The current study examined angiotensin receptor (ATR) regulation in proliferating rat aortic vascular smooth muscle cells (VSMCs) in culture. Radioligand competition analysis coupled with RNase protection assays (RPAs) revealed that angiotensin type 1a receptor (AT1aR) densities (Bmax) increased by 30% between 5 and 7 days in culture [Bmax (fmol/mg protein): day 5, 379 ± 8.4 vs. day 7, 481 ± 12, n = 3, P < 0.05] under conditions in which no significant changes in AT1aR mRNA expression occurred [in RPA arbitrary units (AU): day 5, 0.23 ± 0.01 vs. day 7, 0.24 ± 0.04, n = 4] or in mRNA synthesis determined by nuclear run-on assays [AU: day 5, 0.35 ± 0.14 vs. day 7, 0.33 ± 0.11, n = 5]. In contrast, polysome distribution analysis indicated that AT1aR mRNA was more efficiently translated in day 7 cells compared with day 5 [% of AT1aR mRNA in fraction 2 out of total AT1R mRNA recovered from the sucrose gradient: day 5, 20.9 ± 9.9 vs. day 7, 56.8 ± 5.6, n = 3, P < 0.001]. Accompanying the polysome shift was 50% less RNA-protein complex (RPC) formation between VSMC cytosolic RNA binding proteins in day 7 cells compared with 5-day cultures and the 5′ leader sequence (5′LS) of the AT1aR [5′LS RPC (AU): day 5, 0.62 ± 0.15 vs. day 7, 0.23 ± 0.03; n = 4, P < 0.05] and also with exon 2 [Exon 2 RPC (AU): day 5, 35.0 ± 5.7 vs. day 7, 17.2 ± 3.6; n = 4, P < 0.05]. Taken together, these results suggest that AT1aR expression is regulated by translation during VSMC proliferation in part by RNA binding proteins that interact within exon 2 in the 5′LS of the AT1aR mRNA.
- angiotensin receptor
- RNA binding proteins
ang ii, the effector molecule of the renin-angiotensin system (RAS), plays an integral role in maintenance of blood pressure and fluid and electrolyte homeostasis by increasing arterial vasoconstriction, inducing thirst, stimulating aldosterone secretion, and increasing renal sodium reabsorption (10). ANG II is also a potent growth factor promoting the growth and migration of vascular smooth muscle cells (VSMCs), both in vivo and in vitro. Proliferation of VSMCs is a hallmark of both wall thickening and plaque formation, and vascular structural changes resulting in increased vasoconstriction and decreased vasodilatation can lead to hypertension and associated cardiovascular and renal disease (12, 36, 39).
ANG II mediates many of its physiological effects by binding to the ANG II type 1 receptor (AT1R) (35). ANG II is implicated in the pathogenesis of multiple disease states, and blockade of AT1Rs is widely used clinically to treat various forms of hypertension, atherosclerosis, and cardiovascular and renal disease (7, 26, 28). AT1R antagonists also attenuate VSMC proliferation and migration in vitro and intimal thickening, neointimal formation, and hyperplasia in in vivo models of vascular injury (12, 16, 19, 23).
Given the role of AT1Rs in these physiological and pathophysiological processes, it is no surprise that AT1R expression is controlled at several levels, including during RNA synthesis and processing (6, 11, 14), RNA stability and translation (13, 20, 40, 41), and through posttranslational mechanisms such as receptor desensitization, endocytosis, and trafficking (21, 33).
Perhaps least understood are the posttranscriptional mechanisms that regulate AT1R expression and activity. We have previously shown that AT1Rs can be regulated at the level of translation in stably transfected Chinese hamster ovary (CHO) cells and in transiently transfected A10 rat aortic smooth muscle cells (13, 41). Previous studies have demonstrated that there are alterations in the expression of the AT1R in culture over time. For example, Gunther et al. (9) demonstrated that 125I-labeled ANG II binding is significantly increased in rat mesenteric artery smooth muscle cells after several days in culture compared with freshly isolated cells. In this paper, we describe a similar increase in AT1R expression in proliferating aortic VSMCs in culture and investigate the hypothesis that translational regulation of AT1R mRNA contributes to regulation of the receptor under these conditions.
