to infect MDCK cells at 1.5 x 105 cells per well with 4 µg/ml polybrene per well of a 12-well dish and incubated at 32 C for 18 h (18). The next day, the supernatant was removed and replaced with DMEM/10% FBS and grown at 37 C until confluent. Transfected cells were then selected in 3 µg/ml puromycin.

Glucose/mannitol, rapamycin, and wortmannin treatments
Transfected and nontransfected MDCK cells were grown on glass coverslips and polarized in DMEM/10% FBS. For chronic glucose and serum treatments, cells were grown, polarized, and treated 16 h overnight with one of the following: 1) 25 mM glucose/10% FBS, 2) 5 mM glucose/10% FBS, 3) 25 mM glucose/0.1% BSA, 4) 5 mM glucose/0.1% BSA, 5) 25 mM mannitol/10% FBS, 6) 25 mM mannitol/0.1% BSA, 7) 5 mM mannitol/10% FBS, or 8) 5 mM mannitol/0.1% BSA. For all rapamycin and wortmannin studies, MDCK cells were incubated with rapamycin (Cell Signaling Technology, Danvers, MA) or wortmannin for 30 min at 10 or 100 nM, respectively, in treatment media. Cells were then prepared for immunofluorescence studies, immunoblotting, or glucose uptake assays.

Immunofluorescence
After treatment, cells were washed in PBS and fixed in 4% paraformaldehyde. Fixed cell monolayers were then incubated in 100 mM glycine, permeabilized in 0.1% Triton X-100, blocked in 10% normal swine serum, 5% FBS/PBS for 30 min (all at room temperature), and incubated overnight at 4 C with primary antibody. This was followed by incubation with the appropriate secondary antibody for 1 h at room temperature. After washing with PBS, nuclei were stained with either TOTO-3 (Molecular Probes) at 1:1000 in PBS/0.1% Tween 20 or propidium iodide at 7.5 ng/ml in PBS/0.1% Tween 20 and 1.5 µg/ml RNase (Roche). Coverslips were then mounted in the fluorescent aqueous mountant VectaShield (Vector Laboratories, Burlingame, CA). A Bio-Rad MCR1024 inverted laser scanning confocal microscope (Bio-Rad, Hercules, CA) and Lasersharp 2000 software were used for analysis.

Immunoblotting
Cell pellets were lysed by resuspension in lysis buffer [50 mM Tris (pH 7.5), 150 mM NaCl, 1 mM EDTA, 0.5% Triton X-100, 1 mM phenylmethylsulfonyl fluoride] and drawn through a 21-gauge needle. Collection of tissue samples was approved by the University of Melbourne Human Research Ethics Committee (approval no. 030079) and the St. Vincent’s Health Animal Ethics Committee (approval number 35/00). Frozen tissue was homogenized in lysis buffer (Polytron homogenizer; Kinematica, Lucerne, Switzerland). Cell lysates were then centrifuged at 65,000 rpm for 1 h in a TL700 ultracentrifuge (Beckman-Coulter, Fullerton, CA) with resulting membrane protein pellet resuspended in lysis buffer. Protein concentrations in cell extracts were determined by using Bradford protein assay reagent (Bio-Rad). Each sample (55 µg) was separated by SDS-PAGE and immunoblotted as previously described (14). Filters were stripped for 20 min at room temperature in Re-Blot Plus Mild (Chemicon, Temecula, CA) and protein loading verified by immunolabeling for pan-actin.

Glucose uptake assays
MDCK cells were grown in either 12-well dishes or Transwell Permeable Supports (24 mm diameter, 0.4 µm pore size, polycarbonate membrane; Corning Inc., Corning, NY) for apical/basolateral studies. Before the assay, cells were polarized and treated as previously described. Cells were then equilibrated for 30 min with equal volumes of treatment media and 2x uptake solution. Where used, rapamycin concentration was 10 nM and wortmannin 100 nM. Equal volumes of 2x uptake solution (200 mM NaCl, 4 mM KCl, 2 mM MgCl2, 1.2 mM NaH2PO4, 10 mM HEPES, 4 mM CaCl2) containing either 25 mM or 5 mM cold 2-D-deoxyglucose and 1 µCi/ml 2-[1,2-3H(N)]deoxy-D-glucose (PerkinElmer, Boston, MA) were added. Transport was allowed to proceed for 5 min at room temperature. Where used, cytochalasin B concentration was 15 µM. Uptake was stopped with cold PBS/100 µM phloretin, and cells were washed four times with cold PBS and lysed overnight in 1 ml 1% SDS. Cell lysate (800 µl) was diluted in 3 ml Insta-Gel Plus scintillant (PerkinElmer) and incorporation of radioactivity measured using a 1600A TRI-CARB liquid scintillation analyzer (PerkinElmer). The remaining 200 µl of cell lysate was used for protein estimations using Bradford protein assay reagent (Bio-Rad). Each experiment was performed at least three times with triplicate samples. Representative experiments are shown. Statistical analyses of data were performed using one-way ANOVA and unpaired t test. P values of<0.05 were considered significant.

