1Department of Pathology and 2Departments of Medicine and of Physiology and Biophysics, Case Western Reserve University, Cleveland, Ohio 44106-4951
Submitted 12 February 2003 ; accepted in final form 4 April 2003
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ABSTRACT |
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stomatin; caveolin-1; transferrin receptor; sucrose density fractionation; lipid raft
Results of recent studies in our laboratory suggest that stomatin might be an inhibitory binding partner of Glut1 in Clone 9 cells (49, 50). Stomatin, also known as human erythrocyte protein band 7.2b, is a widely expressed monotopic integral membrane protein (46). Although its function is not clear, it appears to play a role in the regulation of membrane structure, sodium channel activity, and membrane signaling in both mammalian and nematode systems (8, 17, 27, 30, 31, 34, 4144). Recent reports indicate that stomatin is concentrated in cholesterol-rich microdomains ("lipid rafts") in human red blood cells (RBCs) (31, 32). The presence of stomatin in lipid rafts and its interaction with Glut1 suggested that lipid rafts might play an important role in control of Glut1 function.
Lipid rafts represent a revision of the fluid-mosaic model of cell membranes. It has become clear that the heterogeneous mixture of lipids in the plasma membrane and some intracellular membranes form dynamic domains of closely packed lipids that contain specific proteins and high levels of sphingolipids and cholesterol (5, 22, 39). Lipid rafts have been characterized as a class of membrane substructures that include caveolae, which are morphologically distinct structures that are enriched in cholesterol and caveolin (40). Lipid rafts and caveolae are biochemically defined by their lipid content, low density, and resistance to solubilization in buffers containing non-ionic detergents at low temperatures (4°C); hence, they are identified as low-density, detergent-resistant membrane (DRM) domains.
The reported localization of stomatin in DRMs of human RBCs (31, 32), in conjunction with results of our studies suggesting that the association of stomatin with Glut1 correlates with decreased glucose transport (49, 50), prompted us to pose the following set of questions: 1) Is Glut1 also, in part, concentrated in DRMs in Clone 9 cells? 2) Is stomatin localized to the DRM fraction in cells other than the human RBC? 3) Does exposure of cells to inhibitors of oxidative phosphorylation (such as azide) result in a redistribution of stomatin and/or Glut1 between the DRM and non-DRM domains? These and other questions are relevant, especially in light of a recent report indicating that Glut1, but not Glut3, is mostly localized in the DRM fraction in HeLa cells (29). Results of our studies demonstrate that a fraction of cell Glut1 and the bulk of stomatin reside in DRMs under basal conditions and that stimulation of glucose transport in response to azide is associated with a partial redistribution of Glut1, but not of stomatin or caveolin-1, out of the DRM domains.
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MATERIALS AND METHODS |
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Cell culture. Clone 9 cells were maintained in DMEM supplemented with 10% bovine calf serum (vol/vol) and penicillin-streptomycin. Cells were transfected with pcDNA3-human stomatin plasmid (49) or the empty vector, according to the manufacturer's protocol, to enable the detection of stomatin by the available antibody. Stable transfectants were selected with 0.5 mg/ml Geneticin (G418). Surviving clones were harvested using cloning cylinders and maintained in DMEM containing 0.2 mg/ml G418. A clone expressing the human stomatin was employed in these studies. The rate of glucose transport in the chosen clone under basal conditions and following exposure to 5 mM azide was equivalent to nontransfected Clone 9 cells. Culture medium was replaced with fresh medium 1624 h before initiation of experiments. 3T3-L1 fibroblasts were maintained as described (26).
Fractionation by sucrose density step-gradient centrifugation. Four 10-cm plates of confluent Clone 9 cells or 3T3-L1 fibroblasts were washed three times with ice-cold PBS and then scraped into 0.5 ml of ice-cold PBS. All subsequent steps were performed at 4°C. Cell pellets were lysed in 410 µl of buffer A (0.5% Triton X-100, 50 mM HEPES, 150 mM NaCl, 5 mM EDTA, 5 mM EGTA, 20 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 20 mM sodium fluoride, and 0.1 µM PMSF, pH 7.4). Postnuclear lysate (385 µl) was loaded into the bottom of a 4-ml ultracentrifuge tube for the Sorvall TST 60.4 swinging bucket rotor, adjusted to 40% sucrose with 60% sucrose in buffer B (20 mM Tris, 150 mM NaCl, 1 mM EDTA, and 0.1 µM PMSF, pH 7.4) to a total volume of 1 ml, and then overlaid with 2 ml of 35% sucrose and 1 ml 5% sucrose in buffer B. Samples were centrifuged at an average of 150 k g for 17 h at 4°C. Fractions (500 µl) were collected from the top of the tube. Aliquots (30 µl) of each sample were subjected to SDS-PAGE.