MATERIALS AND METHODS
Rat thoracic aortic VSMC were prepared from Fischer 344 rats following an enzyme dissociation method previously described (1). VSMCs were grown in a mixture of Dulbecco's modified Eagle's and Ham's F12 media (DMEM-F12) (Biofluids, Rockville, MD) containing 10% FCS, penicillin (100 U/ml) and streptomycin (100 μg/ml), at 37°C in a humidified atmosphere of 95% air-5% CO2. Cell viability was assessed by trypan blue exclusion or lactate dehydrogenase assay (Roche Molecular Biochemicals, Temecula, CA). Passages of cells between 3 and 25 were used in all subsequent experiments. Cells were plated at a density of 1 × 104 cells/cm2 and grown for 3–7 days. Cells reached confluency by day 7.
Radioligand binding assay.
The cells were washed in PBS, resuspended in Buffer A [10 mM Tris buffer, 0.32 M sucrose, 2 mM EDTA, and 3 mM MgCl2, pH 7.2], and homogenized. The homogenate was centrifuged at 1,000 g for 5 min, and the resultant supernatant was centrifuged at 44,000 g for a further 65 min. The pellet was resuspended in Buffer B [10 mM Tris containing, 3 mM MgCl2, pH 7.2]. The protein content was determined using the Bio-Rad protein assay (Bio-Rad, Hercules, CA). The DNA content was determined by the DNAzol Method (Molecular Research Center, Cincinnati, OH).
For equilibrium binding, membrane fractions were incubated with increasing concentrations (0.05–4 nM) of 125I-labeled [Sar1Ile8ANG II (Peptide Radioiodination Center, Washington University, Pullman, WA) in Buffer B supplemented with 0.2% BSA, for 1 h at room temperature. Nonspecific binding was determined in the presence of 1 μM unlabeled [Sar1Val5Ala8]ANG II (Saralasin) (Sigma-Aldrich, St., Louis, MO). For competition-binding experiments, membrane fractions were incubated with increasing concentrations (0.3–0.5 nM) of Saralasin (AT1R and AT2R antagonist), SK1080 (AT1R antagonist, Korean Research Institute of Chemical Technology, Dae Jeon, South Korea) or PD123319 (AT2R antagonist, Park-Davis Research Laboratories, Ann Arbor, MI). Binding reactions were terminated by addition of ice-cold buffer B and rapid filtration through glass fiber filters with a Brandel cell harvester (Brandel M-24R). The filters were washed with ice-cold Buffer B, and the membrane-bound radioactivity counted in a Packard Cobra II γ-counter. Results are expressed as specific binding (defined as total minus nonspecific binding). The dissociation constant (Kd) and maximum number of specific binding sites (Bmax) were estimated on the basis of nonlinear regression analysis using the software program Prism v4.0 (GraphPad Software, San Diego, CA).
Cells were washed with 10 ml of ice-cold PBS and trypsinized by treatment with 5 ml 0.05% trypsin/0.02% Versene. The cells were recovered in 10 ml of DMEM-F12 containing 10% FBS, then centrifuged at 600 g for 5 min, and the resulting pellet was resuspended in 10 ml PBS. Cells were counted using a hemocytometer, and cell viability was assessed by trypan blue exclusion or lactate dehydrogenase assay (Roche Molecular Biochemicals).
Ribonuclease protection assay.
