Research ArticleCardiovascular Biology

Obligatory role for GPER in cardiovascular aging and disease

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Science Signaling  01 Nov 2016:
Vol. 9, Issue 452, pp. ra105
DOI: 10.1126/scisignal.aag0240

Preventing the ravages of ROS

Ligand-dependent activation of the G protein–coupled estrogen receptor (GPER) has been reported to confer cardiovascular benefits. However, Meyer et al. found that genetic absence of Gper conferred protection from cardiovascular pathologies associated with aging and hypertension. GPER activity was required to increase the abundance of the enzyme Nox1 in vascular smooth muscle cells, blood vessels, and myocardium, and was associated with enhanced production of tissue-damaging superoxide. Aged mice that were deficient in Gper developed much less cardiac fibrosis and hypertrophy and also retained greater cardiovascular function. In addition, a pharmacological inhibitor of GPER reduced blood pressure, superoxide production, and Nox1 abundance in hypertensive mice. Thus, inhibitors of GPER are potential therapies for cardiovascular diseases and conditions characterized by excessive superoxide generation.


Pharmacological activation of the heptahelical G protein–coupled estrogen receptor (GPER) by selective ligands counteracts multiple aspects of cardiovascular disease. We thus expected that genetic deletion or pharmacological inhibition of GPER would further aggravate such disease states, particularly with age. To the contrary, we found that genetic ablation of Gper in mice prevented cardiovascular pathologies associated with aging by reducing superoxide (⋅O2) formation by NADPH oxidase (Nox) specifically through reducing the expression of the Nox isoform Nox1. Blocking GPER activity pharmacologically with G36, a synthetic, small-molecule, GPER-selective blocker (GRB), decreased Nox1 abundance and ⋅O2 production to basal amounts in cells exposed to angiotensin II and in mice chronically infused with angiotensin II, reducing arterial hypertension. Thus, this study revealed a role for GPER activity in regulating Nox1 abundance and associated ⋅O2-mediated structural and functional damage that contributes to disease pathology. Our results indicated that GRBs represent a new class of drugs that can reduce Nox abundance and activity and could be used for the treatment of chronic disease processes involving excessive ⋅O2 formation, including arterial hypertension and heart failure.


G protein (heterotrimeric GTP-binding protein)–coupled receptors (GPCRs) exert both rapid and chronic effects (1). The G protein–coupled estrogen receptor (GPER) is a heptahelical receptor [originally designated GPR30, with amino acid homology to GPCRs for angiotensin II (Ang II) and chemokines] that is found in multiple cell types including vascular cells (24). GPER is localized predominantly to the endoplasmic reticulum and Golgi apparatus (5) and mediates cellular responses to estrogens, selective estrogen receptor modulators, and xenoestrogens (6) through nongenomic and genomic mechanisms (5, 79). GPER activation results in the rapid mobilization of intracellular calcium (5, 10, 11) and activation of nitric oxide (NO) synthase (12), Akt (11, 1316) and extracellular signal–regulated kinase (ERK) (7, 11, 16), among other pathways (6). In addition, GPER, like many other GPCRs (1, 17), may also exhibit basal or intrinsic activities that contribute to the chronic regulation of genomic pathways. Such genomic effects have been suggested to be responsible for the increased vasoconstrictor tone observed in Gper-deficient mice (18). The beneficial cardiovascular effects of GPER-selective synthetic ligands in multiple disease models (4, 12, 19) have led to the currently prevailing concept that activation of GPER conveys organ protection (6, 11, 2023).

Reactive oxygen species (ROS) are short-lived intermediates of oxidative metabolism that are essential for cardiovascular homeostasis (24). Excessive ROS production, however, occurs in many chronic disease processes and aggravates vasoconstriction and cell growth, thereby promoting increased vascular tone, myocardial hypertrophy, fibrosis, heart failure, and aging (25, 26). The NADPH oxidase (Nox) family represents the principal physiological source of ROS in the cardiovascular system and is composed of seven catalytic subunits termed Nox1 to Nox5 and Duox1 and Duox2 (2729). Of these, Nox1, Nox2, and Nox4 have been implicated in both experimental and human hypertension and heart failure, yet their roles in cardiovascular aging are less understood (27, 28).

Given that studies using the GPER-selective agonist (GRA) G-1 have demonstrated that GPER activation conveys partial protection from vascular and myocardial disease (4, 13, 30) and given the central role of ROS in these chronic disease processes (24, 25, 27), we expected that pathologies characterized by increased bioactivity of ROS, particularly those associated with aging, would be exacerbated in the absence of functional GPER, resulting in even greater ROS production. We therefore set out to determine the functional and structural effects of genetic deletion (20) as well as effects of pharmacological inhibition (31, 32) of GPER on ROS-dependent pathologies affecting the cardiovascular system.


GPER increases Nox-dependent vascular ⋅O2 formation and vascular tone in aged arteries

We first examined the effects of Gper deletion on vascular oxidative stress by measuring the production of the unstable free radical superoxide (⋅O2) in the aorta of aged mice. To determine whether Nox enzymes are involved in the generation of ⋅O2, we used a peptide termed gp91ds-tat (33), which is derived from a gp91phox (now named Nox2) sequence in the region that interacts with the organizer protein p47phox, thus disrupting p47phox binding to and activation of associated catalytic Nox subunits [particularly Nox2 (34) but also Nox1] in vascular smooth muscle cells (VSMCs) (29, 3537). We found that in aged wild-type mice, ~50% of ⋅O2 formation was Nox-dependent as it was blocked by gp91ds-tat (Fig. 1A, left). In contrast to our expectation of exacerbated ⋅O2 production, ⋅O2 formation in aged Gper−/− mice was instead blunted by ~50 to 80% compared to wild-type mice (Fig. 1, A and B) and was unaffected by gp91ds-tat treatment (Fig. 1A), suggesting an inactive or absent Nox-dependent ⋅O2-producing pathway.

Fig. 1 Genetic deletion of Gper abrogates age-induced Nox activity and vascular dysfunction.