In separate gels, samples of whole cell postnuclear lysates from control and azide-treated cells were also used for Western blotting. The gels were transferred to nitrocellulose membranes, blocked with 5% (wt/vol) powdered milk, and then incubated overnight with the appropriate primary antibodies. After three washes with TBST (0.05% Tween 20 in Tris-buffered saline), membranes were incubated with the appropriate horseradish peroxidase-conjugated secondary antibody, washed with TBST, and developed by enhanced chemiluminescence.
In each experiment, samples from fractions 2 to 9 from control or azide-treated cells were prepared for Western blotting on the same membrane. Moreover, in each experiment and for each antigen, the densities of the appropriate bands in fractions 2 and 3 were added and divided by the total of band densities in all the fractions. The Bio-Rad Gel Doc 1000 system was used to determine band density.
One-step separation of soluble and insoluble material. Postnuclear cell lysates were prepared as described above using either Triton X-100, Igepal (NP-40), or 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) (all at 0.5% in buffer A). Lysates were centrifuged at an average of 128 k g for 1 h at 4°C in 500-µl tubes in a Beckman tabletop ultracentrifuge. Supernates were removed and pellets were resuspended in an equal volume of fresh buffer. Equal volumes of the supernate and resuspended pellet were analyzed.
Detergent-free isolation of DRMs. This method was adapted from a previously described protocol (45). Cells were lysed with 0.5 M sodium carbonate (pH 11) in buffer A without detergent. Lysates were homogenized with 20 strokes of a Teflon Potter-Elvehjem homogenizer. Finally, aliquots of the homogenate were sonicated with a probe-type sonicator (550 Dismembranator, Fisher Scientific) using four 2-s sessions while on ice. All subsequent steps were performed at 4°C. This material was processed as described for isolation and analysis of the DRM.
Isolation of plasma membrane DRMs and their fractionation. The method of cell surface biotinylation was adapted from a previously described protocol (37). Confluent cells in 10-cm plates were rinsed three times with cold PBS and then incubated with 3 ml of cold biotinylation buffer (120 mM NaCl, 30 mM NaHCO3, 5 mM KCl, and 0.1 mg/ml sulfo-NHS-SS-Biotin, pH 8.5) for 30 min while rocking at 4°C. After three washes with 5 ml of stop buffer (140 mM NaCl, 20 mM Tris, and 5 mM KCl, pH 7.5), cells were scraped into 0.5 ml of stop buffer. All subsequent steps were performed at 4°C. Cell pellets were lysed in 400 µl of buffer A. The postnuclear lysate was fractionated in the sucrose-density step gradient as described above. Fractions (300 µl) 2, 3, 6, 7, and 8 were incubated with streptavidin-agarose beads overnight at 4°C on a rotisserie shaker; fractions 1, 4, and 5 were excluded because only trace amounts of Glut1 immunoreactivity were present in these samples. The supernatant of each sample was saved, and the beads were washed three times with Tris-buffered saline containing 1 mM EDTA (TBS-EDTA, pH 7.4), and then eluted with 2x Laemmli buffer. Forty percent of the eluate, called the pulldown (pd), and 10% of the supernate (sup) were fractionated by SDS-PAGE and analyzed by immunoblot and enhanced chemiluminescence. Films were scanned, and the density of the bands, measured using the Bio-Rad Gel Doc 1000 system, was used to calculate the content of Glut1 in each sample. The fraction of cell Glut1 present in plasma membrane DRMs was calculated by dividing the sum of Glut1 content in the pulldown of fractions 2 plus 3 by the total amount of Glut1 calculated as the sum of Glut1 content in the pulldown and supernate of all fractions. The ratio of Glut1 in plasma membrane DRMs to total cell Glut1 in control and azide-treated cells was corrected by subtracting the ratio of the same from the mock-biotinylated cells prepared in parallel with each experiment.
