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Am J Physiol Cell Physiol 290: C337-C344, 2006. First published September 14, 2005; doi:10.1152/ajpcell.00089.2005
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MEMBRANE TRANSPORTERS, ION CHANNELS, AND PUMPS

Control of Glut1 promoter activity under basal conditions and in response to hyperosmolarity: role of Sp1

Daw-Yang Hwang and Faramarz Ismail-Beigi

Department of Physiology and Biophysics and Department of Medicine, Case Western Reserve University, Cleveland, Ohio

Submitted 1 March 2005 ; accepted in final form 6 September 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We previously identified (Hwang DY and Ismail-Beigi F. Am J Physiol Cell Physiol 281: C1365–C1372, 2001) a 44-bp GC-rich segment of the rat proximal glucose transporter (Glut)1 promoter, located at –104 to –61, as necessary for basal transcription of the Glut1 gene. Using deletion and mutational analysis and expression of transfected reporter constructs, we report in the present study that mutation of the Sp1 site located within this segment of the promoter leads to a marked (~4-fold) decrease in basal promoter activity. Double mutations located in the Sp1 site and in a second downstream GC-rich region (–71 to –51) did not cause a further decrease in promoter activity. Gel shift and supershift assays verified the importance of the Sp1 site. Exposure of cells to trichostatin A resulted in increased expression of the endogenous Glut1 as well as the transfected wild-type construct. Finally, the presence of the Sp1 site was found to be essential for the positive response of the promoter to hyperosmolarity. We conclude that the consensus Sp1 site located in the rat proximal Glut1 promoter is necessary and sufficient for basal expression of the Glut1 gene, as well as for its response to hyperosmolarity.

Glut1 messenger RNA; Sp1; Sp3; electrophoretic mobility shift assay

GLUCOSE TRANSPORTER 1 (Glut1), a member of the Na+-independent facilitative glucose transporter (Glut) family, mediates much of the insulin-independent transport of glucose (15). Although Glut1 is widely expressed, the cellular content of this transporter shows marked tissue-specific expression, with very high levels being present in blood-brain and blood-retinal barriers, placenta, and human erythrocytes (17, 19, 29). Transcriptional regulation of Glut1 gene expression by some stimuli and conditions including hypoxia, inhibition of oxidative phosphorylation, serum and growth factors, and transformation has been studied in some detail, and the roles of specific regions of the promoter and trans-acting factors have been described (2, 4, 5, 9, 14, 21, 24–26, 33). However, the mechanisms controlling the expression of Glut1 gene under basal conditions and in response to hyperosmolarity are less well understood.

In a recent study (14) we identified a 44-bp GC-rich segment within the Glut1 proximal promoter located between –104 and –61 bp upstream of the transcription start site that contains a consensus Sp1 binding site and is necessary for basal transcription of the gene. Moreover, we noted that the 44-bp segment was required for the positive transcriptional response of the gene to hyperosmolarity, because deletion of this GC-rich segment significantly blunted the response to hyperosmolarity. Furthermore, the same Sp1-containing region has been reported to be important for downregulation of Glut1 expression during muscle and heart development (26, 32). A second GC-rich region positioned 20 bp downstream of the above GC-rich segment (but devoid of a consensus Sp1 site) was reported to be crucial to changes in Glut1 gene expression during trophoblast differentiation (21). The importance of the Sp family of transcription factors, first observed in studies of simian virus 40 (SV40) promoter (8), has been established in the regulation of various genes during development, differentiation, cell cycle, and apoptosis (16, 28). Other genes such as Cox-1, p21WAF1/CIP1, and the genes of several enzymes involved in carbohydrate and fatty acid metabolism including pyruvate kinase, aldolase, phosphofructokinase, and acetyl-CoA carboxylase are also regulated by the Sp family of transcription factors (10, 11, 13, 18, 30, 31). Recently, Sp1 activity was shown to be of importance in the development of Huntington disease (7).

To perform a detailed analysis of the importance of these two GC-rich regions for basal activity of the Glut1 promoter and for its response to hyperosmolarity, we used serial deletions and replacement mutations of these GC-rich regions combined with reporter assays under both basal and stimulated conditions. Gel shift and supershift assays were performed to identify proteins that bind to the promoter. Furthermore, the effect of the histone deacetylase (HDAC) inhibitor trichostatin A (TSA) on Glut1 promoter of the transfected plasmid as well as on the endogenous Glut1 gene was examined.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. Clone 9 cells were obtained from the American Type Culture Collection. [{alpha}-32P]dCTP (3,000 Ci/mmol) and [{gamma}-32P]dATP (3,000 Ci/mmol) were purchased from Perkin Elmer Life Sciences. The random primed DNA labeling kit, gel shift assay kit, and Fugene 6 were from Roche Molecular Biochemicals. Nitrocellulose paper (BA-S 85) was obtained from Schleicher & Schuell. The QuikChange site-directed mutagenesis kit and Quickhyb were obtained from Stratagene. DMEM (5.6 mM glucose), Hanks’ balanced salt solution, and calf serum were obtained from Invitrogen. Oligonucleotides used in electromobility gel shift assay- and PCR-based mutations were from Integrated DNA Technologies. The anti-Sp1 and anti-Sp3 antibodies and enhanced chemiluminescence kit were from Santa Cruz Biotechnology. Culture dishes were obtained from Corning. The dual luciferase reporter assay system, pGL2-Basic plasmid, pRL-TK plasmid, Wizard Plus Minipreps system, and restriction endonucleases were purchased from Promega. TSA was from Wako. Horseradish peroxidase-conjugated anti-rabbit antibody and other standard chemicals were obtained from Sigma.