Intact arteries of aged (24-month-old) wild-type (Gper+/+) and Gper−/− mice were analyzed. (A and B) Nox activity was determined by measuring vascular superoxide (⋅O2) production using chemiluminescence (A) or DHE fluorescence (B; scale bars, 200 μm). To quantify the amount of ⋅O2 generated by Nox, subsets of arteries were treated with the Nox inhibitor gp91ds-tat (tat). (C and D) Endothelium-dependent, NO-mediated vasodilation in response to acetylcholine (C) and contractions to Ang II (D) in intact arteries in the presence or absence of gp91ds-tat (tat). Data are means ± SEM; n = 3 to 4 mice per group in (A), n = 5 to 10 mice per group in (B), and n = 4 to 5 mice per group in (C) and (D). *P < 0.05, **P < 0.01 compared to control (CTL); P < 0.05, ††P < 0.01, †††P < 0.001 compared to wild-type mice [analysis of variance (ANOVA) with Bonferroni post hoc tests in (A) and (D); repeated-measures ANOVA with Bonferroni post hoc tests in (C); Student’s t test in (B)].

We next determined the effects of aging on vascular tone, which is characterized by increased ⋅O2 formation that inactivates endothelium-derived vasodilatory NO (38). Impaired endothelium-dependent vasodilation represents an important predictor of mortality in patients with heart failure and hypertension (3841). As expected, NO-mediated endothelium-dependent relaxation induced by acetylcholine (42) was reduced in aged (Fig. 1C) compared to young wild-type mice (fig. S1A).This impairment was completely reversed by incubating arteries with gp91ds-tat, thus restoring vasodilation to an extent similar to that observed in young mice (Fig. 1C and fig. S1A). Further supporting the notion that impaired NO bioactivity was a result of oxidative stress in aged wild-type mice, VSMC sensitivity to NO alone, generated by an exogenous NO donor, was not affected by aging (fig. S2). In contrast, aged Gper−/− mice were completely protected from the impairment in endothelium-dependent vasodilation observed in aged wild-type mice; the vasodilatory capacity was preserved and identical to that of young mice (Fig. 1C and figs. S1A and S2).

In agreement with these observations, we found that in aged wild-type mice, vascular contractions in response to Ang II [a vasoactive peptide that stimulates Nox (43, 44)] were partially (~50%) blocked by gp91ds-tat (Fig. 1D), whereas gp91ds-tat had no effect on Ang II–mediated contractions of arteries from aged Gper−/− mice. In line with the reduced ⋅O2 formation in Gper−/− mice (Fig. 1, A and B, and fig. S1B), contractions triggered by Ang II were reduced by about 50% in aged (Fig. 1D) as well as in young Gper−/− mice (fig. S1C). These findings, which contrast with the protective vascular role of Gper expression and/or GPER stimulation reported in previous studies (4, 12, 13, 30, 45), indicate instead that constitutive Gper expression is essential for increased vascular Nox bioactivity as well as Nox-mediated vasoconstriction and impaired endothelium-dependent vasodilation particularly in the context of vascular aging.

Gper deletion prevents structural and functional cardiac aging and myocardial dysfunction

To determine whether Gper-dependent regulation of oxidative stress also plays a role in age-dependent structural and functional cardiac abnormalities, we next assessed ⋅O2 production in the aging heart. Compared to wild-type mice, myocardial ⋅O2 amounts were markedly lower in aged Gper−/− mice (Fig. 2A). Given that oxidative stress is centrally involved in the structural changes that occur with cardiac aging (25, 26), we next examined myocardial histopathology. Whereas aging increased the wall-to-lumen ratio of the left ventricle (LV) by ~60% in wild-type mice, Gper−/− mice were completely protected from age-dependent myocardial hypertrophy (Fig. 2B and fig. S3A). In addition, histological analyses of the myocardium of Gper−/− mice revealed an absence of cardiomyocyte hypertrophy (Fig. 2, C and D). Organ failure resulting from fibrosis accounts for at least one-third of deaths worldwide (46), with myocardial fibrosis being a key feature of cardiac aging (25, 26). Aging in wild-type mice was associated with prominent and diffuse interstitial myocardial fibrosis and collagen IV accumulation, which again were largely absent in aged Gper−/− mice (Fig. 2, C, E, and F). The cardioprotective effects of Gper deletion on myocardial fibrosis and hypertrophy were already detectable at 12 months of age (although the differences were less prominent because of the reduced disease pathology in the wild-type mice), resulting in a lower LV wall-to-lumen ratio (fig. S3A), reduced cardiomyocyte hypertrophy (fig. S3B), and reduced myocardial fibrosis, as assessed by Sirius Red (fig. S3C) and collagen IV (fig. S3D) staining, although the reduction in the former did not reach significance at this age.

Fig. 2 Obligatory role for GPER in aging cardiomyopathy and diastolic dysfunction.

Hearts of aged (24-month-old) wild-type and Gper−/− mice were analyzed. (A) Myocardial superoxide (⋅O2) production determined by DHE fluorescence. Scale bars, 500 μm. (B) Myocardial hypertrophy as measured by left ventricular (LV) wall-to-lumen ratio in cardiac cross sections. Scale bars, 2 mm. (C) Representative histological myocardial sections stained with hematoxylin and eosin (left, scale bars, 100 μm) and Sirius Red (right, scale bars, 200 μm). (D) Cardiomyocyte hypertrophy analyzed by cross-sectional area. (E) Quantitation of interstitial fibrosis. (F) Representative histological sections stained for type IV collagen (scale bars, 100 μm). (G to I) Dimensions and function of the left ventricle determined by echocardiography. Representative parasternal M mode images (G), calculated LV mass (H), and measures of LV filling pressures and diastolic dysfunction (E/e′ ratio; I) are shown. Data are means ± SEM; n = 4 to 5 mice per group in (A), n = 6 to 7 mice per group in (B), n = 6 to 8 mice per group in (C) to (F), and n = 3 mice per group in (G) to (I). *P < 0.05, **P < 0.01, ***P < 0.001 compared to wild-type mice (Student’s t test).