Neutralizing antibody glucose uptake assays
Blocking studies were done with Transwell filters using MDCK cells stably expressing GLUT12myc. Anti-c-Myc or anti-HA antibody (2.5 µg/ml) was included in the overnight treatment media in the apical (upper) or basolateral (lower) chamber. Anti-c-Myc or anti-HA antibodies were also added to the equilibration buffer at a final concentration of 2.5 µg/ml in the uptake reaction. The glucose uptake assay then proceeded as previously described.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Subcellular distribution of GLUT12 after exposure to glucose and serum
The subcellular distribution of GLUT12 was investigated in polarized MDCK cells using confocal fluorescence microscopy. The GLUT12 polyclonal antibody was raised to a peptide based on the 16 carboxyl-terminal amino acids of the human GLUT12 sequence (16). Fourteen of these 16 amino acids are conserved between the human GLUT12 protein and the predicted dog peptide sequence. To confirm cross-reactivity with human and canine GLUT12, and thus its suitability for detection of GLUT12 in MDCK cells, we performed control Western immunolabeling (supplemental Fig. 1, published as supplemental data on The Endocrine Society’s Journals Online web site at http://endo.endojournals.org). Polarization of MDCK cells was confirmed by immunofluorescent labeling of the tight junction marker ZO-1 as shown in green (Fig. 1Go, A–H). Nonimmune serum control is shown in Fig. 1AGo. After 16 h growth of MDCK cells in medium containing 25 mM glucose and 10% FBS, endogenously expressed GLUT12 was immunolabeled and localized to the cytoplasm and to the plasma membrane. Confocal sections in the Z-plane demonstrated GLUT12 immunolabeling in an arc above the ZO-1 tight junction marker, indicating apical localization (in red, with TOTO-3 nuclear staining in blue, Fig. 1BGo). Some GLUT12 staining was also noted below the ZO-1 tight junction marker (Fig. 1BGo). After 16 h growth in 0.1% BSA and 5 mM glucose, GLUT12 protein was concentrated in a perinuclear position (in red, with TOTO-3 nuclear staining in blue, Fig. 1CGo). Apical localization of GLUT12 was not observed after 16 h growth in osmotic control media containing 25 mM mannitol and no glucose (Fig. 1DGo).


Figure 1
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  		FIG. 1. Localization of GLUT12 in MDCK cells exposed to glucose and serum. Subcellular localization of GLUT12 was studied using confocal microscopy. Polarization of cells was confirmed by immunofluorescent labeling of the tight junction marker ZO-1 as shown in green in panels A–H. GLUT12 was localized by immunofluorescent labeling. A, Negative, nonimmune serum control experiment. After 16 h growth of MDCK cells in high-glucose medium and serum (25 mM glucose, 10% FBS), immunofluorescence detected endogenous GLUT12 in the cytoplasm and at the plasma membrane (in red, panel B). Confocal sections in the Z-plane localized GLUT12 specifically to the apical membrane (in red, with TOTO-3 nuclear stain in blue, panel B). In comparison, immunofluorescent labeling of MDCK cells after 16 h growth in media containing 5 mM glucose, 0.1% BSA localized GLUT12 predominantly to the cytoplasm in a perinuclear position (in red, with TOTO-3 nuclear stain in blue, panel C). After 16 h growth in medium containing 25 mM mannitol and no glucose, GLUT12 was localized to a perinuclear position (in red, panel D). To confirm the differential localization of GLUT12 under these growth conditions, MDCK cells were transfected with the DsRED-tagged GLUT12 cDNA. Polarized, nontransfected cells are shown in panel E. When transfected MDCK cells were grown in high-glucose medium and serum before fixation, DsRED-GLUT12 was localized to the cytoplasm and at the apical plasma membrane (panel F). DsRED-GLUT12 was restricted to a tight, perinuclear localization if cells were grown for 16 h in medium containing 5 mM glucose and no serum (panel G). DsRED-GLUT12 was also localized to a tight perinuclear position after 16 h growth in 25 mM mannitol (panel H). Scale bar, 20 µm; original magnification, x600.