Reporter constructs. Rat Glut1 promoter region was deleted serially from its 5'-end to prepare constructs containing shorter segments of the Glut1 promoter as previously described (14). Reporter construct F (Fig. 1) was prepared by PCR amplification of the specific region of the promoter and then subcloned into pGL2-Basic luciferase reporter vector. Other replacement mutations were made with specific primers and the QuikChange site-directed mutagenesis kit. In all instances, products were verified by sequencing both DNA strands. Plasmid pPL-TK was used as an internal control.



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  		Fig. 1. A GC-rich region is necessary for rat glucose transporter (Glut)1 basal promoter activity. A: nucleotide sequences of Glut1 proximal promoter region from –104 to –51 (A), various 5'-deletions (B–E), and a 6-bp mutation (bold underlined; F) at position –92 to –87. B: Clone 9 cells (in 6-well plate) were transiently transfected with 1 µg of the above plasmids with 50 ng of Renilla luciferase per well as internal control. Cells were harvested 60 h after transfection for measurement of luciferase activity. Firefly luciferase activity of each construct was normalized against Renilla luciferase activity to control for transfection efficiency. Results (in relative luciferase units) represent the average of at least 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with control plasmid F tested using ANOVA.

 
Cell culture, transfection, and luciferase reporter assays. Clone 9 cells were passaged and maintained in DMEM containing 10% calf serum at 37°C with 8% CO2. Cells were used between passages 25 and 37. The medium was changed to serum-free DMEM 16–24 h before all experiments. Culture media were made hyperosmolar by the addition of mannitol at 300 mosmol/kgH2O in DMEM. Cells (in duplicate or triplicate culture dishes) were cotransfected (using Fugene 6 at 70% confluence) with 1 µg of pGL2-Basic constructs expressing firefly luciferase containing various segments of the Glut1 promoter and with 0.05 µg of pRL-TK plasmid (expressing Renilla luciferase). In experiments on hyperosmolarity, media were changed to serum-free isotonic or hypertonic media after 48 h of transfection, and incubation continued for an additional 24 h before measurement of luciferase activity. In Sp1 and Sp3 cotransfection experiments, 1 µg of pCMVSp1, pCMVSp3, or pCMV-null (courtesy of Dr. R. L. Eckert, Case Western Reserve University) was cotransfected with Glut1-luciferase and Renilla luciferase constructs. In experiments designed to measure basal Glut1 promoter activity, the activity of firefly luciferase was corrected against Renilla luciferase activity. In experiments testing the effect of hyperosmolarity, firefly luciferase activity was normalized against the protein content of control and experimental culture plates because Renilla luciferase activity increased dramatically in response to hyperosmolarity.

Electrophoretic mobility shift assay. Nuclear extracts were prepared from Clone 9 cells using the method of Ausubel (1). Protein concentrations of nuclear extracts were determined using the Bio-Rad Dc Protein assay kit. The sequences of various double-stranded (ds) oligonucleotides used for DNA binding and competition experiments are as follows (mutation indicated in boldface). The commercial (Promega) Sp1 consensus oligonucleotide sequence was 5'-ATT CGA TCG GGG CGG GGC GAG-3'. The sequence of the 37-oligonucleotide (probe A), which is identical to the rat Glut1 promoter positioned at –104 to –67 (33), was 5'-TGG TCC TCA GGC CCC GCC CCC CGG CCC ACC TAC ACG C-3', and the sequence of its mutant (probe F) was 5'-TGG TCC TCA GGC AATATT CCC CGG CCC ACC TAC ACG C-3'. ds-Oligonucleotide was prepared by adding equal amounts of the sense and antisense strands in 10 mM NaCl solution, heating for 10 min at 90°C, and gradually cooling to room temperature to anneal the strands. The Gel Shift Assay System (Promega) was used to radiolabel the 5'-end of ds-oligonucleotides. Labeling was carried out at 37°C for 10 min in a mixture containing 4 pmol of ds-DNA, 1 µl of T4 polynucleotide kinase buffer, 1 µl of T4 polynucleotide kinase (Roche Molecular Biochemicals), and 1 µl of [{gamma}-32P]ATP (3,000 Ci/mmol). The reaction was stopped by addition of 1 µl of 0.5 M EDTA and 90 µl of Tris-EDTA buffer. Free unincorporated nucleotides were removed by performing chromatography through a G-25 Quick Spin Column (Roche) using a clinical centrifuge at 1,100 g for 2 min. In gel shift assays, 5 µg of nuclear extract protein was mixed with gel shift binding buffer and incubated for 10 min before addition of 1 µl of 32P-labeled probe (~10,000 cpm) and further incubation for an additional 20 min. Mixtures were loaded on a 4% nondenaturing acrylamide gel that had been prerun for 4 h. After electrophoresis, gels were dried and exposed to X-ray film or analyzed using a PhosphorImager. In supershift experiments, specific antibodies were added to the mixture and incubated for 20 min at room temperature before addition of the radiolabeled DNA probe and incubation for another 10 min.

