Published online before print November 14, 2005, doi: 10.1161/CIRCULATIONAHA.105.541714
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(Circulation. 2005;112:3328-3336.)
© 2005 American Heart Association, Inc.
Vascular Medicine |
Angiotensin Type 2 Receptor Is Expressed in Murine Atherosclerotic Lesions and Modulates Lesion Evolution
From the Cardiovascular Division (V.L.S., M.A.L.-I., V.J.D., R.E.P.) and Donald W. Reynolds Cardiovascular Clinical Research Center (G.K.S., P.L.), Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, Mass. Dr Sales is now at the Department of Cardiac Surgery, Children’s Hospital Boston, Harvard Medical School, Boston, Mass; Dr Dzau is now at the Office of the Chancellor, Duke University Medical Center, Durham, NC.
Correspondence to Virna L. Sales, MD, Department of Cardiac Surgery, Children’s Hospital Boston, 300 Longwood Ave, Bader 279, Boston, MA 02115. E-mail virna.sales{at}childrens.harvard.edu
Received February 8, 2005; revision received August 1, 2005; accepted September 12, 2005.
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Abstract |
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Background— In the vasculature, the angiotensin type 2 (AT2) receptor (AT2R) exerts antiproliferative, antifibrotic, and proapoptotic effects. Normal adult animals have low AT2R expression; however, vascular injury and exposure to proinflammatory cytokines augment AT2R levels. We hypothesized that AT2R expression increases during initiation and progression of atherosclerosis.
Methods and Results— Atherosclerotic lesions of apolipoprotein (Apo) E–/– mice contained AT2Rs, measured by real-time polymerase chain reaction and confirmed by immunohistochemistry. To test the consequences of this expression, male ApoE–/–, angiotensin II type 2 receptor-deficient (Agtr2–), and ApoE–/–, wild-type (Agtr2+) mice consumed a high-cholesterol diet from 4 weeks of age. Ten weeks later, overall area and cellular composition of aortic arch lesions did not differ significantly among genotypes. After 16 weeks, ApoE–/–/Agtr2+, but not ApoE–/–/Agtr2– mice had dramatic decreases in percent positive area of macrophages, smooth muscles, lipids, and collagen. Diminished bromodeoxyuridine incorporation and increased TUNEL staining accompanied these decreases.
Conclusions— Thus, loss of AT2R during the evolution of atherosclerotic lesions augmented the extent of cellularity of atherosclerotic lesions, establishing AT2R as a modulator of atherogenesis.
Key Words: angiotensin • atherosclerosis • inflammation • receptors • renin
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Introduction |
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Angiotensin II (Ang II), the main effector of the renin-angiotensin system, regulates blood pressure by controlling vasoconstriction and maintaining water and salt balance. Ang II also affects diverse cellular systems, including growth and proliferation. Ang II binds to types 1 (AT1) and 2 (AT2) angiotensin receptors, members of the superfamily of heptahelical G protein-coupled receptors. Despite abundant physiological and biochemical characterization of AT1 receptor (AT1R), little is known about the AT2R. Fetal tissues, including the vasculature, express AT2R abundantly and widely, but normal adult tissues have only low levels of AT2R.1,2 We and others have shown augmented expression of AT2R after vascular injury.3–10
Clinical Perspective p 3336
In vivo and in vitro studies demonstrate the growth-promoting effects of AT1R, but evidence suggests that AT2R inhibits cellular growth. For example, gene transfer studies show that AT2R expression in cultured adult smooth muscle cells (SMCs) antagonizes the growth-promoting effects of AT1 receptor and proliferation induced by growth factors such as platelet-derived growth factor.9 Gene transfer studies in vivo demonstrate that AT2R expression attenuates neointimal accumulation in injured carotid arteries.9 Vascular injury studies with mice harboring disruptions of AT2R show exaggerated neointimal development compared with wild-type mice.3–10
Pharmacological studies established that AT2R expression in the fetal vasculature mediates a developmentally regulated decrease in vascular proliferation in late gestation.5,11 Thus, in vivo and in vitro studies support antiproliferative actions of AT2R. We and others have shown AT2R-mediated apoptosis in numerous cells types, including SMCs, in cell culture and in vivo.6,12,13 The reexpression of AT2R in injured vasculature may have important consequences.
