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(Investigative Ophthalmology and Visual Science. 2002;43:2356-2363.)
© 2002 by The Association for Research in Vision and Ophthalmology, Inc.

Glucose-Induced Activation of Glucose Uptake in Cells from the Inner and Outer Blood–Retinal Barrier

Julia V. Busik1, L. Karl Olson1, Maria B. Grant2 and Douglas N. Henry1,3

1 From the Departments of Physiology and 3 Pediatrics and Human Development, Michigan State University, East Lansing, Michigan; and the 2 Department of Pharmacology and Therapeutics, University of Florida, Gainesville, Florida.


   Abstract
Top
 Abstract
Introduction
 Methods
 Results
 Discussion
 References
 
PURPOSE. The purpose of this study was to elucidate in vitro the effect of elevated glucose on glucose uptake in the cells comprising the inner and outer blood–retinal barriers: human retinal pigment epithelial (hRPE) and human retinal vascular endothelial (hRVE) cells.

METHODS. Primary cultures of hRPE and hRVE cells grown in 5.5 or 22 mM glucose or in 22 mM mannitol were used to measure the rates of glucose uptake with [3H]-3-O-methyl glucose as a tracer. GLUT1 expression was measured by Northern and western blot analyses. Cellular fractionation was performed by differential centrifugation. GLUT1 overexpression was accomplished by adenoviral transduction.

RESULTS. Increasing media glucose from 5.5 to 22 mM for 30 minutes caused a 1.9-fold increase in the Vmax of glucose uptake in hRPE cells and a 2.5-fold increase in hRVE cells. These increases were nonosmotic and glucose specific, in that the exposure to 22 mM mannitol did not affect the Vmax of glucose uptake. mRNA, total protein expression, and translocation of GLUT1, the glucose transporter predominantly expressed in hRPE and hRVE cells, were not affected by 22 mM glucose for up to 48 hours. High-glucose effects on Vmax were abolished with 10 µg/mL of the microtubule assembly inhibitor nocodazole. hRPE cells transduced to overexpress GLUT1 showed a 1.5-fold increase in the Vmax for glucose uptake versus control-transduced cells. However, the magnitude of glucose-induced increase in glucose uptake was the same in GLUT1- and control-transduced cells.

CONCLUSIONS. High glucose induced 1.9- and 2.5-fold increases in the Vmax of glucose uptake in hRPE and hRVE cells, respectively. These increases were not due to an increase in GLUT1 expression. The increases were dependent on microtubule integrity, but not on GLUT1 translocation. The mechanism of the increases is unknown. GLUT1 regulating protein(s) and/or novel glucose transporter(s) may be involved in the regulation of glucose uptake by glucose in the cells that comprise the blood–retinal barrier.


   Introduction
Top
 Abstract
 Introduction
Methods
 Results
 Discussion
 References
 
Results of basic and clinical research over the past decade have removed any doubt that hyperglycemia is a major causative factor in the development of diabetic retinopathy. The Diabetes Control and Complications Trial Research Group1 has demonstrated a close correlation between tight glycemic control and prevention of diabetic retinopathy. The hypothesis of glucose toxicity combines several lines of investigation in an attempt to explain the end organ damage in diabetes.2 3 These include glucose-mediated increases in diacylglycerol production and the activation of the PKC pathway,4 5 6 increased flux of glucose through the polyol metabolic pathway,7 8 9 10 11 changes in the redox state,12 and detrimental expression of cytokine-growth factors.13 Glucose transport is a key proximal event prerequisite in all these disturbances. Accumulation of advanced glycation end (AGE) products14 is another part of the glucose toxicity theory. Although elevated extracellular glucose can increase AGE formation, intracellular sugars, such as glucose-6-phosphate and fructose, form AGE at a much faster rate, suggesting that formation of AGE in diabetes is dependent on glucose transport into the cells. To affect the cells within the retina, high glucose must first pass the blood–retinal barrier (BRB) composed of inner (retinal pigment epithelial cells [RPE]) and outer (retinal vascular endothelial cells [RVE]) layers. Thus, expression and regulation of facilitative glucose transporters in human RPE and RVE cells is a key to understanding the downstream effects of elevated glucose in diabetic retinopathy.