Given that Gper deletion prevented the structural cardiac abnormalities observed with aging, we next determined whether this translated into improved myocardial function in vivo. Echocardiography confirmed the marked increase in LV relative wall thickness and mass in wild-type mice compared to Gper−/− mice (Fig. 2, G and H, and table S1). Consistent with the reduced ventricular fibrosis and stiffness, analysis of LV filling and diastolic mitral valve annulus velocities (47) revealed improved diastolic function and lower LV filling pressures in aged Gper−/− mice (Fig. 2I and table S1). Together, the overall absence of myocardial fibrosis and hypertrophy in aged Gper−/− mice translated into increased ventricular elasticity, as indicated by improved LV diastolic filling. These differences were independent of changes in systolic LV function or systemic hemodynamics (table S1).

GPER is essential for ⋅O2 production in murine and human VSMCs through Nox1

Cardiac fibrosis involves an age-dependent localized activation of the renin-angiotensin system (41, 46, 48), with its primary vasoactive peptide Ang II also promoting premature senescence through the induction of Nox (49, 50). Moreover, Ang II–induced ROS promote redox-sensitive cell functions such as intracellular calcium mobilization and VSMC contraction (51). Having established that Gper deficiency abrogates Nox-mediated ⋅O2 production in aged mice, we next examined the underlying mechanisms of the molecular regulation in VSMCs isolated from wild-type and Gper−/− mice (fig. S4, A and B). Consistent with the activation of Nox in intact arteries of wild-type mice (Fig. 1), Ang II–stimulated ⋅O2 production in wild-type VSMCs was completely abrogated by gp91ds-tat (Fig. 3A, left). In cells lacking Gper, the Ang II–stimulating effect on ⋅O2 generation was completely absent (Fig. 3, A and B), which was confirmed by electron paramagnetic resonance (EPR) spectroscopy using 5-tert-butoxycarbonyl 5-methyl-1-pyrroline N-oxide (BMPO) as a spin trap for ⋅O2 (5254) (fig. S5). Similarly, Ang II–induced, Nox-dependent mobilization of intracellular calcium (51) was absent in Gper-deficient VSMCs (Fig. 3, C and D). By contrast, intracellular calcium mobilization responses to the purinergic receptor agonist adenosine triphosphate (ATP) [a Nox-independent stimulus (55)] were comparable in VSMCs from wild-type and Gper−/− mice, thus excluding inherent defects in calcium signaling in VSMCs lacking Gper (Fig. 3D). In addition, absence of Gper did not affect the expression of the genes encoding the Ang II AT1A and AT1B receptors (fig. S4C).

Fig. 3 GPER is required for Ang II–induced superoxide (⋅O2) production and mobilization of intracellular calcium in murine aortic VSMCs.

Nox activity stimulated by Ang II was inhibited using gp91ds-tat (tat). (A and B) Ang II–induced ⋅O2 production detected by DHE fluorescence (A) and chemiluminescence (B). (C and D) Mobilization of intracellular calcium ([Ca2+]i) determined using the Ca2+ sensor dye indo1-AM in response to Ang II [representative tracing in (C) and cumulative data in (D)] and ATP (D). F405/F490, ratiometric detection of emitted light at 405 and 490 nm. **P < 0.01, ***P < 0.001 compared to control (CTL); P < 0.05, ††P < 0.01, †††P < 0.001 compared to VSMCs isolated from wild-type (Gper+/+) mice. Data are means ± SEM; n = 5 to 7 independent experiments per group in (A), n = 5 independent experiment per group in (B), and n = 3 to 5 independent experiments per group in (D), all with VSMCs from two independent isolations. ANOVA with Bonferroni post hoc tests in (A) and (D); Student’s t test in (B).

We next sought to determine whether the effects of GPER on ⋅O2 production observed in murine aortic VSMCs extended to human aortic VSMCs. Knockdown of GPER with small interfering RNA (siRNA) abolished the ability of primary human VSMCs to generate ⋅O2 in response to Ang II (Fig. 4A). To determine whether the effects of GPER were mediated through rapid nongenomic signaling alone or involved long-term genomic effects, we treated human VSMCs with the GPER-selective antagonist G36, a synthetic, small-molecule GPER blocker (GRB) (32). Acute treatment (30 min) with gp91ds-tat, but not G36, abolished Ang II–stimulated ⋅O2 production (Fig. 4B). In contrast, prolonged treatment with G36 (for 72 hours) completely abrogated Ang II–induced ⋅O2 formation (Fig. 4B), suggestive of mechanisms regulating gene transcription. Consistent with the lack of acute effects, G36 did not display any direct antioxidant activity (fig. S6).

Fig. 4 GPER increases Nox1 abundance and activity in human and murine aortic VSMC.

(A) Ang II–induced superoxide (⋅O2) production detected by chemiluminescence in human VSMCs with GPER-targeted gene silencing (siGPER). *P < 0.05 compared to control siRNA (siCTL). siGPER treatment decreased GPER protein abundance by 89 ± 5% compared to siCTL treatment (representative blot shown in inset). (B and C) Ang II–induced ⋅O2 production (detected by chemiluminescence, B) and protein abundance of Nox1, Nox2, and Nox4 (C) in human VSMCs treated with the GPER-selective antagonist G36 (1 μM) for 30 min (acute, B) or 72 hours (chronic, B and C). Nox activity was inhibited using gp91ds-tat (tat). *P < 0.05, **P < 0.01 compared to control (CTL, DMSO 0.01%); P < 0.05 compared to acute treatment. (D and E) Protein (D) and mRNA (E) abundance of Nox1, Nox2, and Nox4 in VSMCs isolated from wild-type (Gper+/+) and Gper−/− mice. **P < 0.01 compared to wild-type mice. (F) Nox1 mRNA abundance in aorta and myocardium of aged (24-month-old) Gper+/+ and Gper−/− mice. *P < 0.05 compared to Gper+/+. (G) Ang II–induced ⋅O2 production detected by chemiluminescence in VSMCs isolated from Gper−/− mice and transduced with Nox1-expressing adenovirus (AdNox1). *P < 0.05 compared to vector control (AdGFP). Data are means ± SEM; n = 3 independent transfections per group in (A), n = 4 to 8 independent experiments per group in (B), n = 3 to 4 independent experiments and Western blots per group in (C) and (D), n = 3 to 4 VSMC preparations in (E), n = 4 to 10 mice per group in (F), and n = 3 independent transductions per group in (G). ANOVA with Bonferroni post hoc tests in (B); Student’s t test in (A) and (C) to (G).