AT1R mRNA in VSMC was measured by ribonuclease protection assay (RPA) using RPA III (Ambion, Austin, TX), according to the manufacturer's protocol. The specific AT1aR cRNA probe (20,000 cpm), which produces a protected fragment of 95 bases was hybridized with total RNA (10 μg, isolated using Tri reagent, Molecular Research Center, Cincinnati, OH) from each cell preparation in a total volume of 10 μl at 42°C for 16 h. For internal control, pTRI-Actin (mouse antisense control template, Ambion) was used to make a 304 base cRNA probe, which produces a protected fragment of 245 bases with T7 RNA polymerase and γ-32P-labeled CTP by in vitro transcription. RNase digestion with RNase T1 was carried out at 37°C for 30 min. After the precipitation of protected fragments, the samples were separated by electrophoresis on a denaturing 5% polyacrylamide gel. After electrophoresis, the gel was transferred to filter paper, dried, and exposed to Phosphor imager screen; the total abundance of AT1R mRNA was quantitated by ImageQuant software (IQMac V1.2) after the image was scanned (Molecular Dynamics Storm Phosphorimager, Amersham Biosciences, Piscataway, NJ). The expression level of AT1R mRNA was expressed as the ratio of the signal in AT1aR cRNA to β-actin.
Nuclear run-on assays.
Nuclei were prepared from VSMCs (1–3 × 107 cells) as described by Nickenig and Murphy (24). In brief, VSMCs were treated with lysis buffer [0.5% Nonidet P-40 in 150 mM KCl, 4 mM magnesium acetate, 10 mM Tris·HCl pH 7.4]. The nuclei were isolated by centrifugation through 0.6 M sucrose then stored at −80°C in glycerol until use in transcription assays. Transcription reactions were started by incubating nuclei for 30 min at 30°C in the presence of dithiothreitol (DTT), ATP, GTP, CTP, and α-[32P]UTP. Reactions were terminated by the addition of TRI reagent (Molecular Research Center). The radioactive RNA was isolated and purified using a P-30 spin column (Bio-Rad) followed by hybridization to nylon membranes immobilized with AT1aR or GAPDH in the pCR3 plasmid. After washing the membranes, the filters were exposed to Phosphor imager screens. De novo AT1R mRNA synthesis levels were expressed as the ratio of the AT1aR to GAPDH signals.
Polysomal distribution analysis.
Cells were homogenized in a buffer containing 20 mM Tris·HCl, pH 7.5, 100 mM NaCl, 1.5 mM MgCl2, 10 mM EGTA, 500 μg/ml heparin, 0.5% Triton X-100, 100 μg/ml cycloheximide, 0.5% deoxycholic acid and 160 U/ml of RNasin inhibitor. After centrifugation at 12,000 g for 10 min at 4°C, the supernatant was loaded onto a 10–50% linear sucrose gradient and centrifuged at 243,000 g at 4°C for 2 h. In the polysome disruption experiment, the supernatant was loaded onto a sucrose gradient prepared in buffer devoid of MgCl2 and supplemented with 20 mM EDTA. Six equal volume fractions were collected from the bottom to the top of the sucrose gradient, digested with 500 μg/ml pronase, and the RNA was isolated by phenol-chloroform extraction and quantitated using the DNAzol Method (Molecular Research Center).
The amount of AT1R mRNA in each fraction was determined by real-time PCR. First-strand cDNA was prepared from total RNA using the Superscript preamplification system (Invitrogen, Carlsbad, CA) with random hexamers. Quantitations of specific mRNAs and 18S rRNA (for control) were performed by real-time PCR using the ABI Prism 7700 Sequence Detection System (Perkin Elmer Applied Biosystems, Foster City, CA). The PCR reaction mixture consisted of RNase-free water, TaqMan Universal PCR Master Mix (Perkin Elmer Applied Biosystems), DNA samples, and 300-nM specific primers and 10 μM probe: [Forward primers: 119F (E1,2,3), 5′-CCA CAT TCC CTG AGT TAA CAT ATG A-3′ and 114F (E1,3), 5′-CTC TGC CAC ATT CCC TGG TC-3′; Reverse primer: 310R (E1,2,3 and E1,3), 5′-TCT TTT GAT ACC ATC TTC AGC AGA A-3′; and Probe: 232T (E1,2,3 and E1,3), 6 FAM-TCG AAT AGT GTC TGA GAC CAA CTC AAC CCA-TAMRA]. PCR conditions were optimized for the probe (232T) and both sets of primers [E1,2,3: 119F and 310R and E1,3: 114F and 310R] using E1,2,3 and E1,3 plasmid DNA for standards. The 18S primers were purchased from Perkin Elmer Applied Biosystems.