Preparation of RBC ghosts. RBC ghosts were prepared from freshly drawn blood as previously described (50). The ghosts were resuspended in buffer A containing 1% Triton X-100 and then subjected to the sucrose density step gradient and the subsequent procedures described above.
Glucose transport. Cytochalasin B-inhibitable glucose transport was measured using 3H-3-O-methylglucose, as previously described (13, 23).
Statistical analysis. All results are expressed as means ± SE. Two-tailed t-test was used, and a P value of <0.05 was considered significant.
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RESULTS |
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We next determined whether the above-noted presence of Glut1 in the DRM fraction is unique to Clone 9 cells. Employing 3T3-L1 fibroblasts and human RBCs, which are known to express Glut1 near exclusively (47, 51, 52), we found Glut1 to be present in DRM fractions of both cells (Fig. 1B for fibroblasts and Fig. 1C for RBCs). The fraction of cell Glut1 localized to the DRM fraction of 3T3-L1 fibroblasts was 29 ± 1%, and for human RBCs, it was 20 ± 3%. In 3T3-L1 fibroblasts, the bulk of caveolin-1 (>95%) was localized in fractions 2 and 3. Similar to the findings in Clone 9 cells, a fraction of Glut1 colocalized with stomatin in DRMs of RBC ghosts; this is presumably the case in 3T3-L1 fibroblasts, although an antibody that recognizes the murine form of stomatin was not available.
The presence of Glut1 in detergent-resistant membrane domains of Clone 9 cells was additionally determined after preparation of cell lysates with buffers containing 0.5% CHAPS or 0.5% NP-40 (Fig. 2). As expected, caveolin-1 remained in the detergent-insoluble fraction, whereas the transferrin receptor was solubilized. Glut1 was present in both fractions employing either of the two additional detergents. However, the fraction of Glut1 remaining in the detergent-resistant fraction (pellet) varied with the detergent used; in two independent experiments employing the above protocol, the percentage of Glut1 in the insoluble fraction averaged 31, 70, and 39% using Triton X-100, CHAPS, and NP-40, respectively. Despite the variability, it is clear that a significant fraction of cell Glut1 is present in the DRM fraction.
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In our attempt to verify the presence of Glut1 in DRMs, we employed a fractionation procedure that does not employ detergents (45). In this method, cells are homogenized in 0.5 M Na2CO3 pH 11 (which will also remove peripheral membrane proteins) followed by fragmentation of membranes by sonication prior to the sucrose-density gradient centrifugation. Employing this procedure and by varying the extent of sonication in several different experiments, we could not obtain ideal conditions that would lead to the expected partitioning of caveolin-1 and the transferrin receptor. We observed that although the amount of Glut1 in the light fractions decreased with increased sonication, caveolin-1 became increasingly associated with high-density fractions (data not shown). We conclude that the carbonate/sonication procedure cannot be applied in a valid manner in these cells. Nevertheless, it appears that a percentage of cellular Glut1 may be present in the low-density fraction.
The observation that a fraction of Glut1 and the bulk of stomatin reside in
DRMs, and considering that stomatin is a Glut1-associated protein that appears
to exert an inhibitory effect on Glut1 function
(49), led us to test the
hypothesis that treatment of cells with azide and the resulting stimulation of
glucose transport is associated with a redistribution of Glut1 and/or stomatin
from the DRM fraction. In keeping with previous results
(13,
36,
37), exposure of Clone 9 cells
to 5 mM azide for 90 min resulted in 4.5 ± 0.8-fold stimulation of
glucose transport (P < 0.05). Upon exposure to azide for 90 min,
we found that the abundance of Glut1 in the low-density fraction decreased,
whereas the distribution of stomatin and caveolin-1 remained unchanged
(Fig. 3A). In repeated
experiments, the content of Glut1 in DRMs of Clone 9 cells decreased by
40% (from 38 to 22% in control and azide-treated cells, respectively),
whereas the fractions of stomatin and caveolin-1 in cell fractions 2
and 3 (DRMs) remained unchanged
(Fig. 3B).