Northern blot analysis. Cytoplasmic RNA was isolated and fractionated as described previously (14). Blots were probed with rat Glut1 or GAPDH cDNA labeled with [{alpha}-32P]dCTP. Membranes were incubated with probe at 68°C overnight with Quick-Hyb and washed four times for 15 min each with a solution containing 0.1% SDS and 1x SSC at 58°C. Blots were autoradiographed using X-ray film or a PhosphorImager (Molecular Dynamics). Relative intensities of the specific mRNA bands were normalized against the 28S ribosomal RNA band measured by ethidium bromide staining of the membrane or by probing for GAPDH.

Statistical analysis. Results are expressed as means ± SE. One-way ANOVA, followed by Tukey’s test, or, where indicated, an unpaired two-tailed Student’s t-test, was used, and a P value of <0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Role of GC-rich sequence located at –95/–77 of rat Glut1 promoter in control of its basal activity. In a previous study (14) we noted that a 44-bp region of the promoter (from –104 to –61) appeared to be required for both its basal expression and its enhanced expression in response to hyperosmolarity. In accordance with our previous results, we find that Glut1 promoter activity is markedly reduced in the absence of this 44-bp region (compare constructs A and D, Fig. 1). Construct B, in which the consensus Sp1 site located within the 44-bp segment had been deleted, showed a reduction of basal activity; further deletion of the promoter to position –68 (construct C) was associated with even greater lowering of the promoter activity to near-negligible levels. Construct E, in which the promoter was deleted to –11 and was devoid of the TATA box, exhibited an activity that was considered to be "zero" because its activity was similar to that of the pGL2-Basic plasmid, which has no promoter. Construct F, which is similar in length to construct A but contains a 6-bp mutation in the Sp1 site at –92, showed markedly reduced activity, although its basal activity was not completely ablated (similarly to construct C). Analysis of the data by ANOVA showed that all promoter activities were significantly different from construct A (P < 0.05).

Further analysis of this region was carried out using a series of replacement mutations and transient transfection assays. All mutations were made with 4-bp replacement with deoxyadenosine (Fig. 2). Mutation M1 did not exhibit decreased activity (the ~50% increase in luciferase activity of M1 was not significant). Mutations M2, M3, and M4 (encompassing positions –95 to –83) caused a relatively large and significant decrease in the promoter activity to from one-third to one-fourth of the activity of the wild-type construct (plasmid A). Constructs M6 and M8 exhibited a modest (~30%) decrease in promoter activity; such variations probably reflect interaction of cis-elements with other trans-acting factors causing modest functional changes. In contrast, mutations M5, M7, and M9–M12 had no significant effect on promoter activity (Fig. 2B). The decrease in the activity of M2, M3, and M4 was similar to the decrease in the promoter activity of construct F, which contains a 6-bp mutation in the center of the 12-bp GC-rich region encompassing –95 to –83 (Fig. 1B). Hence, the 12-bp GC-rich region in the 44-bp segment, which contains a consensus Sp1 site (labeled 1st GC box in Fig. 2A), appears to be important for the basal activity of rat Glut1 promoter. Importantly, mutations made in another GC-rich region spanning positions –70 to –51 (labeled 2nd GC box in Fig. 2A) located near the 3'-end of the above 44-bp region (M8–M12) had no significant effect on promoter activity.



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  		Fig. 2. Effect of mutations in the GC-rich region of the rat Glut1 promoter on its activity. A: replacement mutations were made with PCR-based mutagenesis using specific primers as described in MATERIALS AND METHODS. The Sp1 consensus sequence at position –92 is bold underlined in construct A. Mutations (M1–M12) are bold underlined and named at right. B: Clone 9 cells were transiently transfected with 1 µg of the above plasmids with 50 ng of Renilla luciferase, and cells were harvested 60 h after transfection for measurement of luciferase activity. Firefly luciferase activity was normalized against Renilla luciferase to control for transfection efficiency. The data represent at least 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with control plasmid A tested using ANOVA.

 
We next tested the effect of double mutations on the promoter activity (Fig. 3). Constructs containing mutations in the region from –75 to –51 in addition to the mutation present in construct F (shown in Fig. 1A) did not further reduce the promoter activity, except for mutant F8 (mutations in –92 to –86 and –63 to –59), which showed an approximately twofold increase in promoter activity compared with mutant F (Fig. 3B). Although the activity of mutant construct F8 was still lower than that of the wild-type promoter, construct A, the increase in the activity of F8 suggests that this section acts as a negative element. Construct F10 showed a slight decrease in promoter activity. All mutants shown in Fig. 3 exhibited a higher level of basal activity (3- to 4-fold) compared with the activity of the promoterless pGL2-Basic plasmid (which measured ~60 relative luciferase units above the assay blank). The above results suggest that the first GC-rich segment positioned at –96 to –83 containing a consensus Sp1 binding site, but not the second downstream GC-rich region, appears to be essential for basal activity of the rat Glut1 promoter.