The expression of E1,3 and E1,2,3 mRNA and 18S rRNA in each sample was quantitated using the respective set of primers. PCR reactions without reverse transcription were included to control for contamination by genomic DNA. The standard curves for 18S rRNA and mRNA were made by a series of five dilutions (53, 54, 55, 56, 57, and 58) of the cDNA. The standard curves were calculated based on the control values.
Cells were homogenized in 25 mM Tris buffer, pH 7.4, containing 0.1 mM EDTA, 40 mM KCl, 1% Triton X-100, 0.1 mM phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, 0.2 U/ml aprotinin, and 10 μg/ml antipain. The homogenate was centrifuged at 300 g for 10 min at 4°C, and the supernatant was centrifuged at 20,000 g for a further 20 min at 4°C. The final supernatant was layered onto a 30% sucrose cushion solution containing 10 mM Tris (pH 7.6), 1 mM potassium acetate, 1.5 mM magnesium acetate, and 2 mM DTT, then centrifuged at 230,000 g for 3 h at 4°C. The protein concentration was determined using the Bio-Rad protein assay.
The 5′ leader sequence (5′LS) and exon 2 of the rat AT1aR cDNA were subcloned into the pCR3 vector (Invitrogen) by TA cloning. The plasmids were linearized with Xho1 and then T7 RNA polymerase was used for in vitro transcription with γ-32P-labeled GTP (Promega, Madison, WI). After electrophoresis in an 8 M urea-5% polyacrylamide gel, the probes were eluted and the cRNAs were precipitated.
Cytosolic extracts (30 μg) were incubated with 100 mM DTT, 40 U RNase inhibitor, 10× binding buffer [100 mM N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid, 400 mM KCl, 30 mM MgCl2 and 50% glycerol] and 105 cpm of [32P]-labeled-cRNA probe for 20 min at 30°C. T1 RNase (1 U/μl) and heparin sulfate (100 μg/μl) were then added and the RNA separated on a 4% polyacrylamide gel at 200 V. The bands were quantitated by phosphorimaging using ImageQuant software (IQMac v1.2, GE Healthcare Biosciences, Little Chalfont, UK).
Data are expressed as the means ± se and analyzed using one-way ANOVA followed by the Student-Neuman-Keuls multiple comparison test, two-way ANOVA, followed by Bonferroni's post hoc test, or the unpaired Student's t-test. P < 0.05 was considered to be statistically significant.
AT1R binding in VSMC.
After initial plating at 1 × 104/cm2, the density of cultured rat aortic VSMC steadily increased by 2.7-fold on day 3, 7.8-fold on day 5, and 20-fold on day 7 (Fig. 1A). In these cells, specific binding of the angiotensin receptor ligand 125I-labeled [Sar1Ile8]ANG II increased from day 3 (cpm, 270.5 ± 47.5) to day 7 (cpm, 1,809 ± 221) by 6.7-fold (Fig. 1B). The increase in specific binding was associated with constant total cell protein per cell (Fig. 1C) and DNA content per cell (Fig. 1D). Because of the limited quantities of cells 3 days after passage, we focused the rest of the experiments on cells cultured for 5 and 7 days.