There are conflicting data in the literature on the effect of glucose on glucose transport and metabolism in the BRB. Glucose-specific increases in glucose utilization, lactate production, and aldose reductase (AR) expression in RPE cells have been demonstrated.8 Elevated glucose induces a downregulation of Na+/H+ antiport in hRPE cells15 and a decrease in nitric oxide (NO) synthase expression in RVE cells.16 Incubation of isolated retinas with elevated glucose causes an increase in glycolysis and a higher tissue content of lactic acid and adenosine triphosphate (ATP), indicative of greater glucose utilization.11 Diabetic retina has been shown to have an increased AGE level,17 expression of AR,10 PKC activation,6 decreased expression of NO synthase,18 and decreased Na,K-ATPase activity.19 20 However, elevated glucose has been reported to cause no change in the expression of glucose transporters and in glucose uptake in bovine RVE cells in vitro,21 and short-term diabetes has been reported to downregulate expression of GLUT1 in rat retinal microvessels.22 Changes in the expression of glucose transporters in human diabetic retina have been described only after long-standing diabetes.23 24 The effects of short-term exposure to high glucose on glucose uptake by the cells from human BRB have not been studied. Thus, initial determinants of the glucose toxicity leading to the development of diabetic retinopathy are still unknown. This study is the first to examine the effect of short-term exposure to elevated glucose on glucose uptake and expression of GLUT1 in hRPE and hRVE cells in vitro.


   Methods
Top
 Abstract
 Introduction
 Methods
Results
 Discussion
 References
 
Reagents and Supplies
RPMI 1640 culture medium, antibiotics, HBSS, and trypsin were obtained from GibcoBRL (Gaithersburg, MD); fetal calf serum from Hyclone Laboratories (Logan, UT); and culture dishes and flasks from Sarstedt, Inc. (Newton, NC). Glucose (tissue culture grade) and all other commonly used chemicals and reagents were from Sigma Chemical Co. (St. Louis, MO). {alpha}-32P dCTP, [3H]-3-O-methyl-D-glucose ([3H]-3-O-MG) and [14C]-L-glucose were from NEN Life Science Products (Boston, MA).

Cell Culture Techniques
In the present study, we used primary cultures of hRPE and hRVE cells. hRPE cells were established by a modification of the method of Del Monte and Maumenee,25 as described previously.26 hRVE cells were prepared as previously described.27 28 29 30 hRPE cells were cultured in RPMI-1640 medium supplemented with 15% fetal calf serum, 1% penicillin-streptomycin, and 5.5 mM glucose. hRVE cells were cultured in a 1:1 mix of low-glucose (1000 mg/mL) DMEM and F12 nutrient mix (GibcoBRL) supplemented with 10% fetal calf serum, endothelial cell growth supplement, insulin-transferrin-selenium mix, and 1% penicillin-streptomycin. Glucose concentration in the final media was adjusted to 5.5 mM. The cells were maintained at 37°C in 5% CO2 in a humidified cell culture incubator and passaged at a density of 40,000 to 100,000 cells/cm2 in 75-cm2 flasks. Passaged cells were plated to yield near-confluent cultures at the end of the experiments. Ten-centimeter plates were used for RNA and protein isolation, and six-well plates were used for glucose uptake experiments. The freshly plated cells were allowed to attach in standard growth medium for at least 24 hours before incubation for various periods in different concentrations of glucose or mannitol.