These findings suggest the importance of factors that regulate AT2R expression, especially under pathological conditions. Interferon-
, through the transcription factor IRF-1, augments AT2R expression in cultured SMCs,3,14–16 an example of increased AT2R regulation as a result of inflammatory cytokine signaling.
Induction of AT2R expression by cytokines suggested the involvement of the receptor in other instances of vascular inflammation, eg, development of atherosclerotic lesions. Given the antiproliferative actions and role in vascular remodeling of AT2R, we hypothesized that AT2R expression within atherosclerotic lesions influences lesion development. Here, we examine the functional consequences of AT2R in atherosclerotic lesions in apolipoprotein (Apo) E–/– mice.
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Mouse Breeding
Animal care and experimental procedures were approved by the Animal Care Committees of Harvard Medical School.
Mice homozygous for disruption of AT2R- (sixth-generation backcross to FVB/N background, had 98% FVB/N and 2% 129/SV background)5,10 and ApoE-deficient mice (10th-generation C57BL/6; Jackson Laboratory, Bar Harbor, Me) were used. Male ApoE–/–/Agtr2+ mice were crossed with female ApoE+/+/Agtr2– mice to produce ApoE+/–/Agtr2+/– female and ApoE+/–/Agtr2– male mice. Further brother-sister mating produced an F4 generation containing ApoE–/–/Agtr2– or ApoE–/–/Agtr2+ animals used in our studies. We used only littermates in all our experiments to avoid confounding genetic strain differences. Thus, the generated litters of ApoE–/–/Agtr2– or ApoE–/–/Agtr2+ were neither pure FVB/N nor C57BL/6 strains. Double-knockout (ApoE–/–/Agtr2–) mice were born at expected Mendelian ratios and developed normally. Male ApoE–/–/Agtr2– (n=30) or ApoE–/–/Agtr2+ (n=30) littermates were used at 4 weeks of age; their weight was 15 to 20 g before diet and 30 to 35 g after 10 or 16 weeks of standard or high-cholesterol diet.
Polymerase chain reaction (PCR) confirmed double-knockout (ApoE–/–/Agtr2–) and ApoE-deficient (ApoE–/–/Agtr2+) mice. The primer sequences used were as follows: primer 1, 5'-GCCTAGCCG-AGGGAGAGCCG-3'; primer 2, 5'-TGTGACTTGGGAGCTCTG-CAGC-3'; primer 3, 5'-GCCGCCCCGACTGCATCT-3'; primer 4, 5'-CTCACTGTTTTGTTGTC-3'; primer 5, 5'-CAATGGTTCTGA-CATCC-3'; and primer 6, 5'-CTTCCATTGCTCAGCGGT-3'. Primers 1 and 2 produced a 155-bp product from the wild-type ApoE allele; primers 1 and 3, a 245-bp product from the disrupted ApoE allele; primers 4 and 5, a 365-bp product from the wild-type AT2 allele; and primers 4 and 6, a 350-bp product from the disrupted AT2 allele.
Experimental Design
ApoE–/–/Agtr2– or ApoE–/–/Agtr2+ littermates were weaned at 4 weeks and fed a high-cholesterol diet (1.25% cholesterol, no cholate, No. D12108 Research Diet) or a standard chow diet.17 For tissue harvest, mice were anesthetized with xylazine (10 mg/kg IP) and ketamine (80 mg/kg IP) after 24-hour fasting. Blood (0.5 to 1 mL) was collected from the inferior vena cava for lipid profiling. Plasma total cholesterol and triglyceride were measured in duplicate with standard colorimetric assays (Sigma Chemical). The arterial tree was pressure-perfused for 5 minutes at 100 mm Hg with 10 mL 1x PBS through a catheter (Butterfly 21-inch x 0.75-inch with 12-inch tubing infusion set, Abbot) inserted into the left ventricular apex with fluid drained from a severed right atrium. Aortic arches were dissected and snap-frozen in OCT (OCT compound, Tissue-Tek).
