Isolation of the genomic clone consisting of the human angiotensin type 1 receptor promoter and subcloning into pBluescript II KS vector was described previously (Wyse et al., 2000 ). All DNA manipulations were carried out using standard techniques. By using the genomic clone as a template for polymerase chain reaction (PCR) and oligonucleotides corresponding the published sequence of the hAT1R promoter (Guo et al., 1994 ), fragments of varying length were amplified. The oligonucleotide corresponding to exon 1 (+49 to +7 base pairs) of the hAT1R (5′-CCGCCGCGGCCCGGCAGAGCTG-3′) was used as the antisense primer for each reaction. The varying sense primers (hP1, 5′-GAGGCAGGGAGAGGACACAGACC-3′; hP2, 5′-TAATTAATTGATTCCTTAGGGCT-3′; hP3, 5′-GTCCAATTGCCCTCACTAGAACC-3′; hP4, 5′-GAGGAAGTTCCTATTCCTAGTTT-3′; and hP5, 5′-AATCTAATCTTGCTTTCTGGCATC-3′) yielded fragments of 1717, 1543, 1438, 1278, and 800 base pairs, respectively. Each fragment was cloned into the EcoR V site of pBluescript II KS (Stratagene, La Jolla, CA). To excise the promoter fragments from pBluescript, clones were digested with SalI and XbaI. The released fragments were subsequently inserted into the pCAT-Basic expression plasmid. To generate additional deletion fragments between –1717 and –1543 base pairs, PCR was performed using an antisense primer corresponding –750 base pairs and –771 base pairs (5′-GACTATACACCATGGTCAAGTG-3′) spanning the unique NcoI site and varying sense primers (hP2-1, 5′-ATGCGTCGACGTGGTGAGAAGCC-3′; hP2-2, 5′-ATGCGTCGACTTAATTCCATTTGTTG-3′; and hP2-3, 5′-ATGCGTCGACAACTGATTTTTGTATATTG-3′). Each PCR-amplified fragment and hP1-Bluescript (the pBluescript plasmid containing the –1717-base pairs promoter sequence) were restriction digested with NcoI and SalI and unidirectionally subcloned into the NcoI SalI site in the hP1-Bluescript. The resulting promoter fragments of –1653, -1609, and -1579 base pairs in pBluecript were further restriction digested with XbaI and SalI and inserted into basic pCAT expression plasmid. To insert hAT1R glucose response element (GluRE) sequence downstream of the heterologous SV40 promoter, chloramphenicol acetyl transferase (CAT) vector oligonucleotides were synthesized with XbaI and SalI linker sequences to allow unidirectional insertion. The double-stranded GluRE-containing sequence with linkers was generated by annealing two oligonucletides (forward orientation: sense primer, 5′-CTAGAACTGATTTTTGTATATTGATCTTGTATTG-3′; antisense primer, 5′-TCGACAATACAAGATCAATATACAAAAATCAGTT-3′ and reverse orientation: sense primer, 5′-CTAGATTATGTTCTAGTTATATGTTTTT AGTCAAG-3′; antisense primer, 5′-TCGACTTGACTAAAAACATATAACTAGAAC ATAAT-3′) The double-stranded GluRE was restriction digested with XbaI and SalI, and inserted into pCAT-Promoter (downstream of the reporter CAT gene). The authenticity of clones was confirmed by dideoxy sequencing as described previously (Wyse et al., 2000 ).

The hPTEC cells were maintained in a 1:1 mixture of DMEM/Ham's F-12 nutrient mixture (DMEM/F-12) containing 5% fetal bovine serum, 50 units/ml penicillin, and 100 μg/ml streptomycin (complete medium) as described previously (Wyse et al., 2000 ). For transient transfection of DNA constructs, hPTECs were seeded in six-well plates and grown to 70–80% confluence in complete medium. Cells were transiently transfected with 2 μg of reporter plasmids and cotransfected with 2 μg of pSV-β-galactosidase expression construct (to act as an internal control for transfection efficiency) by using the Trans-IT reagent method according to manufacturers instructions (Mirrus, PanVera, Madison, WI) and grown for 24 h in complete medium.

The CAT assays were performed as described previously (Wyse et al., 2000 ). Briefly, transfected cells were growth arrested in glucose- and insulin-free DMEM medium with 1% fetal bovine serum for 24 h and stimulated with normal glucose (5.5 mM) or high glucose (25 mM). After 24 h, cells were rinsed with phosphate-buffered saline (PBS) three times and harvested in the same buffer. Cells were then centrifuged and the resultant pellet was resuspended in 100 μl of 0.25 M Tris-HCl, pH 7.8. Cellular extracts were prepared by bath sonication at 4°C and 10-min centrifugation (17,530 × g at 4°C). Thirty microliters of the supernatant was removed and β-galactosidase activity was measured using a colorimetric assay according to the previously published method (Wyse et al., 2000 ). The remaining supernatant was heated at 70°C for 10 min to inactivate endogenous acetylases and centrifuged further to remove cell debris. The assay for CAT activity was performed as described previously (Wyse et al., 2000 ). The radioactivity was visualized by autoradiography and its intensity of density quantified by image analysis. The results are expressed as normalized values to β-galactosidase activity.