 
To confirm the localization of GLUT12 after these treatment conditions, MDCK cells were transfected with DsRED-tagged GLUT12 cDNA. In the presence of 25 mM glucose and 10% serum, DsRED-tagged GLUT12 protein was detected at the plasma membrane, and confocal sections in the Z-plane displayed a similar localization pattern to that of endogenously expressed GLUT12 (Fig. 1FGo). In agreement with the localization of endogenous GLUT12, DsRED-GLUT12 was detected in a perinuclear position when cells were grown for 16 h in 5 mM glucose medium and no serum or media containing 25 mM mannitol (Fig. 1Go, G and H). The apical localization of GLUT12 (in red, Fig. 2DGo) was confirmed by colocalization with GLUT3. MDCK cells were retrovirally transfected with GLUT3 cDNA and immunolabeled GLUT3 localized to the apical membrane (shown in confocal sections of the Z-plane in red, Fig. 2CGo). Expression of DsRED-GLUT12 (Fig. 2Go, D and G) and GLUT3 (in green, Fig. 2Go, E and H) confirmed that both proteins localize at the apical surface of MDCK cells. Merged confocal sections demonstrate colocalization of GLUT3 and GLUT12 (shown in yellow in the X-Y and X-Z plane, Fig. 2Go, F and I).


Figure 2
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  		FIG. 2. Colocalization of GLUT3 and GLUT12 in MDCK cells. Polarization of cells was confirmed by immunofluorescent labeling of the tight junction marker ZO-1 as shown in green in panels A–C. A, Negative, nonimmune serum control experiment. B, Negative immunofluorescent labeling confirmed that GLUT3 is not endogenously expressed in MDCK cells. C, After retroviral transfection, GLUT3 was detected by immunofluorescent labeling, with sections in the Z-plane demonstrating localization to the apical membrane (in red). D and G, MDCK cells stably transfected with GLUT3 were transiently cotransfected with the DsRED-GLUT12 construct (in red). E and H, Apical localization of GLUT3 (in green). F and I, Merged images of MDCK cells expressing both GLUT3 and DsRED-GLUT12. Panels F and I also show a Z-plane section confirming the apical localization of GLUT12 when cells were grown in high-glucose medium and serum. Scale bar, 20 µm; original magnification, x600.

 
GLUT12-mediated glucose transport is mitogen stimulated
The functional significance of the differential localization of GLUT12 in renal glucose flux was studied via apical 2-deoxy-D-[H3]glucose uptake assays. This study used Transwell filters, allowing the addition of glucose to either the apical or basolateral surfaces of the epithelial cell and subsequent vectorial transport analysis. Apical glucose transport rates after 16 h growth in medium containing 25 mM glucose and 10% serum were significantly increased when compared with cells grown for 16 h in medium containing 5 mM glucose or in the absence of serum (Fig. 3AGo). Apical glucose transport was not stimulated after 16 h growth of cells in medium containing 25 mM mannitol (Fig. 3AGo). Nonspecific glucose transport levels were determined using cytochalasin B, a specific inhibitor of facilitative glucose transport. The results suggest that glucose transport via the apical membrane is mitogen stimulated, requiring the presence of both glucose and serum.