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  		Fig. 3. Effect of double mutations of the rat Glut1 promoter. A: further mutations (F5–F10) were made in the second GC-rich region (positions –75 to –51) using plasmid F (shown in Fig. 1) as template. B: Clone 9 cells were transiently transfected with 1 µg of each plasmids and 50 ng of Renilla luciferase and were harvested 60 h after transfection for measurement of luciferase activity. Firefly luciferase activity was normalized against Renilla luciferase to control for transfection efficiency. The data represent at least 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with control plasmid F tested using ANOVA.

 
EMSA and Sp1/Sp3 supershift. To identify possible transcription factors that bind to this 44-bp region, EMSA was performed with nuclear extracts from Clone 9 cells; the sequence present in the –104 to –67 region of the rat Glut1 promoter (probe A, Fig. 1A) in conjunction with a 6-bp mutant (–92 to –86) of the same 44-bp region (probe F, Fig. 1A) were used as probes. EMSA detected three DNA-protein complexes after incubation of probe A with nuclear extract, whereas with probe F, in which the 6-bp of the center of the Sp1 binding site had been mutated, the three bands disappeared and a lower (nonspecific) band appeared (Fig. 4A). Use of a commercial Sp1 consensus oligonucleotide (Promega) as a probe also showed a similar three-DNA-protein banding pattern after incubation with nuclear extract from Clone 9 cells (Fig. 4B, lane 2). Inclusion of unlabeled probe A (Fig. 4B, lanes 7 and 8) or Sp1 ds-oligonucleotide (lanes 3–5) resulted in a decrease in the intensity of the three bands with increasing amounts of competitor DNA, signifying specificity.



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  		Fig. 4. EMSA using the GC-rich region of the Glut1 promoter. A: nuclear extracts of Clone 9 cells were incubated with 10,000 cpm of radiolabeled double-stranded (ds) probe A (5'-TGG TCC TCA GGC CCC GCC CCC CGG CCC ACC TAC ACG C-3') or ds-probe F (5'-TGG TCC TCA GGC AAT ATT CCC CGG CCC ACC TAC ACG C-3'). Samples were loaded onto a 4% nondenaturing acrylamide gel and run at 300 V for 2 h. Gels were dried and exposed to film or analyzed using a PhosphorImager. Arrows denote protein-DNA interactions; N.S., nonspecific band. B: gel shift and supershift experiments were performed in the presence of nonradioactive competitor ds-Sp1 DNA (lanes 3–5), nonradioactive competitor ds-probe F (lanes 6–8), anti-Sp1 antibody (lanes 9–11), or anti-Sp3 antibody (lanes 11–14). Thin arrows, protein-DNA interactions; thick arrows, supershift complexes with antibody. Arrows on left denote protein-DNA interaction, and large arrows on right denote supershift bands.

 
Supershift experiments using anti-Sp1 antibody caused a displacement mainly of the upper band (Fig. 4B, lanes 9–11), whereas anti-Sp3 antibody shifted the lower two bands (lanes 12–14). In control experiments, the addition of antibody alone without nuclear extract had no effect on the mobility of the probe (data not shown). Incubation of probe A (but not probe F) with nuclear extract from Clone 9 cells followed by UV irradiation and SDS-PAGE showed one major band at 125 kDa; an identically positioned band was observed when labeled Sp1 probe and recombinant Sp1 protein were used (data not shown). These results suggest that both Sp1 and Sp3 are present in nuclear extracts of Clone 9 cells and that they can bind to the segment of the Glut1 promoter in vitro with requisite specificity. However, the results of these binding assays do not necessarily indicate function.

Effects of Sp1 and Sp3 expression on rat Glut1 promoter. Because mutational analysis, gel shift, and gel supershift assays raised the possibility that Sp1 (and perhaps Sp3) may be playing an important role in Glut1 promoter activity, we tested whether increased Sp1 or Sp3 expression exerts an effect on the activity of the promoter. Construct A or F was cotransfected with plasmid pCMVSp1, pCMVSp3, or pCMV-null to determine the effect of Sp1 and Sp3 overexpression on rat Glut1 promoter. Transient cotransfection with Sp1-expressing plasmid pCMVSp1 significantly increased the promoter activity of the wild-type Glut1 promoter (construct A) by twofold, whereas Sp3-expressing plasmid pCMVSp3 had no effect on either promoter’s activity (Fig. 5). Construct F also responded slightly but significantly to Sp1, but not to Sp3, expression; however, the response of construct F was no greater than that of pGL2-Basic to Sp1 overexpression (data not shown). These results suggested that Sp1 upregulates whereas Sp3 is without effect in modulating Glut1 promoter activity under basal conditions.