Membranes were prepared from cells cultured on days 5 and 7 and saturation binding curves were performed (Fig. 2A). Scatchard transformation of the data (Fig. 2A, inset) revealed a linear distribution. No distinguishable effects on the ligand binding affinity (Kd) were observed over the culture period [Kd (nM): day 5, 0.14 ± 0.02 vs. day 7, 0.15 ± 0.02 nM, n = 3], whereas the AT1R density significantly increased by 30% from day 5 to day 7 [Bmax (fmol/mg protein): day 5, 379 ± 8.4 vs. day 7, 481 ± 12, n = 3, P < 0.05].
Radioligand competition experiments were also performed. Membranes prepared from VSMCs cultured for 5 days were incubated with 125I-labeled [Sar1Ile8]ANG II and the AT1R-specific nonpeptide antagonist, SK1080, or the AT2R-specific nonpeptide antagonist, PD123319 (Fig. 2B). PD123319 was a poor competitor of 125I-labeled [Sar1 Ile8] ANG II binding (Ki > 10 μM). In contrast, SK1080 completely and potently abolished the 125I-labeled[Sar1Ile8]ANG II binding (Ki = 3.9 nM). Identical results were found in competition studies in membranes from VSMC cultured for 7 days (data not shown).
AT1R mRNA expression.
The levels of AT1R mRNA expression in VSMC were determined by an RNA protection assay (RPA) that we previously developed, which differentiates between the AT1aR and AT1bR mRNA subtypes (38) (Fig. 3A). The levels of AT1aR mRNA expression when normalized to β-actin were not significantly different between days 5 and 7 [Arbitrary units (AU): day 5, 0.23 ± 0.01 vs. day 7, 0.24 ± 0.04, n = 4] (Fig. 3B). No AT1bR mRNA was detected in cultured VSMC by RPA. When measured by quantitative real-time PCR, expression of AT1bR mRNA was found to be very low. Levels of AT1aR mRNA were 38-fold higher than AT1bR mRNA (AT1aR mRNA, 4.405 ± 0.450 pg/μg total RNA vs. AT1bR mRNA, 0.115 ± 0.037 pg/μg total RNA, n = 4). We also performed nuclear run-on assays and did not detect significant de novo AT1R mRNA synthesis between days 5 and 7 [AU: day 5, 0.35 ± 0.14 vs. day 7, 0.33 ± 0.11, n = 5].
The AT1aR gene is composed of three exons, which are alternatively spliced to generate two AT1aR mRNA splice variants differing only in the presence (E1,2,3) or absence (E1,3) of exon 2. Real-time PCR was used to determine the levels of E1,3 and E1,2,3 mRNA in VSMC. Greater than 75% of the total AT1aR mRNA expressed in VSMC is composed of the E1,2,3 splice variant [AT1R mRNA (pg/μg total cellular RNA): E1,3, 1.2 ± 0.2; E1,2,3, 4.0 ± 0.6, n = 3]. No significant differences in splice variant levels or in the splice variant ratios were observed between VSMC cultured for 5 and 7 days (data not shown).
Polysome distribution of AT1aR mRNA.