Immunohistochemistry
Serial cryostat sections (6 µm) of aortic arch were cut, fixed in acetone (–20°C, 5 minutes) or 4% paraformaldehyde, air-dried, and stained with antibodies specific for murine smooth muscle 
-actin (FITC conjugate, Sigma) and macrophage marker, Mac-3, 1:1000 (PharMingen) as described.18–21 Immunostaining was performed by use of the ABC method with biotin-labeled secondary antibodies (Vector Laboratories) and using 3-amino-9-ethyl carbazole (AEC, Dako) as substrate. Sections were counterstained with Gill hematoxylin solution (Sigma). Specificity controls used an appropriate nonimmune IgG instead of antibody. Immunohistochemical detection of murine AT2R in lesions used goat anti-mouse AT2 polyclonal antibody (Santa Cruz, diluted 1:250 in 5% normal serum in PBS) and biotin-labeled donkey anti-goat (Santa Cruz, diluted 1:200 in 5% normal serum in PBS) as secondary antibody.
For colocalization of 5'-bromo-2'-deoxyuridine (BrdU) and AT2 with the respective cell types, double immunostaining was performed. Frozen sections were stained for AT2R as described above and for BrdU according to the manufacturer’s protocol (BD Biosciences). After the first staining, sections were treated with an avidin/biotin blocking kit (Vector Laboratories), washed, blocked again with 5% appropriate normal serum for 20 minutes, and stained with monoclonal cell-type-specific antibodies (overnight, 4°C), biotinylated secondary antibody, and finally streptavidin coupled to alkaline phosphatase (30 minutes), and visualized with Fast blue (AT2) and Fast red (BrdU) substrates that produced blue and red reaction products, respectively.20–22 Double-stained cells showed purple staining as a mixture of red (SMCs or macrophages) and blue (AT2R) colors. Double-stained cells showed mixtures of brown nuclei (BrdU) and red cytoplasm (SMC or macrophages). Picrosirius red staining evaluated by polarization microscopy visualized interstitial collagen content, and deposition of lipids was determined by oil red O staining as described.18–21
Quantification and Evaluation of Atherosclerotic Lesions
Evaluation of atherosclerotic lesions in experimental mice was performed by use of longitudinal sections of aortic arches. A 3-mm segment of the aortic arch lesser curvature was defined by a perpendicular line dropped from the right side of the innominate artery, as described previously,18,21 to measure lesion size (intima and media) and percentage of intimal lesion area stained for SMCs, macrophages, interstitial collagen, or lipid. Images captured by a digital imaging system were analyzed with Image-Pro Plus software (Media Cybernetics) by 2 blinded observers independently. A manual threshold process was used to determine the area of positive immunoreactivity. The percentage of total area with positive color was recorded for each mouse.
Measurement of Cellular Proliferation and Apoptosis
Mice received BrdU (Sigma; infusion rate, 13 mg · kg–1 · d–1) by subcutaneous osmotic minipump (Alzet 2001, Alza Corp) for 7 days before euthanasia.23 BrdU and terminal deoxynucleotidyl transferase (TdT)–mediated dUTP nick end-labeling (TUNEL) followed the manufacturer’s protocol (BrdU staining Kit, BD Biosciences; ApopTag Peroxidase In Situ Apoptosis Detection Kit, Intergen) with DAB as a chromogen. Sections were counterstained with methyl green solution (Dako). Ileum and thymus were positive controls for BrdU and TUNEL staining, respectively. For negative controls, BrdU antibody and TdT were omitted from the labeling mixture. Cells were evaluated microscopically at x400. Cells containing dark nuclear BrdU staining were considered to be BrdU+ and to have undergone DNA synthesis during labeling. Apoptotic nuclei were defined as TUNEL-positive nuclei in cells with morphological features of apoptotic cell death (cell shrinkage, aggregation of chromatin into dense masses, and cell fragmentation). Overall proliferation and apoptosis data for intimal lesions were obtained by counting all intimal cells and all intimal BrdU- and TUNEL-positive cells in sections.