Given that Nox inhibition by gp91ds-tat reduced vascular ⋅O2 formation in murine aorta ex vivo as well as in murine and human aortic VSMCs only in the presence of GPER, we next determined whether the vascular abundance of Nox1, Nox2, or Nox4 catalytic subunits, which have been implicated in both experimental and human arterial hypertension (2729), was affected by intrinsic GPER activity. Although gp91ds-tat is traditionally thought only to disrupt Nox2 activity (33), studies have demonstrated that gp91ds-tat also blocks ⋅O2-mediated effects by Nox1 in VSMCs, likely through its interaction with p47phox (56, 57). Thus, p47phox in VSMCs not only facilitates the activation of Nox1 (58), the closest homolog of Nox2 without affecting that of Nox4, but also mediates Ang II–dependent redox signaling (37, 59, 60). Feed-forward mechanisms have also been observed in which ROS production by one Nox subtype (or other sources) results in the activation of additional Nox subtypes, suggesting that inhibition of any intermediate could block downstream events (61, 62). We found that in human VSMCs treated with G36 for 72 hours, the protein abundance of Nox1 was reduced by ~70%, whereas that of Nox2 and Nox4 was unaffected (Fig. 4C). Similarly, only protein abundance of Nox1, but not that of Nox2 or Nox4, was substantially lower in murine VSMCs from Gper−/− mice as compared to wild-type mice (Fig. 4D). The reduced Nox1 protein abundance was commensurate with a similar reduction in mRNA abundance in VSMCs isolated from Gper−/− mice, with gene expression of Nox2 and Nox4 again being unaffected (Fig. 4E). Gper deficiency also reduced Nox1 mRNA abundance in the aorta and myocardium of aged Gper−/− mice (Fig. 4F), both of which displayed markedly reduced ⋅O2 bioactivity compared to wild-type mice (Figs. 1 and 2). To verify that the decreased Nox1 abundance accounted for the inability of Gper-deficient VSMCs to generate ⋅O2 in response to Ang II, we restored Nox1 in these cells using a Nox1-expressing adenovirus. Reintroduction of Nox1 into Gper-deficient VSMCs restored their capacity to generate ⋅O2 in response to Ang II (Fig. 4G), further suggesting an obligatory role for GPER in the maintenance of Nox1 abundance and its associated ROS-dependent cellular functions.

Genetic ablation or pharmacological inhibition of GPER prevents arterial hypertension

To explore whether the protective effects of Gper deletion extended to cardiovascular disease conditions other than those associated with aging, we increased Nox1 abundance and activity in vivo by infusing mice with Ang II, a critical inducer of Nox1-depdendent ⋅O2 production, vascular dysfunction, and increased vascular tone (44). Animals lacking Gper were resistant to the Ang II–induced increase in blood pressure observed in wild-type mice (Fig. 5A). Furthermore, vascular ⋅O2 generation and the increase in vascular Nox1 abundance in response to Ang II infusion required the presence of Gper (Fig. 5, B to D). As previously reported in wild-type mice (44), ⋅O2 generated in response to Ang II impaired endothelium-dependent vasodilation as evident from the blunted NO-dependent relaxation in response to acetylcholine. By contrast, the attenuation of the vasodilator response was completely absent in Gper−/− mice infused with Ang II (Fig. 5E). In line with the ⋅O2-mediated impairment of NO bioactivity, the inherent vascular smooth muscle sensitivity to NO, as determined with an exogenous NO donor, was unaffected by Ang II infusion, Gper deficiency, or GPER inhibition (fig. S7). These data further confirm that Gper is required to increase Nox1 abundance and the resulting ⋅O2 production, vascular dysfunction, and increases in vascular tone.

Fig. 5 Genetic or pharmacological ablation of GPER prevents Ang II–induced hypertension, oxidative stress, and increases in Nox1 abundance.

Wild-type (Gper+/+) and Gper−/− mice were infused with Ang II (0.7 mg kg−1 per day) or vehicle (control, CTL) for 14 days. A subset of wild-type mice was also treated with the GRB G36. (A) Systolic arterial blood pressure in conscious animals. (B and C) Vascular ⋅O2 generation as measured by chemiluminescence (B) and DHE staining (C; scale bars, 300 μm). (D) Vascular Nox1 protein abundance detected by immunofluorescence. (E) Endothelium-dependent, NO-mediated vasodilation in response to acetylcholine. Data are means ± SEM; n = 4 to 5 mice in (A), n = 3 to 5 mice in (B), n = 3 to 4 mice in (C), n = 3 mice in (D), and n = 5 to 9 mice in (E). *P < 0.05, **P < 0.01, ***P < 0.001 compared to genotype-matched CTL; P < 0.05, ††P < 0.01, †††P < 0.001 compared to Ang II–treated wild-type mice [repeated-measures ANOVA with Bonferroni post hoc tests in (A) and (E); ANOVA with Bonferroni post hoc tests in (B) and (D)]. PE, phenylephrine.

The Nox pathway has been recognized as a therapeutic target for ROS-dependent pathologies in humans (46, 63, 64). To determine whether decreasing Nox1 protein by pharmacological GPER inhibition can also be achieved in vivo, we again used the GRB G36 (32). Not only did G36 treatment prevent the Ang II–mediated increase in Nox1 protein abundance (Fig. 5D), it also normalized the increased vascular ⋅O2 production (Fig. 5, B and C) and restored the vasodilatory response (Fig. 5E). These effects of G36 resulted in a substantial inhibition of the Ang II–mediated increase in blood pressure (Fig. 5A). Given that GPER increased Nox1 protein abundance, these results identify GRBs as a member of a new class of drugs that act as Nox down-regulators.