Figure 3
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  		FIG. 3. Mitogen-stimulated apical glucose transport is mediated by GLUT12. A, MDCK cells were polarized in 12-well tissue culture dishes to form intact tight monolayers, allowing assay of glucose transport at the apical surface. Glucose transport was measured by uptake of 2-deoxy-D-[3H]glucose. Nonspecific transport was assessed in the presence or absence of the specific inhibitor of facilitative glucose transport, cytochalasin B (CB). Facilitative glucose transport at the apical surface was significantly increased after 16 h growth in 25 mM glucose and 10% FBS, when compared with glucose transport after 16 h growth in medium containing 5 mM glucose and no serum or mannitol. Mean and SD of transport rates were calculated. Experiments were repeated at least three times, with the results of a representative experiment shown. #, P < 0.001 when compared with all groups. B, Confocal immunofluorescence of MDCK cells expressing GLUT12myc and labeled with anti-c-Myc antibody demonstrated the apical localization of exofacially c-Myc-tagged GLUT12: i, negative, nonimmune serum control experiment, with propidium iodide nuclear stain in red; ii, apical localization of GLUT12 in green with propidium iodide nuclear stain in red. Scale bar, 20 µm; original magnification, x600. C, To determine whether mitogen-stimulated glucose transport at the apical membrane could be attributed to GLUT12 specifically, MDCK cells were stably transfected with the GLUT12myc exofacially tagged cDNA construct. Apical glucose transport assays were then performed on Transwell filters in the presence or absence of neutralizing antibody. Apical transport was inhibited in the presence of neutralizing anti-c-Myc monoclonal antibody but not in control assays performed in the presence of anti-HA monoclonal antibody. D, MDCK cells stably transfected with the GLUT12myc exofacially tagged cDNA construct were grown on Transwell filters and allowed to polarize, and glucose transport was assayed at the basolateral surface. No difference in glucose transport was observed when assays were performed in the presence of either anti-c-Myc or anti-HA neutralizing antibodies. Mean and SD of transport rates were calculated. Experiments were repeated at least three times with the results of a representative experiment shown. #, P < 0.01 when compared with all groups.

 
To determine whether this mitogen-stimulated apical glucose uptake can be attributed specifically to GLUT12, neutralizing antibody experiments were performed. A GLUT12 cDNA was constructed containing a 10-amino-acid c-Myc epitope tag in the large, fifth, exofacial loop. MDCK cells were retrovirally transfected with c-Myc-tagged GLUT12, and immunolabeling localized GLUT12myc to the apical membrane using anti-c-Myc antibodies (Fig. 3BGo). As noted above for endogenously expressed GLUT12, some GLUT12myc labeling was also noted below the ZO-1 tight junction marker. Assays of glucose transport at the apical membrane were performed using Transwell filters. By comparing transport assays in the presence of neutralizing anti-c-Myc monoclonal antibody, we could specifically measure GLUT12-mediated glucose transport at the apical surface. In the presence of neutralizing antibody, glucose uptake via the apical membrane was significantly decreased in comparison with cells treated with control anti-HA antibody (Fig. 3CGo). To address the observation that both endogenously expressed and c-Myc-tagged GLUT12 was detected at the apical membrane and also observed below the ZO-1 tight junction marker, we performed additional experiments measuring glucose transport at the basolateral face of MDCK cells. MDCK cells transfected with GLUT12myc were used in these experiments. No difference in glucose transport was observed when assays were performed in the presence of either anti-c-Myc or anti-HA neutralizing antibodies (Fig. 3DGo).

Mitogen-stimulated glucose transport and GLUT12 apical targeting are inhibited by rapamycin and wortmannin
mTOR controls many mitogen-activated signaling pathways. To further examine the characteristics of mitogen-dependent GLUT12 subcellular localization and GLUT12-mediated glucose transport in renal epithelial cells, we used the specific mTOR inhibitor, rapamycin. After growth in mitogen-stimulated conditions, a 30-min treatment with rapamycin reduced facilitative glucose uptake levels at the apical surface to that observed after 16 h growth in non-mitogen-stimulated conditions (Fig. 4AGo). Wortmannin, a PI3K inhibitor, also reduced apical glucose uptake levels to that of the non-mitogen-stimulated state. Again, nonspecific glucose transport levels were determined using cytochalasin B. Confocal immunofluorescence indicated that the decrease in facilitative glucose transport levels correlated to a change in subcellular localization of GLUT12. Localization of GLUT12 was restricted to a perinuclear location after a 30-min rapamycin or wortmannin treatment of MDCK cells maintained in the mitogen-stimulated state (Fig. 4BGo). Immunoblotting experiments also showed that rapamycin treatment did not alter the levels of endogenously expressed GLUT12 protein (Fig. 4CGo).