RNA Isolation and Real-Time PCR Analysis
Aortic arch tissues were washed briefly with 1x PBS before RNA extraction. Total RNA was homogenized and extracted with TRIZOL reagent (Gibco-BRL Life Technologies). Quantity and quality of each sample were evaluated spectrophotometrically (Beckman Coulter, DU640B) at 260 and 280 nm. RNA quality was examined on a 1% formaldehyde agarose RNA gel. PCR primers and fluorogenic probes for AT2R gene were designed with Primer Express software (Applied Biosystems). TaqMan 18S ribosomal RNA control reagents (Applied Biosystems) served as references to normalize input amounts of RNA for all samples. Probes were selected for a melting temperature (Tm) of 7°C to 10°C higher than the matching primer pair. Fluorogenic probes contained an FAM report dye coupled at the 5' end and a BHQ1 quencher dye coupled at the 3' end and were high-performance liquid chromatography-purified. The forward primer for AT2R mRNA corresponds to residues 30 to 49 of full-length mRNA with a Tm of 59°C (5'-GATGGAGGGAGCTCGGAACT-3'); the reverse primer corresponds to residues 172 to 150 with a Tm of 60°C (5'-TTGAACTGCAGCAACTCCAAATT-3'). TaqMan probe corresponds to residues 51 to 83 with a Tm of 69°C (5'-AAAGCTTACTTCAGCCTGCATTTTAAGGAGTGC-3'). Real-time RT-PCR used iCycler IQ Real-Time Detection Systems (Bio-Rad). SuperScript One-step RT-PCR with Platinum Taq kit (Invitrogen) was used for all RT-PCR amplification in a total volume of 50 µL, containing 200 ng total RNA, 5 mmol/L MgSO4, 500 nmol/L forward and reverse primers, and 200 nmol/L fluorogenic probe. RT-PCR amplification for each RNA sample was performed in triplicate wells. Controls omitted reverse transcriptase for each RNA sample and mRNA for each primer and probe set. The 1-step RT-PCR condition was as follows: 15 minutes at 50°C and 5 minutes at 95°C, followed by 45 two-temperature cycles (15 seconds at 95°C and 1 minute at 60°C). Relative gene expression data analysis was performed by use of a standard curve method.24,25
Statistical Analysis
All results are expressed as mean±SEM. Data were compared for feeding duration and genotype, and differences were determined with Student t test or nonparametric Mann-Whitney test followed by appropriate post hoc test (SigmaStat, Jandel Scientific). Two investigators blinded to feeding duration and genotype made all measurements. A value of P<0.05 was considered significant.
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AT2 Receptor Expression and Colocalization in Atherosclerotic Plaque
On the basis of observations that inflammatory cytokines and vascular injury increased AT2R expression, we hypothesized that atherosclerotic lesions overexpress AT2R. To test this hypothesis, we designed an RT-PCR assay with primers in exons 1 and 3 designed to amplify the mouse AT2R sequence. We also designed a Taqman probe. Amplification of RNA from neonatal mice yielded a PCR product of expected size and sequence (data not shown).
Consumption of the Atherogenic Diet Induced Arterial AT2R mRNA
One group each of 4-week-old of ApoE+/+/Agtr2+ and ApoE–/–/Agtr2+ mice consumed an atherogenic diet (40% fat, 1.25% cholesterol) for 2 months; another set of groups ate normal chow. Two months later, the mice were killed, aortic arches were isolated, and RNA was extracted and analyzed by RT-PCR. In C57Bl/6J mice on normal diets, AT2R mRNA expression in the aortic arch was very low (Figure 1A). ApoE–/–/Agtr2+ mice on normal chow had much greater levels of aortic arch AT2R mRNA than wild-type mice on a comparable diet. ApoE–/–/Agtr2+ mice on an atherogenic diet had 3-fold higher AT2R mRNA levels than genetically identical animals on normal chow.