The results presented in the current study demonstrate that inhibiting GPER activity conveys protection from myocardial and vascular diseases associated with increased Nox1-derived oxidative stress, including cardiovascular aging and arterial hypertension. These data may seem counterintuitive at first when compared to the current body of evidence suggesting the protective role of GRAs in the cardiovascular system (4). GRAs, such as G-1 (10), rapidly activate Nox-independent pathways thought to mediate salutary vascular effects, such as Akt and ERK (4). In particular, G-1, unlike the GRB G36, induces endothelial NO synthase phosphorylation and the subsequent generation of NO, which indirectly mediates antioxidant effects through inactivation of ⋅O2 (12). Thus, both GRBs and GRAs improve disease outcome by reducing ROS bioactivity through the same receptor, albeit through entirely distinct mechanisms. These findings place GPER at the center of the balance between the beneficial l-arginine–NO synthase pathway and harmful excessive ROS generation. Such dichotomous effects also exist for other hormone-receptor systems (such as insulin), with the ultimate (patho)physiogical effect dependent on the stage of the disease process (65).

Our results have identified new and important chronic functions of intrinsic GPER activity that determine Nox1 abundance. Regulation of gene expression through constitutive signaling is prevalent among GPCRs (17). The GPER-dependent increase in Nox1 abundance and Nox1-dependent ROS formation was likely a key factor in the pathogenesis of increased vasoconstriction observed with hypertension and aging, as well as age-dependent cardiac remodeling. Arterial hypertension, LV hypertrophy, and the associated diastolic dysfunction in humans are important predictors of the development and prognosis of heart failure (26), the prevalence of which increases with age and has reached epidemic proportions (66).

The present study has certain limitations. The results were obtained in non-human, experimental models of vascular and myocardial aging, heart failure, and arterial hypertension. Investigating the association between oxidative excess and cardiovascular injury in humans is complicated because most patients are already treated with ROS-inhibiting regimens (such as angiotensin-converting enzyme inhibitors or statins), which makes it difficult to unmask mechanisms as described in the present study (67). Moreover, whether human Nox enzymes such as Nox5, which is not present in the rodent genome (27, 63), are involved in these disease processes requires further study. Given the results from our in vitro studies with human aortic VSMCs it remains to be determined whether G36 inhibits Nox1 activity in humans, which would also require establishing the safety of GRB treatment. Because ROS are implicated in the progression of many chronic noncommunicable diseases, GRBs may find application in a broader array of indications.

In summary, this study has identified an obligatory role for the intrinsic activity of GPER as an activator of Nox1 expression that mediates structural and functional injury in vascular and myocardial diseases and hypertension. Although the role of ROS in the aging process and chronic diseases is well recognized, simple scavenging of ROS with antioxidants has been largely unsuccessful therapeutically (68), likely due to the highly localized production and ensuing effects of ROS. Inhibition of Nox activity through the use of small molecules, some with limited selectivity (69), is currently being evaluated in clinical trials. The present study, however, introduces a new class of drugs, Nox down-regulators, which, as shown for the GRB G36, reduce Nox protein abundance, thus directly limiting both ⋅O2 production and the abundance of one of its main cellular sources. Therapeutically reducing the expression of Nox could provide an effective approach to targeting chronic disease conditions involving excessive Nox-mediated ⋅O2 formation. Nox down-regulators such as GRBs may find therapeutic application in chronic diseases, such as arterial hypertension and heart failure, as well as other diseases with a high prevalence in the aged population, including chronic kidney disease and cancer, and even rare disorders of accelerated aging such as progeria (3941).


Study design

The aim of this study was to explore whether and through which mechanisms GPER regulates cardiovascular disease processes. For this purpose, we used both young and aged wild-type and Gper-deficient mice, as well as mice chronically infused with Ang II. In addition, the GRB G36 was used to test whether pharmacological targeting of GPER altered oxidative stress. Detailed mechanistic studies were carried out in primary VSMCs isolated from wild-type and Gper−/− mice, as well as in human VSMCs. Mice were randomly and equally assigned to different treatment groups. Animal studies were conducted in a controlled and nonblinded manner. Study designs included the following: (i) two-way factorial designs comparing Gper+/+ and Gper−/− mice, in the presence or absence of inhibitors; (ii) similar two-way factorial designs with repeated measures (concentrations, time); (iii) one-way factorial repeated-measures designs within the Gper+/+ cohort, controlling for baseline differences, over time; and (iv) two-way factorial designs with repeated-measures designs between the Gper+/+ and Gper−/− groups, controlling for baseline differences, over time.

Transgenic mice and aging model

Gper−/− mice (provided by J. S. Rosenbaum, Procter & Gamble Co.) were generated as previously described (12) and backcrossed 10 generations onto the C57BL/6 background (Harlan Laboratories). Wild-type C57BL/6 and Gper−/− mice were housed at the Animal Resource Facility of the University of New Mexico (UNM) Health Sciences Center under controlled temperature (22° to 23°C) on a 12-hour light-dark cycle with unrestricted access to standard chow and water. Male mice aged 24 months, which show functional and structural changes resembling human cardiovascular aging (25), were used as a model of aging. Animals were killed at 4, 12, or 24 months of age by intraperitoneal injection of sodium pentobarbital (2.2 mg g−1 body weight). All procedures were approved by and carried out in accordance with UNM institutional policies and the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals.

Chronic angiotensin II infusion model

Micro-osmotic pumps (Alzet model 1002, Durect) were implanted subcutaneously in the midscapular region of wild-type and Gper−/− mice under isoflurane (3%) anesthesia. Pumps continuously delivered phosphate-buffered saline (PBS) or Ang II (MP Biomedicals) at a rate of 0.7 mg kg−1 per day for 14 days (43, 44). Three days before pump implantation, pellets continuously releasing the GRB G36 (33 μg/day, Innovative Research of America) (32) or placebo were implanted subcutaneously into the right hindlimb of a subset of wild-type mice.