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  		Fig. 5. Effect of Sp1 and Sp3 expression on Glut1 promoter activity. Clone 9 cells were transiently cotransfected with 1 µg of construct A (left) or F (right) (shown in Fig. 1), 50 ng of Renilla luciferase, and 1 µg of either Sp1 or Sp3 mammalian expression plasmid per well. Cells were harvested 60 h after transfection for measurement of luciferase activity. Firefly luciferase activity was normalized against protein content of cells. The data represent at least 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with respective control (construct A or F) tested using ANOVA.

 
Effect of TSA and Sp1 binding site on Glut1 promoter activity. To further delineate the role of the Sp1 binding site in the rat Glut1 promoter, we tested the promoter’s response to an agent that modifies chromatin structure by acetylation of histones and thereby helps enhance the interaction of transcription factors with DNA (12, 20). In Clone 9 cells transiently transfected with the wild-type Glut1 promoter (construct A), luciferase activity increased 10-fold after 24 h of incubation with 300 ng/ml TSA, whereas construct F showed <2-fold increase, suggesting that an intact Sp1 binding site is critically important for the response (Fig. 6A). The internal control pRL-TK plasmid (whose promoter contains several Sp1 binding sites) cotransfected in both experiments also showed an ~10-fold increase in Renilla luciferase activity after 24 h of TSA incubation (data not shown).



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  		Fig. 6. Effect of trichostatin A (TSA) on activity of Glut1 promoter construct and on endogenous Glut1 mRNA expression. A: Clone 9 cells were transiently transfected with 1 µg of plasmid A or F and 50 ng of Renilla luciferase. After 48 h, TSA was added to culture medium to a final concentration of 300 ng/ml, and incubation continued for an additional 24 h before measurement of luciferase activity. Because the expression of Renilla luciferase was enhanced 10-fold in response to TSA treatment, firefly luciferase activity was normalized against protein content of cells. The data represent 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with control plasmid A or F, respectively, analyzed using a t-test. B: Clone 9 cells were incubated with 300 ng/ml TSA for times indicated, and cytoplasmic RNA was collected. Lane + contains RNA from cells treated with 250 µM CoCl2 for 12 h. Glut1 mRNA and GAPDH mRNA were probed as described in MATERIALS AND METHODS.

 
To test whether TSA also stimulates Glut1 gene expression, Clone 9 cells were incubated with 300 ng/ml TSA, and RNA was isolated at various intervals (Fig. 6B). Glut1 mRNA content increased after 6 h of incubation; in a repeated experiment the increase at 6 h averaged 2.0 ± 0.2-fold compared with control (P < 0.05). Although the response of the endogenous gene was significantly lower than that of the transfected plasmid, the results suggest that both the endogenous and exogenous forms of the Glut1 promoter respond to TSA.

Response of mutant Glut1 promoter constructs to hyperosmolarity. In a previous study (14), we found that the activity of Glut1 promoter activity under basal conditions and in response to hyperosmolarity is dependent on a 44-bp region of the proximal promoter. Here we tested the response of the wild-type and mutant Glut1 promoter constructs to hyperosmolarity. All three mutants tested (constructs F, M3, and M4) with mutations in the Sp1 consensus sequence showed no significant response to hyperosmolarity, whereas the activity of the wild-type promoter was stimulated (Fig. 7). The slight, nonsignificant increases in luciferase activity of the mutant plasmids were similar to the response of the empty pGL2-Basic plasmid after exposure to hyperosmotic medium. These results indicate that the Sp1 binding site is important not only for basal activity of the rat Glut1 promoter but also for its response to hyperosmolarity.



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  		Fig. 7. Effect of hyperosmolarity on proximal Glut1 promoter activity. Clone 9 cells were transiently transfected with 1 µg of pGL2-Basic, constructs A, F, M3, and M4, and 50 ng of Renilla luciferase as internal control. After 48 h, the medium was changed to either isotonic or hypertonic DMEM, and the incubation continued for an additional 18 h before measurement of dual luciferase activity. Luciferase activity in cells under basal conditions was normalized against Renilla luciferase to control for transfection efficiency. Because expression of Renilla luciferase was enhanced in response to hyperosmolarity, the effect of hyperosmolarity on firefly luciferase activity was normalized against protein content of cells exposed to isotonic and hypertonic medium. The data represent 3 independent experiments performed in triplicate and are means ± SE. *P < 0.05 compared with cells maintained in isotonic medium analyzed using a t-test.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In the present study, we have analyzed the importance of the GC-rich region of the rat Glut1 proximal promoter to its basal activity and for the response of the Glut1 gene to hyperosmotic stimulation. The proximal promoter of rat Glut1 contains three consensus Sp1 sites located at –144, –118, and –92 of the transcription start site (33). In a previous study (2, 14) we found that the 104-bp proximal Glut1 promoter region that included only one of these Sp1 sites, namely, that located at –92, was as active as the 200-bp promoter containing all three Sp1 sites (which was as active as promoters containing up to 6,000 bp of DNA). Using deletion and mutational analysis of Glut1 promoter combined with gel shift and supershift experiments, we now show that the GC-rich Sp1 binding region located –95 to –83 bp upstream of the transcription start site is crucial for basal activity of the promoter. In addition, the Sp1 binding region appeared to be critical for the promoter’s response to hyperosmolarity and TSA. DNA-protein complexes formed during UV cross-linking experiments provided additional support for the role of Sp1 transcription factor in control of the rat Glut1 promoter.