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We next examined localization of AT2R using double immunostaining with cell type-specific markers. As above, ApoE–/–/Agtr2+ mice consumed an atherogenic diet starting at 4 weeks of age, and at 2 and 4 months the aortic arches were removed and embedded in OCT. AT2R immunoreactivity in vessel walls localized predominantly in intimal lesions, with little or no immunoreactivity in the tunica media. Double immunohistochemistry colocalized AT2R with both macrophages and SMCs within intimal lesions of ApoE–/– mice (Figure 1B). Most of the Mac-3–positive macrophages (left bottom, red) and -actin-positive SMC (right bottom, red) also displayed AT2R staining (purple staining, left and right, top).
AT2 Receptor Expression and Evolution of Atherosclerotic Lesions
AT2R Deficiency Does Not Alter Lesion Size
ApoE–/–/Agtr2– double-knockout mice and ApoE–/–/Agtr2+ littermates consumed an atherogenic diet starting at 4 weeks of age. After 10 weeks of atherogenic diet, the lesion area in ApoE–/–/Agtr2– mice was slightly but not significantly greater than that in ApoE–/–/Agtr2+ mice. After 16 weeks of atherogenic diet, lesion areas in ApoE–/–/Agtr2+ and ApoE–/–/ Agtr2– mice exceeded those at 10 weeks and both had increased similarly (1.5 versus 1.6). Lesions in aortae of ApoE–/–/Agtr2– were slightly but not significantly greater than those of ApoE–/–/Agtr2+ mice. Medial area did not differ among the 4 groups (data not shown).
AT2R Deficiency Increases SMC and Collagen Content of Mature Atheromata
Sections were stained for smooth muscle, macrophages, interstitial collagen, and lipid. After 10 weeks of atherogenic diet, ApoE–/–/Agtr2+ mice developed large atherosclerotic lesions on the aortic arch inner curvature (Figures 2 and 3
). Staining with smooth muscle- or macrophage-specific antibodies revealed substantial accumulation of both cell types (25% of area SMCs and 34% of area macrophages).
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Staining with oil red O revealed considerable lipid accumulation within lesions. Collagen content was also high (31%). Between 10 and 16 weeks, the composition of lesions in ApoE–/–/Agtr2+ mice changed substantially. The area occupied by macrophages and lipid each fell by some 65% to 70% in the ApoE–/–/Agtr2+ mice (Figure 2). Smooth muscle and collagen decreased even more markedly over this time, by 65% to 85% (Figure 3).
Effects of AT2R Deficiency on Atherosclerotic Plaques
After 10 weeks on the diet, macrophage, lipid, smooth muscle, and collagen contents in lesions of ApoE–/–/Agtr2– mice did not differ significantly from those in ApoE–/–/Agtr2+ mice (Figures 2 and 3
). Thus, in these early lesions, loss of AT2R had little effect on plaque composition but did profoundly affect plaque evolution. The marked decrease in percent positive area in aortic lesions of ApoE–/–/Agtr2+ mice between 10 and 16 weeks did not occur in ApoE–/–/Agtr2– mice. However, the percent positive area for collagen in ApoE–/–/Agtr2– mice increased by 55% from 10 to 16 weeks.
These altered lesion compositions did not associate with differences in plasma cholesterol or triglyceride levels. After 10 weeks of the high-cholesterol diet, total serum cholesterol for ApoE–/–/Agtr2+ (n=9) and ApoE–/–/Agtr2– (n=9) mice was 1021±102 and 991±144 mg/dL, respectively. Similarly, total serum triglyceride was 170±18 and 207±39 mg/dL, respectively. After 16 weeks of the high-cholesterol diet, total serum cholesterol for ApoE–/–/Agtr2+ (n=9) and ApoE–/–/Agtr2– (n=11) mice was 986±144 and 1017±66 mg/dL, respectively. Similarly, total serum triglyceride was 242±34 and 233±22 mg/dL, respectively. No statistical difference was observed between ApoE–/–/Agtr2+ and ApoE–/–/Agtr2– at either 10 or 16 weeks.