Figure 4
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  		FIG. 4. Mitogen-stimulated glucose transport and GLUT12 apical targeting are inhibited by rapamycin and wortmannin. A, Glucose uptakes were performed as described. After a 30-min incubation with the specific inhibitor of mTOR, rapamycin (R), or with the PI3K inhibitor wortmannin (W), facilitative glucose transport rates were reduced to levels observed in the non-mitogen-stimulated state. Mean and SD of transport rates were calculated. Experiments were repeated at least three times, and the results of a representative experiment are shown. #, P < 0.001 when compared with all groups. B, Confocal immunofluorescence also revealed that 30 min incubation with rapamycin or wortmannin prevented apical targeting of GLUT12 in high-glucose medium and serum. Immunolabeling of the tight junction marker ZO-1 is shown in green and immunolabeling of GLUT12 in red (B, i–iv): i, negative nonimmune serum control experiment; ii, apical localization of GLUT12; iii and iv, cells treated with either rapamycin or wortmannin, respectively. Scale bar, 20 µm; original magnification, x600. C, Immunoblotting of endogenously expressed GLUT12. Primary antibody was preincubated in the presence of either non-competitive (NCP) or competitive (CP) peptide before immunoblotting. Filters were stripped and subsequently immunoblotted with pan-actin monoclonal antibody to verify protein loading. Growth of MDCK cells in the presence of rapamycin (R) for 30 min did not alter expression levels of GLUT12.

 
High glucose and serum does not alter basolateral glucose transport rate or subcellular localization of GLUT1
In control experiments, Transwell filters were used to assay glucose transport at the basolateral surface of MDCK cells. MDCK cells were grown on Transwell filters and allowed to polarize. After 16 h growth in either media containing high glucose and serum or 5 mM glucose and no serum, glucose transport was assayed at the basolateral surface. Facilitative glucose transport rates at the basolateral membrane were unaltered after 16 h growth in high glucose and serum (Fig. 5AGo). As shown by confocal sections in the X-Y and X-Z planes, the subcellular localization of the endogenously expressed basolateral glucose transporter, GLUT1, was also unchanged (Fig. 5BGo). Thus, apical glucose transport via GLUT12 but not basolateral glucose transport via GLUT1 is altered after growth in high glucose and serum.


Figure 5
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  		FIG. 5. Growth in high glucose and serum does not alter basolateral glucose transport rate or the localization of GLUT1. A, MDCK cells were grown on Transwell filters and allowed to polarize, and glucose transport was assayed at the basolateral surface. Glucose transport was measured by uptake of 2-deoxy-D-[3H]glucose. Nonspecific transport was assessed in the presence of the specific inhibitor of facilitative glucose transport, cytochalasin B (CB). The rate of glucose transport at the basolateral surface did not differ between cells that had been grown for 16 h in high-glucose media and serum and those grown for 16 h in media containing 5 mM glucose and no serum (5 mM glucose, 0.1% BSA). B, Localization of GLUT1: i, immunolabeling of the tight junctional marker ZO-1 in green; ii, GLUT1 immunofluorescent labeling, with sections in the Z-plane demonstrating cytoplasmic staining and localization to the basolateral membrane in media containing 5 mM glucose and no serum (in red); iii, growth in high glucose and serum did not alter the localization of GLUT1 (in red). Scale bar, 20 µm; original magnification, x600.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Renal glucose flux is controlled by members of two glucose transporter families: the sodium-dependent glucose transporter (SGLT) and the facilitative glucose transporter (GLUT) proteins. In the normal kidney, the bulk of glucose reabsorption occurs in the S1 section of the proximal tubules via the coordinated action of SGLT2 and GLUT2 (19). In diabetic, hyperglycemic rats, increased glucose transport has been shown to be associated with both increased expression and altered localization of GLUT proteins. Modulation of GLUT1, GLUT2, and the fructose-specific transporter GLUT5 has been reported in response to the intracellular concentration of glucose (11, 12). More recently, up-regulation and basolateral localization of both GLUT8 and GLUT9 have been demonstrated in the renal distal tubules of diabetic mice (20, 21). Apical (brush border) targeting of GLUT2 has been demonstrated in the proximal region of the kidney in diabetic rats (22). Interestingly, in this study, Marks et al. (22) showed that GLUT2 insertion into the brush border membrane was a specific response to hyperglycemia, because apical targeting was abolished after overnight fasting. However, in in vitro studies, exogenously expressed GLUT2 has been localized only at the basolateral surface of MDCK cells, even when the cells were grown in high-glucose medium and serum (23, 24). Thus, it is unclear at this time whether the mechanisms controlling mitogen-activated GLUT12 apical targeting may also regulate localization of GLUT2.