Downregulation of Glut1 expression, associated with decreased tissue Sp1 content, has been observed during development of the heart and differentiation of skeletal muscle in rats, and an inhibitory role of Sp3 has been proposed (9, 26, 32). A GC-rich region exhibiting Sp1 binding activity was identified as a hyperosmotic stress-regulated region in the proximal promoter of serum- and glucocorticoid-inducible protein kinase (Sgk) gene (3). Sp transcription factors also have been shown to be essential for both basal and hypoxia-induced expression of enolase and pyruvate kinase, where hypoxia caused an increase in the Sp1-to-Sp3 ratio (by decreasing Sp3 expression) in C2C12 myocytes without the involvement of the hypoxia-inducible factor pathway (6).

The results of experiments using Sp1 and Sp3 overexpression deserve comment. One explanation for the observed small stimulation of plasmid F in response to Sp1 expression is the presence of unidentified or unoccupied Sp binding sites that are available to the higher levels of Sp factors, as reported previously (21). However, it is unlikely that a similar phenomenon was present in our studies, because plasmids with mutations in both GC-rich regions of the promoter had no significant effect on basal activity. In addition, the 37-bp oligonucleotide containing a mutation in the Sp1 binding site (probe F) did not show the typical Sp1 or Sp3 binding patterns in electromobility shift assays (Fig. 4). Alternatively, it is possible that the small stimulation of construct F (which was not much higher than that of the empty vector) reflects an indirect effect of Sp1 overexpression, because the expression of other putative Glut1-stimulating genes that contain Sp1 binding sites may be stimulated by Sp1 overexpression.

Overexpression of Sp3, in contrast to that of Sp1, resulted in no change in the promoter activity of construct A or F (Fig. 5), whereas the results of the gel shift and supershift assays (Fig. 4B) clearly show that Clone 9 cell nuclei contain both Sp1 and Sp3 proteins. These results suggest that although Sp3 is present, and even though Sp3 can bind to this DNA segment in vitro, the protein does not appear to have a modulatory function in vivo on the Glut1 promoter region under study.

The results of the present experiments using mutational analysis, gel shift assays, and overexpression of Sp1 suggest that Sp1 plays an important stimulatory role in the regulation of Glut1 transcription under both control conditions and hyperosmolarity, whereas Sp3 appears to have no or only a minor modulatory role. Nevertheless, Sp3, with its bifunctional transcription, may have an important role in the regulation of Glut1 gene expression in different cell types and under different conditions (9). Results showing changes in Glut1 promoter activity following mutations in the GC-box and Sp1 and Sp3 overexpression were presented by Zorzano and coworkers (9, 26, 32); other potential binding proteins and regions involved in this regulation remain to be determined. A region from –67 to –80 (C8 box) reported to be inhibitory was stimulated by fetal serum growth factor (25). This location is similar to the second mutation site in our construct F8 with promoter activity close to that of wild-type promoter. The role of Sp1 and Sp3 in Glut1 promoter function requires further elucidation by in vivo studies using chromatin immunoprecipitation and other assays.

It has been suggested that transcriptional activation requires chromatin to become accessible to transcription factors, RNA polymerase complex II, and recruitment of coactivators and corepressors and that histone acetylation plays a key role in the activation process (12, 20). Acetylation of lysine residues at the amino terminus of histones, regulated by histone acetyltransferase and HDAC, results in conformational change in chromatin structure, making it more accessible to the transcriptional machinery. TSA, sodium butyrate, and HDAC inhibitors have been shown to stimulate the transcription of a number of genes by histone hyperacetylation, whereas other mechanisms such as acetylation of other nonhistone proteins or effects on non-HDAC proteins might also be involved (19, 20). Here we tested the role of histone acetylation on rat Glut1 promoter activity by incubation of Clone 9 cells with TSA. TSA resulted in an increase in Glut1 mRNA content, a 10-fold increase in luciferase activity of the plasmid containing the wild-type rat Glut1 promoter, and only a 2-fold increase in the Sp1-mutated Glut1 promoter; these results are consistent with the possibility that the effect of TSA is mediated by, and is dependent on, Sp binding. The discrepancy between the 10-fold increase in plasmid promoter activity and the 2-fold increase in Glut1 mRNA content, a phenomenon described for other genes, might be due to several mechanisms. First, the transfected DNA is believed to be more "open" and accessible to proteins compared with cellular chromatin (27). Second, the transfected construct contains only a small portion of the promoter, whereas the chromosome containing the endogenous Glut1 gene probably contains several negative regulatory elements that might affect the overall response. Finally, it is possible that TSA has a differential effect on Glut1 mRNA vs. luciferase mRNA turnover.