Importantly, the sumtotal of Glut1 content in all the gradient fractions per
unit protein was equal in control and azide-treated cells (P = not
significant). Moreover, and in keeping with previous results
(36,
37), the content of Glut1 in
whole cell lysates was equal in control cells and in Clone 9 cells, exposed to
5 mM azide for 90 min. The distribution of transferrin receptor was unaltered
by treatment of cells with azide.
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To examine the time course of action of azide on Glut1 redistribution and glucose transport, we performed experiments after 30 min of exposure to azide. In three experiments performed after 30 min of exposure to 5 mM azide, the content of Glut1 in the DRM fraction decreased by 25 ± 2% (means ± SE, P < 0.05), whereas the distribution of stomatin remained unaltered; the rate of glucose transport increased 2.5 ± 0.5-fold in cells treated with azide for 30 min (P < 0.05) (Table 1).
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The effect of azide on the abundance of Glut1 in the DRM fraction was also
determined in 3T3-L1 fibroblasts. Similar to the finding in Clone 9 cells, the
content of Glut1 in the low-density fractions decreased by 55% from 29
± 1 to 13 ± 5% in control cells and in cells treated with 5 mM
azide for 2 h, respectively (n = 4; P < 0.05). In
parallel experiments, the rate of glucose transport in fibroblasts increased
2.8 ± 0.5-fold in cells exposed to 5 mM azide for 2 h (P <
0.05).
The above results in Clone 9 cells and 3T3-L1 fibroblasts led us to conclude that treatment of cells with azide, and the resulting stimulation of Glut1-mediated glucose transport, is associated with a net decrease in the content of Glut1 in DRMs. It is, hence, possible that the movement of Glut1 away from the relatively low-membrane fluidity of DRMs (10) and the predicted reduction in the association of Glut1 with stomatin might mediate, in part, the activation (unmasking) of Glut1. This possibility, along with the fact that DRM microdomains are present in both the plasma membrane and intracellular membranes (11), prompted studies on the distribution of Glut1 in plasma membrane DRMs (PM-DRMs) in control and stimulated cells. In these experiments, cells were first surface-biotinylated (37), and after lysis in Triton X-100 and sucrose-density centrifugation, samples were incubated with streptavidin-agarose beads. This protocol yields an estimate of the content of Glut1 in the PMDRM of control and stimulated cells, although the value may represent an underestimate because the biotinylation reaction and isolation procedure may be incomplete. Nevertheless, in three independent experiments, after 90 min of exposure to 5 mM azide, the percentage of total cell Glut1 present in the plasma membrane-DRM fraction decreased by 52 ± 12% (P < 0.05) from 5.9 to 2.8% in control and azide-treated cells, respectively (Fig. 4, A and B).
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DISCUSSION |
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Although the mechanism by which Glut1 is associated with the DRMs is not known, we speculate that palmitoylation may be a potential mechanism. We propose this for several reasons, including 1) palmitoylation is reversible and can occur in the time scale of the observed stimulation of glucose transport in response to azide (2, 21), 2) Glut1 contains an ideally positioned cysteine (Cys 207) that can potentially serve as a substrate for palmitoylation (6, 7, 15), and 3) Glut1 has been reported to be palmitoylated (25). Furthermore, protein acyltransferase activity has been localized to the DRM fraction in some cell types, thereby implicating its compartmentalization as a factor in the recruitment of other proteins to this domain (9).
At present, the signals and sequence of events leading to the movement of Glut1 out of DRMs in response to treatment with azide are not known. Likewise, the role of the redistribution of Glut1 in the observed enhancement of glucose transport following inhibition of oxidative phosphorylation remains unresolved. Nevertheless, on the basis of previous observations that stomatin acts as a Glut1-binding protein and appears to have an inhibitory effect on Glut1 function (49, 50), it is tempting to speculate that the movement of Glut1, but not of stomatin, out of the DRMs in response to azide may contribute to "activation" of Glut1. At the present time, the magnitude of this contribution is not known, especially considering the relative change in the content of Glut1 in DRMs compared with the larger increment in the rate of glucose transport. Further studies are necessary to explore the underlying mechanisms and to determine the role of Glut1 redistribution in the stimulation of glucose transport in response to the inhibition of oxidative phosphorylation.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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