Vascular smooth muscle cells

Primary aortic VSMCs from wild-type and Gper−/− mice (n = 13 per genotype) were isolated and cultured as previously described (12). Human aortic VSMCs (Lonza) were cultured according to the provider’s recommendations. Experiments were performed with cells derived from passages 2 to 5 for murine and passages 2 to 8 for human VSMCs. For functional assays, cells at subconfluence were rendered quiescent by overnight serum starvation.

Measurement of ⋅O2 by lucigenin-enhanced chemiluminescence

After euthanasia, the aorta was immediately excised, carefully cleaned from perivascular adipose and connective tissue, opened longitudinally, and cut into segments of identical size (3 mm) in cold (4°C) physiological saline solution (PSS; 129.8 mM NaCl, 5.4 mM KCl, 0.83 mM MgSO4, 0.43 mM NaH2PO4, 19 mM NaHCO3, 1.8 mM CaCl2, and 5.5 mM glucose; pH 7.4). Tissues were transferred and equilibrated in Hepes-buffered PSS (134 mM NaCl, 6 mM KCl, 1 mM MgCl, 10 mM Hepes, 2 mM CaCl2, 0.026 mM EDTA, and 10 mM glucose; pH 7.4) in a humidified incubator at 37°C for 60 min. In addition to intact isolated arteries, VSMCs and a cell-free⋅O2 generating system [produced by adding the substrate xanthine (100 μM, Calbiochem) to xanthine oxidase (0.05 mU, Calbiochem)] were used. Chemiluminescence was measured in dark-adapted Hepes-PSS containing 5 μM lucigenin (Enzo Life Sciences) at 37°C (70). After equilibrating for 15 min, ⋅O2 production was induced by Ang II (100 nM) (70). Where indicated, tissues or cells were pretreated with the Nox-selective inhibitor gp91ds-tat (Anaspec, 3 μM) (28, 33), the GRB G36 (10 nM, 100 nM, or 1 μM) (32), the ⋅O2 dismutase mimetic tempol (100 μM, Tocris Bioscience) (71), or vehicle [dimethyl sulfoxide (DMSO) 0.01%]. Luminescence was measured 10 times in 20-s intervals using a Synergy H1 Multi-Mode Microplate Reader (BioTek), and readings were averaged to reduce variability (70). A background reading lacking Ang II was subtracted, and ⋅O2 production was normalized to the surface area of vascular segments (72) or to VSMC number (70).

In situ detection of ⋅O2 by dihydroethidium

The thoracic aorta was equilibrated in Hepes-PSS in a humidified incubator at 37°C for 60 min and treated with the Nox-selective inhibitor gp91ds-tat (3 μM) (28, 33) for 30 min when indicated. Tissues were frozen in Tissue-Tek optimum cutting temperature (O.C.T.) compound (Sakura Finetek), cut on a cryostat into 10-μm-thick sections, and stored on glass slides at −80°C. For staining, sections were incubated with dihydroethidium (DHE) (5 μM, Invitrogen) in Hepes-PSS for 15 min at room temperature in the dark (73). In separate experiments, VSMCs were grown on coverslips coated with poly–l-lysine, and incubated with DHE (5 μM) in Hepes-PSS for 30 min at 37°C in the dark (73). Where indicated, VSMCs were pretreated with the Nox-selective inhibitor gp91ds-tat (3 μM) (28, 33) for 30 min, and ⋅O2 production was stimulated by Ang II (100 nM) for 20 min before imaging. Slides with VSMCs or aortic sections were carefully washed, mounted in Hepes-PSS with coverslips, and immediately imaged by epifluorescence microscopy (Axiovert 200M, Zeiss) using a rhodamine filter with exposure intensity adjusted to background fluorescence (73). Signal intensity was quantified using ImageJ software (NIH).

⋅O2 detection by spin trapping combined with EPR spectroscopy

BMPO (Enzo Life Sciences) was used as the spin trap for ⋅O2 generated from VSMC, which was monitored using EPR spectroscopy as previously described (5254). Briefly, serum-starved VSMCs were suspended in serum-free medium supplemented with BMPO (50 mM) and diethylenetriaminepentaacetic acid (100 μM), and ⋅O2 production was stimulated by Ang II (100 nM, 30 min at 37°C). Supernatant containing spin-trapped ⋅O2 was snap-frozen in liquid nitrogen and stored at −80°C for less than 1 week. After thawing, the supernatant was immediately transferred to custom-made gas-permeable Teflon tubing (Zeus Industries) folded four times, and inserted into a quartz EPR tube open at each end. The quartz EPR tube was inserted into the cavity of an EPR spectrometer (EleXsys 540 X-band, Bruker) operating at 9.8 GHz and 100-kHz field modulation, and the spectra of BMPO-OOH were recorded after spectrometer tuning at room temperature. The EPR spectrum was acquired with a scan time of 40 s, and 20 scans were obtained and averaged to produce significant signal-to-noise ratio. Instrument settings were as follows: magnetic field, 3509 G; scan range, 70 G; microwave power, 21 mW; modulation frequency, 100 kHz; modulation amplitude, 1.0 G; time constant, 20 ms. The EPR spectra were collected, stored, and manipulated using Xepr software (Bruker).