Immunoblotting of GLUT1
hRPE and hRVE cells were grown in 10-cm plates in experimental media for up to 48 hours. Each plate was rinsed twice with 5 mL ice cold phosphate-buffered saline (PBS) containing 130 mM NaCl, 8.2 mM Na2HPO4, and 1.8 mM NaH2PO4 (pH 7.4). The cells were then harvested and homogenized in a ground-glass homogenizer in 1 mL of the lysis buffer containing 50 mM HEPES (pH 7.5), 150 mM NaCl, 1.5 mM MgCl2, 1 mM EGTA, 1% TritonX-100, and 10% glycerol, with the freshly added inhibitors 1 mM sodium orthovanadate, 0.15 U/mL aprotinin, and 100 mg/mL phenylmethylsulfonyl fluoride (PMSF). Homogenates were centrifuged at 16,900g for 20 minutes at 4°C, and 100 µL of each supernatant was combined with 50 µL 3x SDS-containing sample buffer (30 mM Tris, 9% SDS, 15% glycerol, 0.05% bromphenol blue). Then 3.3 µL ß-mercaptoethanol was added to each sample. Samples corresponding to 10 µg cell protein were electrophoresed on 10% polyacrylamide minigels (Bio-Rad, Hercules, CA) along with prestained molecular weight standards. The separated proteins were electrophoretically transferred to nitrocellulose by conventional procedures.31 Nitrocellulose sheets were blocked for 60 minutes at room temperature in Tris-buffered saline (TBS; 130 mM NaCl, 100 mM Tris/HCl [pH 7.5]) containing 5% powdered milk and 0.05% Tween-20 and then incubated for 120 minutes at room temperature in a blocking buffer containing a 1:5000 dilution of a rabbit polyclonal antibody against the COOH terminus of rat GLUT1 (Chemicon International, Inc., Temecula, CA). After rinsing with TBS, the nitrocellulose sheets were incubated for 90 minutes in blocking buffer containing a 1:5000 dilution of a commercial (Sigma Chemical Co.) goat anti-rabbit IgG-peroxidase conjugate and then developed using a chemiluminescent substrate (Super Signal West Pico; Pierce, Rockford, IL) followed by immediate exposure to autoradiograph film (X-Omat Kodak, Rochester, NY) for 1 to 30 seconds. Blots were quantitated by scanning densitometry on a high-resolution optical scanner (AGFA; Orangeburg, NY) and NIH Image software (version 1.60; W. Rasband, National Institutes of Health; available by ftp from zippy.nimh.nih.gov or on floppy disc from NTIS, Springfield, VA, part number PB95-500195GEI).

Cellular Fractionation
Cellular fractions were prepared by the method of differential centrifugation with slight modifications.32 hRPE and hRVE cells grown in 10-cm diameter plates were exposed to experimental media for 30 minutes. The plates were then rapidly rinsed with ice-cold PBS and allowed to swell for 10 minutes at 4°C in buffer A (10 mM Tris [pH 7.5]; 35 mM NaF; 5 mM MgCl2; 1 mM EGTA with the freshly added inhibitors 1 mM sodium orthovanadate, 0.15 U/mL aprotinin, and 100 mg/mL PMSF). The cells were then scraped from the plates and homogenized with 20 strokes in a glass homogenizer with a tight-fitting pestle (Dounce; Kontes Glass, Vineland, NJ). The resultant homogenates were centrifuged for 5 minutes at 500g to pellet the nuclei. The supernatants were centrifuged at 16,900g for 1 hour at 4°C to obtain a sedimented plasma membrane–enriched fraction (P16.9). The P16.9 fraction was rinsed twice with buffer A and resuspended in buffer B (50 mM HEPES [pH 7.5]; 150 mM NaCl; 1.5 mM MgCl2; 1 mM EGTA; 1% TritonX-100, 10% glycerol with the freshly added inhibitors 1 mM sodium orthovanadate; 0.15 U/mL aprotinin and 100 mg/mL PMSF). The supernatants were subjected to ultracentrifugation at 210,000g for 90 minutes at 4°C to obtain a sedimented microsomal fraction (P210). The P210 fraction was rinsed twice with buffer A and resuspended in buffer B. Expression of Na,K-ATPase33 34 was used as a plasma membrane marker. The microsomal fraction was identified using ß1,4-galactosyltransferase for Golgi membranes. Anti-ß1,4-galactosyltransferase antibody was generously provided by Joel H. Shaper (The Johns Hopkins Oncology Center, Baltimore, MD). Cytosolic fraction was identified with cytosolic enzyme lactate dehydrogenase (LDH) as a marker.35 Anti-LDH antibody was the gift of John Wilson (Michigan State University, East Lansing, MI). Cell fractions containing equal amounts of proteins (10 µg) were resolved by Western Blot analysis, as described earlier.