Initially, cells were grown to 80–90% confluence in complete medium containing 5% fetal bovine serum, 50 U/ml penicillin, and 100 μg/ml streptomycin. Twenty-four hour growth-arrested cells in RPMI medium (without glucose) were stimulated with normal or high glucose for indicated times. Alternatively, high glucose-treated cells were primed with cyclohexamide (1 μg/ml) for 15 min. Cells were rinsed two times with PBS, scraped into PBS without Mg²⁺ and Ca²⁺ (pH 7.4), and cytosolic and nuclear extracts were prepared as described previously (Wyse et al., 2000 ). Briefly, harvested cells were centrifuged at 650 × g for 10 min at 4°C, and cell pellets were resuspended in ice-cold hypotonic buffer containing 10 mM HEPES, 10 mM KCl, 1.5 mM MgCl₂, and 0.5 mM dithiothreitol (DTT) supplemented with protease inhibitors (Complete Mini protease inhibitor mixture tablets, EDTA-free; Roche Diagnostics). The cells were Dounce homogenized (14 strokes) and centrifuged at 1000 × g for 10 min at 4°C. The supernatant, cytoplasmic extract was stored at -70°C for later use, and the pellet was resuspended in high-salt buffer containing 20 mM HEPES, 25% glycerol, 400 mM NaCl, and 1 mM EDTA supplemented with protease inhibitors. The high salt nuclear extract was dialyzed (membrane molecular weight cut-off of 8 kDa) overnight in a buffer containing 20 mM Tris-HCl, pH 7.9, 0.1 M KCl, 0.2 mM EDTA, 0.5 mM DTT, 20% glycerol, and 50 μg/ml phenylmethylsulfonyl fluoride (PMSF). Protein concentration was determined using Bio-Rad protein assay reagent based on Bradford method (Bradford, 1976 ), and extracts were stored at -70°C for later use.

Mobility shift assays were performed as described previously (Wyse et al., 2000 ). A double-stranded GluRE-containing sequence with a 5′ overhang was generated by annealing two oligonucleotides (sense primer, 5′-AACTGATTTTTGTATATTGATCTTGTA-3′; antisense primer, 5′-AATACAAGATCAATATACAAAAATCAG-3′) at 65°C. The double-stranded GluRE was labeled by filling the overhang with DNA polymerase Klenow in the presence of [α-³²P]dTTP. The labeled probe was purified using a Sephadex G-25 column. Nuclear extracts were preincubated with 2 μg of poly(dI-dC) in a total volume of 20 μl of binding buffer comprising of 0.1 M Tris-HCl, 50% glycerol, 0.2 M KCl, 0.5 M EDTA, and 1.0 M DTT at 22°C for 20 min. In addition, the reaction mixture was supplemented with either proteinase K, varying concentrations (0–50-fold excess) of double-stranded, unlabeled GluRE; 50-fold excess of nonspecific double-stranded GAGA probe (5′-GAGAGGGAGGAG-3′); or 50-fold excess of unlabeled mutant GluRE (M1, 5′-ACAGAATTTTTGTATATTGATCTTGTATT-3′; M2 5′-AACTGAGGGGTGATAT TGATCTTGTATT-3′; M3, 5′-AACTGATTTTTGGCGCTTGATCTTGTATT-3′; M4, 5′-AACTGATTTTTGTATATTGAGAGGGTATT-3′; M5, 5′-ACAGAAGGGGTGTTATTGATCTTGTATT-3′; M6, 5′-AACTGATTTTTGGCGCTTGAGAGGGTATT-3′ and M7, 5′-AACTGATTTTTGTAGCGGGATCTTGGCGG-3′). Then, the labeled probe (400,000 cpm) was added and the reaction mixture further incubated for 30 min at 22°C. Complexes were separated on a 4% native polyacrylamide gel containing 0.5× Tris borate-EDTA buffer (25 mM Tris, 25 mM boric acid, and 0.5 mM EDTA). The gels were run at 240 V at 4°C, dried, and exposed to Kodak XR-film at -70°C with intensifying screens.

Southwestern analysis was performed as described previously (Wyse et al., 2000 ) by separating nuclear proteins on an 8% SDS-polyacrylamide gel and electrophoretically transferring them to a nitrocellulose membrane. The membrane was rinsed in PBS, prehybridized in buffer containing 10 mM Tris-HCl (pH 8.0), 50 mM KCl, 1 mM EDTA, 1 mM DTT, 5% glycerol, 0.5 mM PMSF, and 500 ng/ml salmon sperm DNA for 30 min at 22°C. Membrane was hybridized in the same buffer containing [α-³²P]dTTP-labeled GluRE probe (2.5 × 10⁶ cpm/10 ml of buffer) for 45 min at 22°C. Nonspecific binding was removed by adding 2 U of DNase I for 15 in at 37°C. The membrane was washed twice in PBS at 22°C and exposed to XR-film at -70°C with intensifying screen.