Cytosolic extracts of VSMC cultured for 5 or 7 days were layered onto sucrose gradients for polysome distribution analysis. Six fractions/gradient were collected on the basis of the heaviest (fraction 1) to the lightest (fraction 6) densities. Quantitation of AT1aR mRNA expression by real-time PCR in each fraction revealed that the AT1aR mRNA distribution shifted from the lighter polysome fractions observed on day 5 to the heavier and more actively translated fractions on day 7 (Fig. 4). The amount of total AT1aR mRNA from 7-day cultures of VSMC that was extracted from fraction 2 was 2.7-fold greater than the amount using 5-day cultures [% of AT1aR mRNA in fraction 2 out of total AT1R mRNA recovered from the sucrose gradient: day 5, 20.9 ± 9.9 vs. day 7, 56.8 ± 5.6, n = 3, P < 0.001]. Furthermore, there was 60% less AT1aR mRNA found in fraction 3 in VSMC cultured for 7 days compared with cells cultured for 5 days [% of AT1aR mRNA in fraction 3 out of the total AT1R mRNA recovered from the sucrose gradient day 5, 60.9 ± 11.1 vs. day 7, 24.6 ± 3.1, n = 3, P < 0.05]. Real-time PCR also revealed that the AT1aR E1,2,3 splice variant was enriched in the denser polysome fractions on day 7 compared with day 5. There was a significantly greater proportion of E1,2,3 mRNA in the densest sucrose fraction (fraction 1) on day 7 compared with day 5 [% of E1,2,3 mRNA out of total AT1aR mRNA (E1,3 + E1,2,3) recovered from fraction 1: day 5, 7.16 ± 4.6 vs. day 7, 68.1 ± 1.0, n = 3, P < 0.01]. Consequently, the proportion of E1,3 mRNA in this fraction was reduced [% of E1,3 mRNA out of total AT1aR mRNA (E1,3 and E1,2,3) recovered from fraction 1: day 5, 92.8 ± 4.6 vs. day 7, 31.9 ± 1.0, n = 3, P < 0.01].
RNA EMSA of AT1R 5′LS and AT1R exon 2.
We have previously shown that alterations in the levels of cytosolic proteins, which bind to the 5′LS of the AT1aR are associated with inverse changes in AT1R protein expression (17, 18, 22). To determine whether the levels of RNA-protein complex (RPC) formation in the 5′LS are altered in proliferating VSMCs, we performed an RNA EMSA with radiolabeled 5′LS on cytosolic extracts prepared from VSMC cultured for 5 and 7 days (Fig. 5A). The degree of 5′LS RPC formation was reduced by 62% in VSMC cultured for 7 days compared with cells cultured for 5 days [5′LS RPC (AU): day 5, 0.62 ± 0.15 vs. day 7, 0.23 ± 0.03; n = 4, P < 0.05]. We observed a similar 49% decrease using radiolabeled exon 2 [Exon 2 RPC (AU): day 5, 35.0 ± 5.7 vs. day 7, 17.2 ± 3.6; n = 4, P < 0.05].
The present study demonstrates that the cell membrane density of AT1Rs increases in proliferating aortic VSMCs isolated from Fischer 344 rats. Furthermore, the data suggest that this increase in AT1R expression is in part due to more efficient translation of AT1aR mRNA. Additionally, we provide evidence suggesting that translation of the AT1aR during VSMC proliferation is regulated by proteins binding to exon 2 within the 5′LS of the AT1aR mRNA.
Scatchard analysis (Fig. 2A, inset) of radioligand saturation curves (Fig. 2A) indicates a single population of angiotensin receptors are expressed in membranes isolated from this rat aortic VSMC culture. Radioligand competition studies on VSMC membranes showed that the AT2R antagonist PD123319 was a poor competitor (Ki > 10 μM), whereas the AT1R antagonist SK1080 potently (Ki = 0.14 nM) and completely inhibited 125I-labeled [Sar1,Ile8]ANG II radioligand binding (Fig. 2B). Taken together, these data are consistent with previous reports showing that the AT1R subtype is the predominant subtype expressed in the vasculature (8). Previous studies have demonstrated that in VSMC cultures from Wistar-Kyoto (37), spontaneously hypertensive (37) and Sprague-Dawley (25) rats, binding of radiolabeled ANG II can be abolished by treatment with the AT1R specific nonpeptide antagonist losartan, whereas treatment with the AT2R-specific nonpeptide antagonist PD123177, has no effect on binding of radiolabeled ANG II to VSMC from Sprague-Dawley rats (25).