GLUT1 mRNA Expression
GLUT1 mRNA levels were measured by Northern blot analysis. Total RNA from hRPE and hRVE cells grown in 5.5, 11, and 22 mM glucose for up to 48 hours was obtained by a modification of the acid-phenol extraction method and 10 µg RNA was resolved on denaturing 2.2-M formaldehyde-1% agarose gels.26 After electrophoresis, RNA was transferred to nylon filters (ZetaBind; Cuno Laboratory Products, Inc., Meriden, CT) by capillary blot. Filters were stained with methylene blue to examine the integrity of the RNA and to assess the uniformity of loading and transfer. The filters were fixed and hybridized at high stringency.36 Human GLUT1 cDNA generously provided by Frank C. Brosius (University of Michigan, Ann Arbor, MI),37 was used for making the GLUT1 probe. Chicken ß-actin cDNA38 was used for making the ß-actin probe. GLUT1 and ß-actin cDNA probes were labeled with 32P-dCTP using random primers to a specific activity of 109 dpm/mg and separated from unincorporated nucleotides by gel filtration. After 18 hours, hybridized filters were washed at high stringency.36 Autoradiograms were obtained with multiple exposures to remain within the linear range of the film and were quantitated by scanning densitometry, with the high-resolution optical scanner (AGFA) and NIH Image software. After initial determination that GLUT1 cDNA hybridizes to a single band of an appropriate size, all the blots were cohybridized with both GLUT1 and ß-actin.

Glucose Transport Assays
Initial rates of glucose transport were determined by using the principle that the glucose analogue 3-O-methyl glucose (3-O-MG) is transported into the cells through D-glucose transporters, but is not phosphorylated or further metabolized. Thus, the rate of intracellular accumulation of [3H]-3-O-MG during the linear portion of its uptake versus time represents the initial transport rate. The nontransported stereoisomer [14C]-L-glucose was used simultaneously to determine nonspecific association of hexose with the cell layer. hRPE and hRVE cells were plated on six-well plates, allowed to attach for at least 24 hours in standard medium, and exposed to experimental medium for 30 minutes, as indicated in the figure legends (Figs. 2 3 and 8) . The cells were rapidly washed three times at 37°C with 3 mL PBS containing different concentrations of glucose. The cells were then incubated at 37°C with 1 mL PBS containing 0 to 22 mM glucose, 3.26 µCi/mL [3H]-3-O-MG and 0.0022 µCi/mL [14C]-L-glucose. At the end of the incubation period, the uptake solution was aspirated, and 3 mL of blocking solution (ice-cold PBS containing 10 µM cytochalasin B) was added to the plate. Two additional 3-mL washes were performed, and the cells were solubilized in 1 mL 0.05 N NaOH. An 800-µL aliquot was used for beta scintillation counting in 3H and 14C windows, and the remainder was saved for protein determination. The rate of uptake of [3H]-3-O-MG was linear for only the first 20 seconds in cultures of hRPE cells (Fig. 1A) and the first 10 seconds in cultures of hRVE cells (Fig. 1B) . Based on these data, the kinetics of glucose uptake was measured at 12 seconds in hRPE and 5 seconds in hRVE cells.



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  		Figure 2. Kinetics of glucose uptake by hRPE cells. (A) Kinetics of glucose uptake was measured in hRPE cells incubated in 5.5 ({blacksquare}) or 22 mM (•) glucose for 30 minutes. (B) A Hanes plot calculation of the data was used to determine Km and Vmax. Data are the mean ± SE of results in three experiments, each performed in triplicate.

 


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  		Figure 3. Kinetics of glucose uptake by hRVE cells. (A) Kinetics of glucose uptake was measured in hRVE cells incubated in 5.5 ({blacksquare}) or 22 mM (•) glucose for 30 minutes. (B) A Hanes plot calculation of the data was used to determine Km and Vmax. Data are the mean ± SE of results in three experiments, each performed in triplicate.