We have previously shown that the facilitative glucose transporter protein GLUT12 is expressed in renal distal tubules, specifically at the apical membrane (14). Our work has also demonstrated that GLUT12 resides in both a perinuclear and plasma membrane localization, and we have hypothesized that GLUT12 undergoes regulated trafficking to the cell surface (16). In the current study, we have used the MDCK epithelial cultured cell model to investigate GLUT12 trafficking after growth in an environment with elevated glucose levels. This model is ideal because the cells express GLUT12 endogenously and are derived from renal distal tubular epithelium. In addition, MDCK cells form polarized monolayers and have been used previously to characterize the targeting and transport properties of GLUT proteins (23).

To test the hypothesis that targeting of GLUT12 occurs in response to elevated glucose levels, we grew MDCK monolayers for 16 h in media containing either physiological (5 mM) or high (25 mM) glucose concentrations. Confocal immunofluorescence microscopy localized GLUT12 to a perinuclear position in physiological glucose conditions. When cells were grown in high-glucose media, GLUT12 was detected both intracellularly and specifically at the apical plasma membrane. High-glucose conditions did not alter the basolateral localization of GLUT1. To confirm specificity of detection, we used MDCK cells transiently transfected with DsRED-tagged GLUT12 cDNA. The pattern of localization of tagged GLUT12 in normal and high-glucose media was identical to that shown by immunofluorescence. Further confirmation of the apical localization of GLUT12 in high-glucose media was obtained by colocalization with GLUT3. The GLUT3 facilitative glucose transporter has previously been shown to be specifically targeted at the apical membrane of MDCK cells (23).

The mechanisms that dictate apical protein targeting are not well understood. Some specific amino acid targeting domains have been identified. For example, a domain in the carboxyl-terminal region of GLUT3 governs apical targeting of this transporter. Similar domains are not present in the GLUT12 sequence. In the main, in contrast to basolateral sorting, apical sorting signals are not restricted to specific motifs but also include posttranslational modifications such as glycosylation (25) and possible association with lipid rafts (26). Potential sites for N-linked glycosylation are present in the large predicted exofacial loop between GLUT12 membrane helices 9 and 10. The apical localization of GLUT12 in MDCK cells is consistent with our previous in vivo findings where GLUT12 was localized to the apical membrane of renal distal tubules (14). In our present study, endogenously expressed and tagged GLUT12 was detected at the apical membrane and also observed below the ZO-1 tight junction marker. It is possible that GLUT12 may transiently cycle via the basolateral to the apical membrane. Models of transcytotic apical targeting have been described, although there is some controversy as to the activity in MDCK cells (26). Our control basolateral transport assays show that neutralizing antibodies have no effect on basolateral glucose uptake. Thus, if GLUT12 were trafficking to the apical surface via a transcytotic route, across the basolateral surface, it would appear that any basolateral delivery is transient or unable to mediate functional transport. The nature of the intracellular compartment(s) in which GLUT12 resides in the absence of high glucose is not known. The trans-Golgi network is a major site of protein sorting, and recent evidence also indicates that the recycling endosome compartments may also be important (27). The apical targeting of GLUT12 in response to elevated external glucose concentrations implicates this transporter in glucose reabsorption when the glucose transport capacity of the proximal tubules is exceeded.

Interestingly, we observed that targeting of GLUT12 to the apical membrane in the presence of high glucose occurred only if serum was also present in the cell culture media. This finding led us to consider whether targeting of GLUT12 in epithelial cells is controlled by the mitogen-stimulated mTOR pathway (28). We subsequently demonstrated that the specific mTOR-Raptor complex inhibitor rapamycin acutely blocks mitogen-stimulated apical targeting of GLUT12. The nutrient-sensitive mTOR signaling pathway is a major regulator of cell mass and proliferation and is regulated by signals including amino acids, glucose, ATP, and insulin. mTOR signaling predominantly controls downstream increases in the activity of transcriptional activators (29, 30). A role for the rapamycin-insensitive mTOR-Rictor cascade in cytoskeleton regulation has also been described (31), and mTOR signaling has also been implicated in endocytosis (32). Our work now suggests an additional role for the rapamycin-sensitive mTOR-Raptor signaling pathway in protein trafficking. Increased osmotic load has also been shown to activate mTOR signaling cascades (33). In our current experiments, GLUT12 was retained to a tight perinuclear position in the presence of 25 mM mannitol, indicating that the apical targeting of GLUT12 in high-glucose and serum conditions was not a response to osmotic pressure.