Vascular reactivity studies

The aorta was immediately excised after euthanasia, transferred into cold (4°C) PSS, carefully cleaned from perivascular adipose and connective tissue, and cut into 2-mm-long rings. Aortic rings were mounted in myograph chambers (multichannel myograph system 620M, Danish Myo Technology) onto 200-μm pins (18, 74). A PowerLab 8/35 data acquisition system and LabChart Pro software (ADInstruments) were used for recording isometric tension. Experiments to determine the vascular reactivity of aortic rings were performed as previously described (18, 74). Briefly, rings were equilibrated in PSS (37°C; pH 7.4; oxygenated with 21% O2, 5% CO2, and balanced N2) for 30 min and stretched stepwise to the optimal amount of passive tension for force generation. Functional integrity of vascular smooth muscle was confirmed by repeated exposure to KCl (PSS with substitution of 60 mM potassium for sodium), with resulting contractions demonstrating no differences between groups. Selected arteries were pretreated with the Nox-selective inhibitor gp91ds-tat (3 μM) (28, 33) for 30 min. Contractions to Ang II (100 nM) were studied in the abdominal aorta in the presence of the NO synthase inhibitor l-NG-nitroarginine methyl ester (300 μM, incubation for 30 min, Cayman Chemical) (75) to exclude Ang II–mediated release of NO (76). Ang II–induced contractions exhibit rapid desensitization in the mouse vasculature with a nearly complete loss of tension after about 2 min, thus preventing the recording of responses to increasing concentrations (75, 77). To study endothelium-dependent, NO-mediated relaxations, rings from the thoracic aorta were precontracted with phenylephrine (Sigma-Aldrich) to 80% of KCl-induced contractions, and responses to acetylcholine (0.1 nM to 10 μM, Sigma-Aldrich) were recorded. Similarly, endothelium-independent, NO-mediated relaxations to sodium nitroprusside (1 to 10 μM, MP Biomedicals) were determined. Precontraction did not differ between groups. To exclude any GPER-dependent effects on vasoconstrictor prostanoids (18, 75), responses were obtained in the presence of the cyclooxygenase inhibitor meclofenamate (1 μM, incubation for 30 min, Cayman Chemical). Contractions were calculated as the percentage of contraction to KCl, and relaxation was expressed as the percentage of phenylephrine-induced precontraction.

Cardiac histopathology

After sacrifice, hearts were excised, fixed in 4% paraformaldehyde, and embedded into paraffin blocks. Histological sections (2 μm) were stained with hematoxylin and eosin, with Sirius Red, or by immunohistochemistry using a polyclonal goat antibody recognizing collagen IV (SouthernBiotech) as previously described (78). Morphometric analysis of free LV wall thickness and ventricular lumen area was performed using light microscopy at 400-fold magnification and cellSens software (Olympus), with LV wall thickness based on analysis of 10 randomly selected measure points. Cardiomyocyte cross-sectional area was determined by analysis of 15 anterolaterally located cardiomyocytes using cellSens software. Myocardial fibrosis on Sirius Red– or collagen type IV–stained paraffin sections was graded using a semiquantitative fibrosis score (0 = no staining, 1 = less than 25%, 2 = 26 to 50%, 3 = 51 to 75%, and 4 = more than 75% of cardiac tissue with positive staining). For each heart, the mean score evaluated on 10 fields at 200-fold magnification was calculated.

High-resolution, high-frequency echocardiography

Mice were lightly sedated using inhaled isoflurane anesthesia and placed on a heat pad to maintain body temperature, and echocardiography was performed using a Vevo LAZR photoacoustic imaging system (VisualSonics) using high-resolution, high-frequency ultrasound at 40 MHz (47, 79). Conventional B mode, M mode, pulsed-wave Doppler, and tissue Doppler images were acquired by an experienced, blinded operator to ensure a standardized, consistent technique, and LV dimensions were quantified as previously described (47, 79). LV ejection fraction was determined by speckle tracking–based wall motion analysis (47, 79, 80) using VevoStrain software (VisualSonics). Analysis of diastolic function included transmitral flow velocity waveforms obtained from pulsed-wave Doppler to calculate the ratio of early (E)-to-late (atrial, A) LV filling velocities (E/A ratio). In addition, mitral annulus diastolic velocity (e′ waves) obtained from pulsed-wave tissue Doppler imaging as well as the calculated E/e′ ratio were obtained as measures of diastolic function and LV filling pressures.

Measurement of arterial blood pressure

Systolic and diastolic blood pressure were measured in conscious mice using a volume-pressure recording noninvasive monitoring system (CODA-6, Kent Scientific) as previously described (12), which produces blood pressure readings with similar sensitivity and specificity to invasive measurements (12). This blood pressure measurement technique has been successfully applied in the chronic Ang II infusion model (43, 44).

Intracellular calcium mobilization

VSMCs were loaded with 5 μM indo1-AM (Invitrogen) and 0.05% pluronic F-127 (Invitrogen) in Hanks’ balanced salt solution (HBSS) supplemented with NaCl (150 mM), CaCl2 (2 mM), and Hepes (20 mM; pH 7.4) for 30 min at room temperature in the dark. Cells were washed and resuspended in HBSS (106 cells/ml), and calcium mobilization in response to Ang II (100 nM) and ATP (1 μM, Sigma-Aldrich) was determined ratiometrically using a λex value of 340 nm and λem values of 405 and 490 nm at 37°C in a QM-2000-2 spectrofluorometer (Photon Technology International).

VSMC transduction with Nox1 adenovirus or transfection with GPER-targeted siRNA

VSMCs from Gper−/− mice were plated at ~600,000 cells per T25 flask, washed in PBS, and infected with Nox1GFP or green fluorescent protein (GFP) control adenovirus constructs (59) at a multiplicity of infection of 400 overnight in low-serum (1% fetal bovine serum) Dulbecco’s modified Eagle’s medium. Cells were allowed to recover for 48 hours before experiments. Transduction efficiency was determined by GFP expression. For siRNA, human aortic VSMCs were transfected with siGPER (Dharmacon ON-Targetplus J-005563-08) or control (siCTL) siRNA (Dharmacon ON-Targetplus D-001810-02) using Lipofectamine 2000 (Invitrogen) for 6 to 8 hours in serum-free medium, washed, and returned to normal medium as described by the manufacturer. Subsequent experiments were performed 72 hours after transfection.