 


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  		Figure 8. Kinetics of glucose uptake in hRPE cells transduced with GLUT1. The kinetics of glucose uptake was measured in hRPE cells transduced with either a control adenoviral vector AdCMV-ßGAL, (A) or with adenoviral vector containing GLUT1 cDNA (AdCMV-GLUT-1; C). The cells were incubated in 5.5 mM ({blacksquare}) or 22 mM (•) glucose for 30 minutes. A Hanes plot calculation of the data was used to determine Km and Vmax in (B) hRPE-ß-Gal or (D) hRPE-GT1 cells. Data are the mean ± SE of results in three experiments, each performed in triplicate.

 


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  		Figure 1. Time course of glucose uptake in hRPE (A) and hRVE (B) cells. The cells were incubated in 5.5 or 22 mM glucose for 30 minutes. Glucose uptake was measured using [3H]-3-O-MG as a tracer. Data are the mean ± SE of results in three experiments, each performed in triplicate.

 
The specific inhibitor of facilitative glucose transport, cytochalasin B, inhibited glucose uptake in hRPE and hRVE cells with IC50s of 2.5 and 3.2 µM, respectively. Complete inhibition of [3H]-3-O-MG uptake was achieved at 10 µM cytochalasin B in both cell lines.

Uptake of glucose was calculated as follows. The amount of 3-O-MG accumulated (in micromoles) equaled total 3H on the plate after washing (microcuries/plate) divided by the [3H]-3-O-MG specific activity (81 Ci/mmol). The amount of L-glucose nonspecifically associated with the cell layer equals total 14C on the plate after washing (microcuries/plate) divided by the [14C]-L-glucose specific activity (55 mCi/mmol). Because the concentrations of 3-OMG and L-glucose in the uptake solution were the same (40 nM), subtracting the amount of L-glucose from the amount of 3-O-MG gives the specific uptake of 3-O-MG by the cell layer (nmol/plate). The glucose uptake was calculated as 3-O-MG uptake (nmole/plate) multiplied by the concentration of glucose in the uptake solution (mM) divided by the concentration of 3-O-MG in the uptake solution (mM). The glucose uptake was then normalized to total protein on the plate (in nmoles/mg protein) and plotted against time for the time course experiments or against glucose concentration in the uptake solution for determining the kinetics of glucose uptake. Km and Vmax were calculated from a linear regression fit to a Hanes plot (mM glucose/uptake versus mM glucose).

Adenoviral Transduction of hRPE Cells
Recombinant adenoviral vectors that express GLUT1 (AdCMV-GLUT-1) and ß-galactosidase (AdCMV-ßGAL)39 were generously provided by Christopher B. Newgard (University of Texas Southwestern Medical Center, Dallas, TX). AdCMV-GLUT1 and AdCMV-ßGAL were amplified in HEK-293 cells, as described by Becker et al.40 Seventy to 90% confluent hRPE cells were treated with each viral stock for 1 hour, rinsed with PBS, and returned to their regular medium. Transduced cells were then incubated for 48 hours at 37°C in 5% CO2 in a humidified cell culture incubator. The cells were then analyzed by Western blot to confirm GLUT1 overexpression and were used for glucose uptake experiments.

Statistical Analysis
Results are expressed as the mean ± SE of results of at least three experiments performed in triplicate. Statistical significance of differences between experimental groups was determined using Student’s t test.


   Results
Top
 Abstract
 Introduction
 Methods
 Results
Discussion
 References
 
Kinetics of Glucose Uptake in hRVE and hRPE Cells
The kinetics of the initial rate of glucose uptake was measured in hRPE and hRVE cells incubated in media containing 5.5 or 22 mM glucose for 30 minutes. The results of these experiments are shown in Figures 2 and 3 . Km and Vmax were calculated by using a Hanes plot conversion of the kinetics data (Figs. 2B 3B) . Glucose induced an increase in the Vmax of glucose uptake from 7.50 to 14.50 pmol/mg protein per second in hRPE (Figs. 2A 2B) and from 22.40 to 56.00 pmol/mg protein per second in hRVE cells (Figs. 3A 3B) . Km was 5.6 mM in hRPE and 6.3 mM in hRVE incubated in 5.5 mM glucose and was not significantly affected by 22 mM glucose (7.1 mM in hRPE and 8.2 mM in hRVE). Mannitol was used as an osmotic control. Vmax and Km for glucose uptake in hRPE and hRVE cells exposed to 22 mM mannitol containing medium (5.5 mM glucose and 16.5 mM mannitol) for 30 minutes were not significantly different from the control (5.5 mM glucose) cells (data not shown).