There are two known subtypes of the AT1R (AT1aR and AT1bR) present in both rats and mice. These receptors are 95% homologous at the amino acid level. Though only minor differences exist in their affinity for ANG II peptides, these receptors differ considerably in their tissue distribution. AT1aRs are widely distributed throughout the body, while AT1bRs are enriched in the pituitary and adrenal cortex (30, 31, 34). RNase protection assays failed to detect AT1bR, whereas only very low levels could be detected by real-time PCR in our VSMC cultures. These data indicate that the vast majority of AT1R mRNA expressed in these cells is of the AT1aR subtype. This is consistent with previous reports demonstrating that expression of AT1bR mRNA is either exceedingly low (4) or undetectable (27) in rat VSMC.
During proliferation of VSMC in culture, cell density increases from the initial plating at 1 × 104/cm2 to 20 × 104 on day 7 (Fig. 1A). Under these conditions, AT1R-specific binding steadily increases over the 7 days in culture (Fig. 1B, Fig. 2A). During VSMC proliferation, total cellular protein (Fig. 2C) and DNA content (Fig. 2D) did not significantly change, suggesting that the increase in AT1R binding is due to higher AT1R densities per cell rather than to increases in cell size or total cell protein. This conclusion is supported by Scatchard analysis of saturation binding curves on VSMC membranes, which revealed that AT1R densities increased by 30% between days 5 and 7 in culture (Fig. 2A).
Although we observed a marked increase in AT1R density, this increase was not associated with significant increases in AT1aR mRNA expression determined by RPA (Fig. 3) or by real-time PCR. We also did not observe any detectable de novo AT1R mRNA synthesis by nuclear run-on assay. Taken together, these data suggest that rates of AT1aR transcription are not increased during VSMC proliferation and therefore do not account for the increased receptor density. However, this study cannot rule out the possibility that the reason AT1aR mRNA levels did not detectably increase is because an increase in AT1aR transcription was offset by a decrease in AT1aR mRNA stability or vice versa. We hypothesized that posttranscriptional mechanisms contribute to the observed increases in AT1R density during VSMC proliferation since we could not detect increases in AT1aR mRNA levels under conditions in which AT1R densities were increased.
To determine whether the AT1R is translationally regulated in proliferating VSMC, we performed polysomal distribution analysis of AT1aR mRNA in VSMC cultured for 5 and 7 days. The basis of polysome distribution assays is that sucrose gradients separate polyribosomal fractions based on density. Because efficiently translated mRNA species have more ribosomes attached than less efficiently translated species, these more efficiently translated mRNAs are enriched in the denser fractions of the sucrose gradient (3, 29). We have previously used polysome distribution assays to demonstrate translational control of adrenal AT1Rs (38). VSMC AT1aR mRNA was predominantly found in sucrose fraction 3 on day 5. On day 7, the AT1aR mRNA was shifted to the heavier polysomal fraction 2 (Fig. 4). These data suggest that more ribosomes bind per AT1aR mRNA, and receptor translation is more efficient after 7 days in culture compared with 5 days. These findings suggest that the observed increase in AT1R density in proliferating VSMC is due, at least in part, to increased AT1R protein synthesis. However, the possibility that posttranslational mechanisms (e.g., attenuation of AT1R degradation or AT1R endocytosis) also contribute to the observed increase in AT1R densities cannot be excluded.
Two AT1aR splice variants are produced by alternative splicing: the first contains all three exons (E1,2,3), while the second contains only exons 1 and 3 (E1,3). The two-splice variants code for identical receptor proteins since exon 3 encompasses the entire open reading frame. However, we have shown in stably transfected CHO cells (13) and in transiently transfected rat aortic A10 cells (41) that exon 2 is inhibitory to translation resulting in reduced AT1R densities in cells expressing the E1,2,3 variant compared with cells expressing E1,3. Therefore, one mechanism by which AT1aR densities could increase is through an increased rate of excision of exon 2 during alternative splicing, despite a constant total AT1aR mRNA level. However, the present data do not suggest alternative splicing of the AT1aR is regulated in proliferating VSMC because the relative proportions of the two-splice variants were unchanged between days 5 and 7.