To further define the region responsible for glucose-mediated repression of CAT activity, we generated additional deletion constructs between –1543- and –1717-base pair sequences (Figure 2A). Reporter gene assays using pCAT-hP2-1 (-1579 base pairs), pCAT-hP2-2 (1609 base pairs), and pCAT-hP2-3 (-1653 base pairs) revealed that the repressor activity resides within the pCAT-hP2-2 construct, suggesting the repressor element is located between –1582 and –1610 base pairs (Figure 2B). Although computer analysis indicated a number of putative binding sites located within this region, the 29-base pair sequence (5′-AACTGATTTTTGTATATTGATCTTGTATT-3′) showed no significant homology with any known cis-regulatory elements. Therefore, we refer to this sequence as the GluRE. To determine whether this region (between -1582 and -1610 base pairs) was capable of binding transcription factors, we performed a series of gel shift assay experiments with nuclear extracts of glucose-exposed hPTECs by using the α-³²P–labeled GluRE as a probe. Gel shift assays showed a distinct mobility shift of the labeled probe (indicated by arrow) with the nuclear extracts (Figure 3A, lane 2), which was abolished on pretreatment of the extracts with proteinase K (Figure 3A, lane 4), indicating the formation of a glucose-induced protein–DNA complex. The protein-DNA binding was specific for nuclear factor(s) as evidenced by observed lack of mobility shift with the cytoplasmic extract (Figure 3A, lane 3). The presence of increasing concentrations (0–50 times) of unlabeled GluRE probe progressively inhibited the appearance of labeled DNA–protein complex (Figure 3B), demonstrating the specificity of the nuclear protein(s) binding to GluRE DNA. This specificity was further confirmed by competitive mobility shift assay with a nonspecific competitor. A previously identified enhancer element (GAGA box) in the hAT1R promoter (Wyse et al., 2000 ) was used as a nonspecific competitor. Although the unlabeled specific GluRE probe (50-fold excess) efficiently competed with the ³²P-labeled GluRE probe, there was no competition observed in binding when the nonspecific GAGA box was used as a competitor (Figure 4). Occasionally, we observed an additional band in the mobility shift assay that could be due to degradation of nuclear binding factors or alterations in binding factor affinity. In an attempt to identify the essential DNA sequences within the GluRE necessary for nuclear trans-acting factor(s) to bind GluRE-DNA, mobility shift assays were performed using nuclear extracts from high glucose-treated cells in the presence of 50-fold excess of various unlabeled mutant GluRE-DNAs. Results in Figure 5 show that although certain mutations within the GluRE have no effect and allowed effective competition with the original sequence, there are three specific regions (“ACTG” motif, and two “TATT” motifs) within the GluRE has high affinity for nuclear protein(s) binding. Sequence alteration of any one region significantly reduced the ability of the GluRE to bind nuclear trans-acting factor(s), suggesting that multiple elements within the hormone response unit are required for full repressor activity. To further define the importance of high glucose in GluRE to bind nuclear trans-acting factor(s), we evaluated time-dependent changes in GluRE binding. Exposure of cells to high glucose caused a time-dependent increase in GluRE binding activity to nuclear trans-acting factor(s) (Figure 6). The maximum binding activity was observed at 4 h (44% increase over basal activity) and maintained up to 24 h after high glucose exposure.

The direct interaction between GluRE and the nuclear transacting protein(s) was further examined by Southwestern analysis. Figure 7 (top) shows that in high glucose, two DNA binding proteins (34 and 36 kDa) showed induced DNA binding activity with the GluRE sequence. Exposure of hPTEC to high glucose for increasing times shows that the 34- and 36-kDa proteins were activated and reached a maximum at 4 h. Furthermore, the induced activity of these binding proteins (BPs) was sustained above basal level for up to 24 h. The delayed activation of these DNA binding proteins by glucose suggests that de novo synthesis may be required for their induced activity. To investigate whether new protein synthesis is required for glucose dependent trans-activation of DNA binding proteins, cells were pretreated with protein synthesis inhibitor cyclohexamide (1 μg/ml; a concentration shown to inhibit ongoing protein synthesis by >98%, as determined by the [³H]leucine incorporation assay (unpublished data), and glucose-induced DNA binding activity was measured (Figure 7, bottom). Our results show that inhibition of protein synthesis did not reduce glucose-induced activity of 34- and 36-kDa proteins to bind the hAT1R GluRE, suggesting that de novo protein synthesis (expression) of the trans-acting factors is not prerequisite for the observed increase in protein DNA binding activity.