Effect of Nocodazole, the Microtubule Assembly Inhibitor, on Glucose-Induced Increase in the Vmax of Glucose Uptake
Nocodazole was used to determine whether translocation is responsible for the observed glucose-induced increase in the Vmax of glucose uptake. Nocodazole at 10 µg/mL significantly inhibited the glucose-induced increase in glucose uptake in hRPE (Fig. 4A) and hRVE (Fig. 4B) cells.



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  		Figure 4. Effect of the microtubule assembly inhibitor nocodazole on glucose uptake. HRPE (A) and hRVE (B) cells were exposed to normal (5.5 mM) or pathophysiological (22 mM) concentrations of glucose in the absence ({square}) or presence () of 10 µg/mL nocodazole for 30 minutes. *P < 0.05 compared with 22 mM in the absence of nocodazole. Data are the mean ± SE of results in three experiments, each performed in triplicate.

 
Effect of Glucose on GLUT1 Expression and Cellular Distribution
The observed increase in the Vmax of glucose uptake must occur through one of the following: an increase in the total expression of the transporter, recruitment of new transporters into the plasma membrane, or activation of the transporters already residing in the plasma membrane. GLUT1 is known to be the predominantly expressed glucose transporter in human BRB.23 41 No other isoforms (GLUT2, -3 and -4) were detected by Northern blot analysis in hRPE and hRVE cells (data not shown). Therefore, we determined the effect of elevated glucose on GLUT1 expression and translocation. No change in GLUT1 mRNA (Fig. 5) and total protein expression (Fig. 6) was shown by Northern and western blot analyses, respectively in hRPE and hRVE cells incubated in media containing 5.5 or 22 mM glucose for 4 to 72 hours (the 48-hour time point is shown in Figs. 5 and 6 ).



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  		Figure 5. GLUT1 mRNA expression in hRPE and hRVE cells. Top: Northern blot analysis of hRPE and hRVE cells cultured in 5.5 to 22 mM glucose for 48 hours. The blots were cohybridized with GLUT1 and ß-actin cDNAs. GLUT1 cDNA hybridized to a single band of an appropriate size (2.8 kb) for the GLUT1 mRNA transcript. Bottom: Quantitation of GLUT1 mRNA expression from Northern blot analysis. The data were normalized to ß-actin mRNA expression. Data are the mean ± SE of results in three experiments.

 


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  		Figure 6. GLUT1 total protein expression in hRPE cells. Top: immunoblot of GLUT1 protein from hRPE cells cultured in 5.5 to 22 mM glucose for 48 hours. The GLUT1 protein appeared as a single band of approximately 55 kDa. Bottom: Quantitation of GLUT1 protein expression. Results represent the mean ± SE of results in three experiments.

 
Cell fractionation studies were performed by differential centrifugation. Figure 7A shows the expression of marker proteins in different fractions. hRPE and hRVE cells incubated in 5.5 or 22 mM glucose-containing medium for 30 minutes showed no difference in GLUT1 expression in plasma membrane-enriched (P16.9) and microsomal (P210) fractions (Fig. 7B) .



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  		Figure 7. GLUT1 protein expression in plasma membrane–enriched (P16.9) and microsomal membrane–enriched fractions (P210). (A) Expression of marker proteins in different fractions. The plasma membrane marker Na,K-ATPase was detected in the P16.9 fraction, the Golgi marker ß1,4-galactosyltransferase in the P210 fraction, and the cytosolic enzyme LDH in the S210 fraction. (B) Immunoblot and quantitation of GLUT1 expression in P16.9 and P210 fractions from hRPE and hRVE cells incubated in 5.5 or 22 mM glucose for 30 minutes. GLUT1 transcript was detected at 55 kDa (glycosylated form45 ) in the P16.9 fraction ({square}) and 45 kDa (nonglycosylated form) in the P210 fraction (). Data are the mean ± SE of results in three experiments. Typical experiments are shown in (A).