Do Differences in Cellular Proliferation and/or Cellular Apoptosis Explain Differences in Cellular Composition?
We assessed the BrdU and TUNEL labeling index in these atherosclerotic lesions. Four-week-old animals were fed an atherogenic diet. At 9 or 15 weeks, pumps administering BrdU were implanted intraperitoneally; 1 week later, the aortic arch was dissected and embedded in OCT. Immunohistochemical analysis of lesions revealed that BrdU incorporation during week 10 did not differ significantly between the 2 strains (ApoE–/–/Agtr2+ versus ApoE–/–/Agtr2–). By week 16, BrdU incorporation in lesions of ApoE–/–/Agtr2+ mice fell significantly from that at 10 weeks, whereas rates of incorporation into lesions of ApoE–/–/Agtr2– was unchanged (Figure 4A). Thus, differences in cellular proliferation may contribute to altered cellular composition at 10 and 16 weeks. Double immunostaining suggested that both macrophages and SMCs in intimal lesions proliferated during this period (Figure 4B).
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Does Loss of AT2R Influence Apoptosis?
At 10 weeks, rates of apoptosis in lesions assessed by TUNEL staining were virtually identical between strains (ApoE–/–/Agtr2+ versus ApoE–/–/Agtr2–). At 16 weeks, the frequency of apoptotic cells in lesions of ApoE–/–/Agtr2+ mice was more than twice that in ApoE–/–/Agtr2– mice. Thus, loss of AT2R results in decrease in apoptotic cells (Figure 5).
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These data indicate that differences in cellular composition during evolution of plaque depend on both changes in rates of proliferation and apoptosis in macrophages and SMCs within lesions.
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The renin-angiotensin system crucially regulates the structure of nonatherosclerotic arteries by balancing growth-promoting, antiapoptotic, and profibrotic effects of AT1R and antigrowth, proapoptotic, and antifibrotic effects of AT2R.3–13 The present study demonstrates that during atherogenesis in ApoE–/– mice, the loss of AT2R elevated macrophage, smooth muscle, and collagen content of atherosclerotic lesions, probably because of enhanced proliferation and decreased apoptosis of lesional cells.
High AT2R expression in many fetal and early neonatal tissues declines precipitously in the first postnatal days. A few adult tissues have residual AT2R expression: adrenal gland, brain, myometrium, and atretic ovarian follicles. We and others have shown that injury elicits reexpression of AT2Rs, perhaps in part through an inflammatory pathway, because interferon- induces AT2R expression via IRF-1 transactivation in cultured SMCs.16
An in vivo model of vascular inflammation (placing a polyethylene cuff around the femoral artery) shows expression of AT2R coinciding with expression of interferon-, interleukin (IL)-1ß, IL-6, tumor necrosis factor-, and IRF-1.3 Causal relationships among expression of cytokines, IRF-1, and AT2R were provided by cuff injury studies using recombinant activating gene-1– and IRF-1–knockout animals: Cuff injury failed to induce AT2R expression (V.S., V.D., and R.P., unpublished data, 2002). Thus, expression of AT2R in our report agrees with known patterns of receptor regulation.