Neutralizing antibodies can be used to block cell surface receptors such as HER-2/neu (34). Here we have adapted this methodology and used neutralizing monoclonal antibodies directed to an exofacial epitope tag between helices 9 and 10 to specifically block GLUT12-mediated glucose transport. This approach verified functional mTOR-regulated, mitogen-stimulated GLUT12 apical targeting. GLUT12 is a class III facilitative glucose transporter. This class is unique from other GLUTs in that they possess a large predicted exofacial loop between helices 9 and 10. Our results also suggest a critical role for this type of structure in membrane topology and glucose transport function. Parallel to results of our localization studies, GLUT12-mediated glucose transport at the apical surface of MDCK cells was stimulated after growth in high-glucose media. Also paralleling our localization studies, stimulation required both high glucose and serum and was acutely inhibited by rapamycin. GLUT12-mediated glucose transport was blocked by cytochalasin B, a specific inhibitor of facilitative glucose transporters. Again reflecting our localization studies, GLUT12-mediated glucose transport was not increased by osmotic load. The Km of GLUT12-mediated glucose transport and thus the substrate concentration at which saturation is reached have not as yet been determined. Therefore, substrate concentration gradients, as well as GLUT12 targeting to the membrane, may also contribute to glucose transport rates.

The mTOR complexes initiate diverse signaling cascades, many of which are incompletely understood. Two major pathways involve either the rapamycin-sensitive association of mTOR-Raptor or the rapamycin-insensitive mTOR-Rictor complex (31). GLUT12 apical targeting and increased glucose transport in response to nutrients were completely abolished after 30 min exposure to rapamycin, implicating involvement of the mTOR-Raptor cascade. Apical targeting of GLUT2 in the gut has recently been demonstrated in response to elevated luminal glucose levels. In comparison with our results, glucose-responsive targeting of GLUT2 was dependent on protein kinase CβII and calcium levels (35).

Class III PI3K has been implicated in nutrient stimulation of the mTOR-Raptor complex, and it is thought that the wortmannin-sensitive target of amino acid-induced mTOR-Raptor complex 1 is distinct from the class I PI3K-PKB/Akt signaling pathway (36). Our data demonstrating inhibition of GLUT12 mitogen-stimulated targeting and glucose transport by wortmannin is consistent with this prediction. Glucose is thought to regulate mTOR in concert with AMP-activated kinase (36). It is possible that glucose-responsive GLUT12 targeting and glucose transport is a result of increased glycolysis and regulated by altered ATP levels.

Our current study demonstrates apical targeting of GLUT12 in response to nutrients. GLUT12-mediated glucose transport is mitogen stimulated and inhibited by rapamycin, suggesting regulation by the mTOR-Raptor signaling pathway. These findings implicate GLUT12 glucose reabsorption in the diabetic kidney under conditions of increased glomerular glucose flux. The mTOR pathway has recently been linked to the progression of diabetic nephropathy (37). High glucose is thought to induce mTOR-dependent changes in protein synthesis and diabetes-induced renal hypertrophy (38). Our current results provide evidence for a mechanistic link between activation of mTOR, elevated renal glucose transport, and diabetic kidney disease.


   Acknowledgments
 
We thank James Best, Natalie Burgess, and Maria Macheda for reading of the manuscript, Darren Kelly for useful discussions, James Camakaris for provision of the MDCK cell line, and Glenn McConell and Frank Alford for tissue samples.


   Footnotes
 
This research was supported by a National Health and Medical Research Council (Australia) Project Grant No. 350424 and the Diabetes Australia Research Trust.

Disclosure Statement: The authors have nothing to disclose.

First Published Online November 26, 2007

See editorial p. 913.

Abbreviations: FBS, Fetal bovine serum; GLUT, glucose transporter; HA, hemagglutinin; MDCK, Madin-Darby canine kidney; mTOR mammalian target of rapamycin; PI3K, phosphatidylinositol 3-kinase; ZO-1, zonula occludens-1.

Received July 18, 2007.

Accepted for publication November 12, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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H. Schmid, M. Bertoluci, and T. M. Coimbra
Glucose Transporter 12 and Mammalian Target of Rapamycin Complex 1 Signaling: A New Target for Diabetes-Induced Renal Injury?
Endocrinology, March 1, 2008; 149(3): 913 - 916.
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