 
Thus, an increase in GLUT1 expression and/or translocation was not involved in the glucose-induced increase in the Vmax of glucose uptake observed in hRPE and hRVE cells.

Glucose-Induced Glucose Uptake in Cells Transduced to Overexpress GLUT1
To confirm that an increase in GLUT1 expression is not involved in the mechanism of glucose-induced glucose uptake, we transduced hRPE cells to overexpress GLUT1. The cells were transduced with either control adenoviral vector (AdCMV-ßGAL) or with the vector containing GLUT1 cDNA (AdCMV-GLUT1). GLUT1 overexpression caused the expected increase in the Vmax of glucose uptake from 7.75 pmol/mg protein per second in RPE-ß-Gal cells to 11.67 pmol/mg protein per second in RPE-GT1 cells. Increasing glucose in the medium from 5.5 to 22 mM for 30 minutes caused the same order of magnitude increase in Vmax of glucose uptake in RPE-ß-Gal (Figs. 8A 8B) and RPE-GT1 (Figs. 8C 8D) cells; from 7.75 to 19.01 pmol/mg protein per second and from 11.67 to 27.50 pmol/mg protein per second in RPE-ß-Gal and RPE-GT1 cells, respectively. The Kms for glucose transport in RPE-GT1, RPE-ß-Gal, or nontransduced control cells were similar in both 5.5 and 22 mM glucose.


   Discussion
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
References
 
The effects of hyperglycemia on glucose transport are not well understood.42 Hyperglycemia was reported to decrease or to have no effect on glucose transporter expression in smooth muscle cells,43 44 endothelial cells,43 adipose tissue, and the blood–brain barrier.45 Knowing the adverse effects of glucose in the retina in diabetes, the detailed study of the effect of elevated glucose on glucose uptake in the human BRB was warranted. Studies by Kumagai et al.23 described changes in the expression of glucose transporters in human subjects with long-term diabetes. They demonstrated a virtually identical pattern of GLUT1 immunoreactivity in normal and diabetic subjects with the exception of the absence of GLUT1 from neovascular endothelium. In a separate study, GLUT1 expression in retinal microvasculature from three individuals with long-standing diabetes and minimal or no clinical retinopathy were compared with expression in two without diabetes. In a subpopulation of diabetic microvessels a dramatic increase in GLUT1 expression was noted. These studies, although very important for the understanding the pathologic course of diabetic retinopathy, do not explain how high glucose initially enters the retina in diabetes. The present study is the first to examine short-term effects of pathophysiological levels of glucose on glucose transport in cells comprising the inner (hRVE) and outer (hRPE) human BRB in a well-controlled in vitro setting. Our results demonstrate that the level of glucose typical of poorly controlled diabetes (22 mM) causes an increase in the Vmax of glucose uptake into hRVE and hRPE cells. A member of the facilitative glucose transporter family, GLUT1, is predominantly expressed in the cells comprising the BRB.23 41 46 The Km for glucose uptake by GLUT1 was reported to be 2 to 7 mM,47 indicating that GLUT1-mediated transport should be saturated at 22 mM glucose. The results presented in this article demonstrate that exposure to 22 mM glucose increased the Vmax of glucose uptake in hRPE and hRVE cells without significantly changing the Km of the transport. This increase in the capacity of glucose transport can explain how high levels of glucose initially enter the retina. The increase in the Vmax of glucose uptake could be elicited by a change in glucose transporter expression, translocation into the plasma membrane or activation. We and others have shown that glucose induces an increase in GLUT1 expression in mesangial cells,37 48 and that mesangial cells transduced to overexpress GLUT1 mimic the diabetic phenotype, even when cultured in normal glucose.49 The results of the present study demonstrate that the cells from the retina must use a different mechanism, because hRPE and hRVE cells exposed to high ambient glucose failed to demonstrate upregulation of GLUT1 mRNA and protein expression. Moreover, hRPE cells transduced to overexpress GLUT1 had the same magnitude of glucose-induced increase in the Vmax of glucose uptake as the control cells, confirming that changes in GLUT1 expression are not responsible for the increase in glucose uptake.