We and others3,8,16 have found a marked decrease in SMC and collagen content during atherosclerotic plaque evolution between 10 and 16 weeks of consumption of an atherogenic diet. This change in cellular composition most likely results in part from inflammatory signaling. For example, when ApoE–/– mice are exposed to high levels of HDL, macrophage and T-cell composition within plaque decreases, whereas smooth muscle and collagen content increases.26 Moreover, interruption of CD40 L signaling produces similar changes in composition.19 In ApoE–/– mice treated with IL-10 or transforming growth factor, macrophages decrease and collagen increases.27,28 Human plaques, especially unstable plaques, show a strong inverse correlation between inflammation and smooth muscle content.28–32
We found the presence of AT2R to be strikingly reduced the evolution of atheroma, marked by a decrease in smooth muscle and collagen, implicating AT2R in the regulation of plaque cellularity and composition. In other models of vascular lesion formation, intimal SMCs that populate lesions express AT2R.3,4,7–10 Consistent with known actions of AT2R, ApoE–/–/Agtr2– mouse aorta incorporated more BrdU and had less TUNEL staining, indicating an increase in proliferation and a decrease in apoptosis, respectively, and providing a likely mechanism for changes in cellular composition. We and others showed that AT2R inhibits collagen accumulation,7,10 thus permitting speculation that AT2R is reexpressed during plaque evolution, consistent with the in vivo role demonstrated here for AT2R in limiting smooth muscle and collagen accumulation during the progression of atheromata.
The effect of AT2R inactivation on accumulation of macrophages was unanticipated. Although smooth muscle, endothelial cells, and fibroblasts express AT2R, only 1 in vitro study reports its presence in monocytes or macrophages.33 Double immunostaining with cell type-specific antibodies showed AT2R localized in macrophages and SMCs within atherosclerotic lesions, but whether the decrease in macrophage content in evolving plaque of ApoE–/– mice results directly from engagement of the receptor on the macrophage or indirectly because of a paracrine effect of AT2R expressed on smooth muscle remains unclear. Although knowledge of the AT2R signaling pathway is incomplete, stimulation of the receptor produces a kinin-dependent stimulation of NO production. Actions of locally produced NO may include a decrease in oxidative stress and subsequent decrease in attraction of monocytes, decreased proliferation, and/or increased macrophage apoptosis.
Cell death in atheromata may have complex effects on lesion evolution and complication. Apoptosis of SMCs may limit lesion bulk and reduce fibrosis but, by impairing the collagen synthesis needed to maintain the fibrous cap, may render some lesions more susceptible to rupture and hence thrombosis.34,35 However, apoptosis of macrophages should combat inflammation, reduce local cytokine and protease release, and thus overall stabilize plaques, albeit contributing to formation of the lipid core of the lesion.
Most known proatherogenic effects of Ang II are generally accepted as being mediated by AT1R.36–42 The lesional composition in AT2-disrupted mice could be caused by a compensatory increase in AT1R. AT1R levels rise in hyperlipidemia,37–39 as demonstrated by markedly high serum cholesterol levels in ApoE–/– mice after an atherogenic diet.38,43 Cuff-induced vascular injury and pressure-overload cardiac hypertrophy demonstrated the effects of AT1R expression in AT2R-disrupted mice.3,6,8,10 The effect of AT2R inactivation does not require pronounced hyperlipidemia. We found similar effects of enhanced lesion formation in aortae of ApoE–/–/Agtr2– compared with ApoE–/–/Agtr2+ mice after 16 weeks on a chow diet (data not shown).
There are several limitations within the present study. First, we were unable to obtain plasma Ang II, renin, and circulating angiotensin peptide measurements in our mouse model of atherosclerosis. However, we believe that our results on the enhanced atherosclerotic lesion formation in ApoE–/–/Agtr2– are likely to be reflected in the modulatory role of AT2R. Recently, Daugherty and colleagues44 reported that absence of AT2R in LDL receptor-knockout mice had no effect on plasma aldosterone. Indeed, previous studies by our group and others demonstrate that levels of plasma renin, ACE, Ang II, and circulating angiotensin peptides are comparable in both Agtr2+ and Agtr2– mouse strains.4,45 Wu and colleagues7 also observed that the inhibitory effects of a nonselective Ang II receptor antagonist, such as Sar1,Ile8–Ang II, on cardiac hypertrophy, coronary thickness, and perivascular fibrosis were not different between Agtr2+ and Agtr2– mice. However, Levy and colleagues46 reported that chronic blockade of AT2 receptor by PD 123319 had no effect on plasma Ang II levels. On the basis of these data, we can predict that in our mouse model of atherosclerosis, there will be no elevations in systemic Ang II, renin, and angiotensin peptide concentrations that might contribute to the atherogenic process.