A second mechanism by which splice variants could play a role in modulating AT1R densities is by cellular regulation of the translational efficiency of the two-splice variants. For example, if conditions made translating the E1,2,3 variant more efficient then subsequent increases in AT1R densities could occur. Examination of the splice variant composition of the AT1aR mRNA in the sucrose gradient fractions suggests that the E1,2,3 transcript is more rapidly translated on day 7 than on day 5 since the proportion of E1,2,3 mRNA found in the heaviest polyribosomal fraction significantly increased on day 7 compared with day 5.
One possible mechanism by which the translational efficiency of AT1aR mRNAs could be controlled is through RNA binding proteins interfering with ribosomal scanning and translation initiation by binding to secondary structures in the 5′LS of the mRNA. We have previously shown that the ability of cytosolic proteins to bind the 5′LS of the AT1aR mRNA is regulated in a manner that inversely correlates with changes in AT1R densities under a variety of conditions, including dietary sodium manipulation (18), estrogen deficiency (17), and renal mass ablation (22).
To determine whether AT1aR 5′LS RNA binding proteins are regulated in proliferating VSMCs, we performed RNA EMSA on cytosolic extracts prepared from VSMCs cultured for 5 and 7 days using radiolabeled 5′LS and exon 2 RNA (Fig. 5A). RPC formation with the 5′LS was reduced by > 60% in cytosolic extracts prepared from day 7 VSMC compared with day 5. Furthermore, RPC formation was reduced by 50% in the day 5 cytosolic extracts compared with day 7 when using exon 2 RNA in the EMSA (Fig. 5B). These findings raise the possibility that a reduction in inhibitory RNA binding proteins interacting within exon 2 in the 5′LS may result in increased efficiency of AT1R translation in proliferating VSMC. To date, the identity of the protein components of this RPC and their mode of action remain unknown. One possible mechanism is suggested from studies on transferrin, in which binding of regulatory proteins to a hairpin loop in the 5′LS of the mRNA inhibits translation initiation and ribosome scanning by steric hindrance (15). Potentially, similar mechanisms involving RNA binding proteins interfering with ribosomal scanning could also be involved in translational regulation of the AT1aR. These data also raise the possibility that the reason the E1,2,3 transcript is enriched in the heavier polysomal fractions on day 7 is because of the reduction in the amount of RPC formation to exon 2 compared with day 5.
It is interesting to note that the alteration in translational efficiency of AT1aR mRNA between days 5 and 7 coincides with the cells reaching confluence. This observation raises the possibility that the increase in AT1R binding might be related to the proliferative state of the cells and that translational regulation of AT1R expression is regulated by changes in rates of cell proliferation. Increased proliferation of VSMCs is associated with events such as intimal thickening which contribute to the formation of atherosclerotic lesions (2, 12, 16, 19, 23). Increased expression of various components of the RAS, including AT1Rs, has been associated with a number of disease states, including atherosclerosis, various forms of hypertension, and cardiovascular and progressive renal disease (5, 32). The present data suggest that AT1R expression increases as VSMC proliferate in culture by a mechanism that involves translational regulation of the receptor by proteins that bind to exon 2 within the 5′LS. Therefore, translational regulation of the AT1aR during VSMC proliferation may play a role in processes mediating vascular disease. Future studies directed toward understanding the role 5′LS RNA-binding proteins in translational regulation of AT1Rs may lead to the development of novel therapeutics for disease states in which dysregulation of AT1R expression occurs.
This research was supported by National Institutes of Health Grants HL-7502 and AG-19291 to K. Sandberg and an American Heart Association Jocelyn Beard Moran Memorial Fellowship grant to A. Hassan.
Present address for S. Lee: Dept. of Biotechnology and Informatics, Sangmyung University, San 98–20, Anseo-Dong, Cheonan, Republic of Korea, 330–7204 (e-mail: firstname.lastname@example.org.)
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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