Protein-protein interactions, in addition to DNA-protein interactions, appear to play an important role in the control of Glut1 gene expression. c-Myc has been reported to activate the expression of Glut1 and phosphofructokinase genes (22), an effect that may be mediated through an Sp1-dependent mechanism (18). c-Myc interacts with the carboxy-terminal region of the zinc finger domain of Sp1 or Sp3, suggesting that transcriptional activation by c-Myc involves its binding to Sp1 and Sp3 proteins on promoters of specific genes (10). Moreover, interaction of Sp1/Sp3 factors with other corepressor or coactivator complexes, such as CRSP (cofactor required for Sp1 activation) and TFIID (23), might play a significant role in Glut1 gene expression.

Results showing the importance of the same first GC-rich region of the proximal Glut1 promoter to expression both in the basal state and in response to hyperosmotic challenge leads to a new working hypothesis, namely, that hyperosmolarity exerts its effect on Glut1 gene expression through interaction with Sp1, and perhaps with other coactivators and corepressors (including potentially Sp3). In addition, a conformational change in chromatin containing the Glut1 gene appears to play an important role in the response. We propose that the expression of the Glut1 gene is modulated by interaction of putative proteins with Sp1. In this scenario, Sp1 serves as an anchoring protein. This proposal is consistent with our finding that mutation of the Sp1 binding site disrupted not only the basal activity but also the response of the Glut1 gene to hyperosmolarity. Further studies using in vivo protein-DNA binding assays are necessary to further delineate the response and to identify the set of mediating proteins.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK-45945-A1 supported this research.


   FOOTNOTES
 

Address for reprint requests and other correspondence: F. Ismail-Beigi, Clinical and Molecular Endocrinology, Case Western Reserve Univ., Cleveland, OH 44106-4951 (e-mail: fxi2{at}po.cwru.edu)

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Ausubel FM. Current Protocols in Molecular Biology. New York: Wiley Interscience, 1987.

2. Behrooz A and Ismail-Beigi F. Dual control of glut1 glucose transporter gene expression by hypoxia and by inhibition of oxidative phosphorylation. J Biol Chem 272: 5555–5562, 1997.[Abstract/Free Full Text]

3. Bell LM, Leong ML, Kim B, Wang E, Park J, Hemmings BA, and Firestone GL. Hyperosmotic stress stimulates promoter activity and regulates cellular utilization of the serum- and glucocorticoid-inducible protein kinase (Sgk) by a p38 MAPK-dependent pathway. J Biol Chem 275: 25262–25272, 2000.[Abstract/Free Full Text]

4. Boado RJ and Pardridge WM. Glucose deprivation and hypoxia increase the expression of the GLUT1 glucose transporter via a specific mRNA cis-acting regulatory element. J Neurochem 80: 552–554, 2002.[CrossRef][ISI][Medline]

5. Chen C, Pore N, Behrooz A, Ismail-Beigi F, and Maity A. Regulation of glut1 mRNA by hypoxia-inducible factor-1. Interaction between H-ras and hypoxia. J Biol Chem 276: 9519–9525, 2001.[Abstract/Free Full Text]

6. Discher DJ, Bishopric NH, Wu X, Peterson CA, and Webster KA. Hypoxia regulates {beta}-enolase and pyruvate kinase-M promoters by modulating Sp1/Sp3 binding to a conserved GC element. J Biol Chem 273: 26087–26093, 1998.[Abstract/Free Full Text]

7. Dunah AW, Jeong H, Griffin A, Kim YM, Standaert DG, Hersch SM, Mouradian MM, Young AB, Tanese N, and Krainc D. Sp1 and TAFII130 transcriptional activity disrupted in early Huntington’s disease. Science 296: 2238–2243, 2002.[Abstract/Free Full Text]

8. Dynan WS and Tjian R. Isolation of transcription factors that discriminate between different promoters recognized by RNA polymerase II. Cell 32: 669–680, 1983.[CrossRef][ISI][Medline]

9. Fandos C, Sanchez-Feutrie M, Santalucia T, Vinals F, Cadefau J, Guma A, Cusso R, Kaliman P, Canicio J, Palacin M, and Zorzano A. GLUT1 glucose transporter gene transcription is repressed by Sp3. Evidence for a regulatory role of Sp3 during myogenesis. J Mol Biol 294: 103–119, 1999.[CrossRef][ISI][Medline]

10. Gartel AL, Ye X, Goufman E, Shianov P, Hay N, Najmabadi F, and Tyner AL. Myc represses the p21WAF1/CIP1 promoter and interacts with Sp1/Sp3. Proc Natl Acad Sci USA 98: 4510–4515, 2001.[Abstract/Free Full Text]

11. Girard J, Ferre P, and Foufelle F. Mechanisms by which carbohydrates regulate expression of genes for glycolytic and lipogenic enzymes. Annu Rev Nutr 17: 325–352, 1997.[CrossRef][ISI][Medline]

12. Grunstein M. Histone acetylation in chromatin structure and transcription. Nature 389: 349–352, 1997.[CrossRef][Medline]

13. Hamm-Kunzelmann B, Schafer D, Weigert C, and Brand K. Redox-regulated expression of glycolytic enzymes in resting and proliferating rat thymocytes. FEBS Lett 403: 87–90, 1997.[CrossRef][ISI][Medline]