We have demonstrated that the microtubule formation inhibitor nocodazole inhibits the glucose-induced increase in glucose uptake in both hRPE and hRVE cells (Fig. 4) . Thus, the mechanism of the increase was dependent on the integrity of microtubules and may require a translocation step. However, GLUT1 translocation was not involved in the mechanism of glucose-induced increase in glucose uptake, because plasma membrane GLUT1 expression did not change in the hRPE and hRVE cells exposed to high versus normal glucose.

The results of this study demonstrated a glucose-induced increase in glucose uptake independent of the expression of glucose transporter. The exact mechanism of glucose-induced activation of glucose uptake is unknown, but there are several possibilities. GLUT1 C-terminal binding protein (GLUT1CBP), which has been recently isolated and characterized,50 51 was shown to bind specifically, through a PDZ domain, to the C terminus of GLUT1. GLUT1CBP is coexpressed with GLUT1 in wide variety of tissues. It interacts with cytoskeletal proteins myosin VI, {alpha}-actin-1, and the kinesin superfamily protein KIF-1B.51 The protein interacts with itself, suggesting that a dimeric form of GLUT1CBP may exist.51 GLUT1CBP may, therefore, be involved in localizing GLUT1 to specific membrane domains, stabilizing GLUT1 by cross-linking GLUT1 monomers and regulating the transport activity by colocalizing GLUT1 with other regulatory proteins. We hypothesize that GLUT1CBP’s binding to GLUT1 C terminus or GLUT1CBP-mediated binding of GLUT1 to as yet unknown regulatory proteins could be involved in the glucose-induced activation of glucose uptake.

A family of novel glucose transporters has been recently discovered. These novel transporters were first proposed to exist based on intriguing data from GLUT4 knockout mice. The mice did not become diabetic, exhibited normal glucose transport in muscle without compensatory increase in GLUT1 or GLUT3, and demonstrated an increase in muscle glucose uptake in response to insulin.52 This observation, along with other suggestive findings, led to the discovery of a novel transporter, GLUT8.53 54 55 Based on similarity to GLUT8, several other transporters from the novel GLUT family were identified.56 57 58 These transporters have approximately 30% homology to the GLUT1 family (GLUT1 to -5). The most well-studied transporter from the novel family is GLUT8. GLUT8 mRNA expression has been demonstrated in a wide variety of human, rat, and mouse tissues, with the highest expression in testis, muscle, brain, liver, kidney, and mouse blastocysts.53 54 55 GLUT8 is regulated by insulin in mouse blastocysts, which translates into increased glucose uptake, a process that is inhibited by antisense oligoprobes.53 GLUT8 was shown to contain a dileucine internalization domain in the amino terminal tail. Mutation of the dileucine domain induced GLUT8 translocation into plasma membrane.55 Expression of GLUT8 in the retina has not been studied. The possible involvement of these novel transporters in the mechanism of glucose-induced activation of glucose uptake by the cells from human BRB remains to be elucidated.


   Acknowledgements
 
The authors thank Seth R. Hootman and Cameron Devarmon Henry for helpful discussions and comments on the manuscript, Caroline A. Greenidge for technical assistance with adenoviral transduction experiments, and Gregory Fink for assistance with statistical analysis of the data.


   Footnotes
 
Supported by a grant from The Center for Clinical Neuroscience and Ophthalmology and the Michigan State University Foundation, Michigan State University, East Lansing, Michigan (DNH).

Submitted for publication August 28, 2001; revised January 16, 2002; accepted January 29, 2002.

Commercial relationships policy: N.

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Corresponding author: Julia V. Busik, Michigan State University, Department of Physiology, 3155 Biomedical and Physical Sciences Building, East Lansing, MI 48824; busik{at}msu.edu.


   References
Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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