The delay in the effects of increased AT2R expression was a result of a number of factors. First, the difference in the genetic background of the mice needed to be taken into account. Because the genetic background of our ApoE–/–/Agtr2– or ApoE–/–/Agtr2+ littermates used in Figures 2 through 5 is a mixed background of FVB/N and C57 BL/6 mice, we used only littermates in the experiments described in Figures 2 through 5 to avoid confounding genetic strain differences. The mice used in Figure 1 are C57 BL/6 (ApoE–/– mice). The conflicting results may suggest that the heterogeneity effects of AT1 and/or AT2 stimulation may be a result of different tissues, cells, experimental conditions, species, and/or genetic background.8 These issues will be addressed in future experiments. In the present context, our results provide the role of AT2R in modulating atherosclerotic lesion formation. Second, the detailed temporal pattern of AT2R expression during the initiation and progression of atheroma is undefined. After the initiation of atherosclerotic lesion development, SMCs migrate from the medial layer into the intima, dedifferentiate, and proliferate. Monocytes then enter the intimal lesions, become activated, and differentiate into macrophages. During this process, the AT2R is upregulated as a result of cytokine expression. During plaque evolution, AT2R expression on intimal smooth muscle results in a decrease in smooth muscle in intima. This decrease is because of a decrease in proliferation and increase in apoptosis. Our observations suggest a temporal association of AT2R expression with the reduction in SMC content in the lesion evolution. These results suggest strongly that AT2R modulates atherosclerotic lesion formation, perhaps by controlling the stimulatory effects of developmentally regulated growth factors or through another mechanism, such as apoptosis.
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We know of no publications demonstrating that AT2R modulates atherosclerotic lesion formation. The demonstration of the antiinflammatory and antiproliferative actions of AT2R has important implications in the pathogenesis of human atherosclerosis. Long-term consequences of increased AT2R signaling in the atheroma are still unknown, leading to its important pathophysiological relevance. If stimulation of AT2R decreases inflammation and proliferation in atherosclerotic lesion evolution, therapies that favor receptor stimulation, eg, blockade of AT1R that leaves Ang II available to stimulate AT2R, may prove protective in humans.47 Our findings in this study may be of clinical significance, because the regression of atherosclerotic lesions by ARBs may be partly a result of an unopposed antigrowth and antiinflammatory effect of Ang II mediated via AT2. The combination of ARB with ACE inhibitor represents an appealing therapeutic strategy, because of the possibility of its producing more complete blockage of the renin-angiotensin system while preserving the beneficial effects mediated by AT2 receptor stimulation and increased bradykinin levels.48 An AT1R antagonist plus an ACE inhibitor may stabilize plaques, a possible clinical extension of the role for AT2R in the regulation of the composition of experimental atheroma, as demonstrated here.
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Acknowledgments |
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This work was supported by National Institutes of Health (NIH) grant RO1-HL-61661 (to Dr Pratt); by the Leducq Foundation (to Drs Libby and Sukhova); and by NIH/National Heart, Lung, and Blood Institute grants RO1-HL-34636 to Dr Libby and RO1-HL-67249 to Dr Sukhova. Dr Dzau is a recipient of an NIH Merit award. Dr Lopez-Ilasaca is the recipient of an American Heart Association National Scientist Development Grant (0435427T). The authors acknowledge the excellent technical assistance of Eugenia Shvartz, Weining Lu, and Margaret Lei and the editorial expertise of Jaylyn Olivo (Brigham and Women’s Hospital).
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References |
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Guest Editor for this article was John F. Keaney, Jr, MD.
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