14. Hwang DY and Ismail-Beigi F. Stimulation of GLUT-1 glucose transporter expression in response to hyperosmolarity. Am J Physiol Cell Physiol 281: C1365–C1372, 2001.[Abstract/Free Full Text]

15. Joost HG, Bell GI, Best JD, Birnbaum MJ, Charron MJ, Chen YT, Doege H, James DE, Lodish HF, Moley KH, Moley JF, Mueckler M, Rogers S, Schurmann A, Seino S, and Thorens B. Nomenclature of the GLUT/SLC2A family of sugar/polyol transport facilitators. Am J Physiol Endocrinol Metab 282: E974–E976, 2002.[Abstract/Free Full Text]

16. Kaczynski J, Cook T, and Urrutia R. Sp1- and Krüppel-like transcription factors. Genome Biol 4: 206, 2003.[CrossRef][Medline]

17. Kumagai AK, Glasgow BJ, and Pardridge WM. GLUT1 glucose transporter expression in the diabetic and nondiabetic human eye. Invest Ophthalmol Vis Sci 35: 2887–2894, 1994.[Abstract/Free Full Text]

18. Le HB, Vaisanen PA, Johnson JL, Raney AK, and McLachlan A. Regulation of transcription from the human muscle phosphofructokinase P2 promoter by the Sp1 transcription factor. DNA Cell Biol 13: 473–485, 1994.[ISI][Medline]

19. Maher F, Vannucci SJ, and Simpson IA. Glucose transporter proteins in brain. FASEB J 8: 1003–1011, 1994.[Abstract]

20. Naar AM, Lemon BD, and Tjian R. Transcriptional coactivator complexes. Annu Rev Biochem 70: 475–501, 2001.[CrossRef][ISI][Medline]

21. Okamoto Y, Sakata M, Yamamoto T, Nishio Y, Adachi K, Ogura K, Yamaguchi M, Takeda T, Tasaka K, and Murata Y. Involvement of nuclear transcription factor Sp1 in regulating glucose transporter-1 gene expression during rat trophoblast differentiation. Biochem Biophys Res Commun 288: 940–948, 2001.[CrossRef][ISI][Medline]

22. Osthus RC, Shim H, Kim S, Li Q, Reddy R, Mukherjee M, Xu Y, Wonsey D, Lee LA, and Dang CV. Deregulation of glucose transporter 1 and glycolytic gene expression by c-Myc. J Biol Chem 275: 21797–21800, 2000.[Abstract/Free Full Text]

23. Ryu S, Zhou S, Ladurner AG, and Tjian R. The transcriptional cofactor complex CRSP is required for activity of the enhancer-binding protein Sp1. Nature 397: 446–450, 1999.[CrossRef][Medline]

24. Sanchez-Feutrie M, Santalucia T, Fandos C, Noe V, Vinals F, Brand NJ, Ciudad CJ, Palacin M, and Zorzano A. A novel muscle DNA-binding activity in the GLUT1 promoter. Cell Mol Life Sci 61: 709–720, 2004.[CrossRef][ISI][Medline]

25. Sanchez-Feutrie M, Vinals F, Palacin M, and Zorzano A. Identification of a novel proliferation-dependent C-rich element that mediates inhibition of the rat GLUT1 promoter. Gene 322: 47–55, 2003.[CrossRef][ISI][Medline]

26. Santalucia T, Boheler KR, Brand NJ, Sahye U, Fandos C, Vinals F, Ferre J, Testar X, Palacin M, and Zorzano A. Factors involved in GLUT-1 glucose transporter gene transcription in cardiac muscle. J Biol Chem 274: 17626–17634, 1999.[Abstract/Free Full Text]

27. Smith CL and Hager GL. Transcriptional regulation of mammalian genes in vivo. A tale of two templates. J Biol Chem 272: 27493–27496, 1997.[Free Full Text]

28. Suske G. The Sp-family of transcription factors. Gene 238: 291–300, 1999.[CrossRef][ISI][Medline]

29. Takata K and Hirano H. Mechanism of glucose transport across the human and rat placental barrier: a review. Microsc Res Tech 38: 145–152, 1997.[CrossRef][ISI][Medline]

30. Taniura S, Kamitani H, Watanabe T, and Eling TE. Transcriptional regulation of cyclooxygenase-1 by histone deacetylase inhibitors in normal human astrocyte cells. J Biol Chem 277: 16823–16830, 2002.[Abstract/Free Full Text]

31. Vaulont S, Vasseur-Cognet M, and Kahn A. Glucose regulation of gene transcription. J Biol Chem 275: 31555–31558, 2000.[Free Full Text]

32. Vinals F, Fandos C, Santalucia T, Ferre J, Testar X, Palacin M, and Zorzano A. Myogenesis and MyoD down-regulate Sp1. A mechanism for the repression of GLUT1 during muscle cell differentiation. J Biol Chem 272: 12913–12921, 1997.[Abstract/Free Full Text]

33. Williams SA and Birnbaum MJ. The rat facilitated glucose transporter gene. Transformation and serum-stimulated transcription initiation from identical sites. J Biol Chem 263: 19513–19518, 1988.[Abstract/Free Full Text]




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