Quantification of gene and protein expression

RNA was extracted, reverse-transcribed, and analyzed using SYBR Green–based detection of amplified gene-specific cDNA fragments by quantitative polymerase chain reaction performed in triplicate as previously described (18). The following primer pairs have been used: 5′-CAT CCA GTC TCC AAA CAT GAC A-3′ (forward) and 5′-GCT ACA GTG GCA ATC ACT CCA G-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse Nox1 (GenBank ID: NM_172203.1); 5′-ACT CCT TGG GTC AGC ACT GG-3′ (forward) and 5′-GTT CCT GTC CAG TTG TCT TCG-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse Nox2 (GenBank ID: NM_007807.4); 5′-TGA ACT ACA GTG AAG ATT TCC TTG AAC-3′ (forward) and 5′-GAC ACC CGT CAG ACC AGG AA-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse Nox4 (GenBank ID: NM_015760.4); 5′-GCG GTC TCC TTT TGA TTT CC-3′ (forward) and 5′-CAA AGG GCT CCT GAA ACT TG-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse AT1A receptor (GenBank ID: NM_177322.3); 5′- TAT TTT CCC CAG AGC AAA GC-3′ (forward) and 5′-TGT TGC TTC CTT GTC CCT TG-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse AT1B receptor (GenBank ID: NM_175086.3); and 5′-TTC ACC ACC ATG GAG AAG GC-3′ (forward) and 5′-GGC ATG GAC TGT GGT CAT GA-3′ (reverse) for amplification of a specific cDNA fragment encoding mouse glyceraldehyde-3-phosphate dehydrogenase (GenBank ID: NM_008084.2), which served as the housekeeping control.

For determination of protein abundance by Western blot, VSMCs were lysed in NP-40 buffer supplemented with protease inhibitor (1 μg/ml), 10% SDS, 0.5% sodium fluoride, and 0.5% sodium orthovanadate. Lysates (20 or 40 μg) were loaded on 10% SDS–polyacrylamide gel electrophoresis gel (Thermo Scientific), blotted onto polyvinylidene fluoride membrane (Millipore), and blocked with 3% newborn calf serum in tris-buffered saline with Tween-20 (0.1%). Blots were incubated overnight at 4°C with primary antibodies recognizing Nox1 (Sigma-Aldrich), Nox2 (Boster), or Nox4 (Boster); washed; incubated with secondary horseradish peroxidase–conjugated antibodies (1:5000) for 1 hour at room temperature; and developed with SuperSignal West Pico Chemiluminescent substrate (Thermo Scientific). Blots performed in duplicate were imaged and quantified using ImageJ densitometry analysis software (NIH).

Immunofluorescence of Nox1 and GPER

Aortic sections frozen in Tissue-Tek O.C.T. compound were fixed in 4% paraformaldehyde, blocked, and permeabilized in PBS containing normal goat serum (3%) and Triton X-100 (0.01%, EM Science). Sections were incubated with rabbit antibody recognizing murine Nox1 (1:100, Sigma-Aldrich) or negative control immunoglobulin G (IgG) (1:100, Sigma-Aldrich) overnight at 4°C, washed, incubated with goat antibody recognizing rabbit IgG conjugated to Alexa Fluor 488 (Invitrogen) for 1 hour, washed, mounted in Vectashield (Vector Laboratories), and imaged using a Leica SP5 confocal microscope. Signal intensity was quantified using ImageJ software. VSMCs were stained with a rabbit antiserum recognizing murine GPER as previously described (12).

Statistical analysis

Statistical analysis of in vitro and in vivo experiments was performed using GraphPad Prism version 5.0 for Macintosh (GraphPad Software). When comparing two groups, the two-tailed, unpaired Student’s t test was performed. When comparing multiple groups, data were analyzed by two-way ANOVA, with repeated measures as appropriate, followed by the Bonferroni post hoc test to correct for multiple comparisons. Values are expressed as means ± SEM; n equals the number of independent animals or cell preparations used. Statistical significance was accepted at a P value of <0.05.


Fig. S1. Contribution of Nox activity to vascular reactivity in young (4-month-old) wild-type and Gper−/− mice.

Fig. S2. Endothelium-independent, NO-mediated vasodilation in young (4-month-old) and aged (24-month-old) wild-type and Gper−/− mice.

Fig. S3. Effect of Gper deletion on LV hypertrophy and fibrosis in adult mice.

Fig. S4. Determination of GPER protein in cultured VSMCs isolated from wild-type and Gper−/− mice.

Fig. S5. Ang II–induced ⋅O2 generation in VSMCs isolated from wild-type and Gper−/− mice.

Fig. S6. Effect of G36, a small-molecule GRB, on ⋅O2 generation in a cell-free system.

Fig. S7. Endothelium-independent, NO-mediated vasodilation in wild-type and Gper−/− mice with Ang II–induced hypertension.

Table S1. Echocardiographic measurements and hemodynamic parameters in aged (24-month-old) wild-type and Gper−/− mice.


Acknowledgments: We thank C. Hu, D. Cimino, M. Reutelshöfer, K. Schmitt, and S. Söllner for expert technical assistance; J. Weaver for assistance with EPR spectroscopy; and B. Deeley (FUJIFILM VisualSonics Inc.) for support with the echocardiography studies. We acknowledge K. K. Griendling and B. Lassègue (Emory University School of Medicine, Atlanta, GA, USA) for providing the Nox1/GFP adenovirus. We thank J. S. Rosenbaum (Procter & Gamble Co.) for providing the Gper−/− mice. Funding: This study was supported by the NIH (NIH R01 CA127731 and CA163890 to E.R.P.), Dedicated Health Research Funds from the UNM School of Medicine allocated to the Signature Program in Cardiovascular and Metabolic Diseases (to E.R.P.), the Swiss National Science Foundation (grants 135874 and 141501 to M.R.M. and grants 108258 and 122504 to M.B.), and the Interdisciplinary Centre for Clinical Research Erlangen, project F1 (to K.A.). E.R.P. was also supported by the UNM Comprehensive Cancer Center (NIH grant P30 CA118100). N.C.F. was supported by NIH training grant HL07736. The EPR core facility of the UNM Biomedical Research and Integrative Neuroimaging Center was supported by NIH grant P30 GM103400, and the UNM and UNM Comprehensive Cancer Center Fluorescence Microscopy Shared Resource was supported by NIH grant P30 CA118100 as detailed at Biostatistics support was provided by the UNM Clinical and Translational Science Center supported by NIH grant UL1 TR001449. Author contributions: M.R.M., N.C.F., C.D., and G.S. performed the experiments; J.B.A. synthesized the G36; M.R.M., N.C.F., C.D., G.S., M.B., and E.R.P. analyzed the data; M.R.M., N.C.F., K.A., M.B., and E.R.P. interpreted the results of the experiments; M.R.M., M.B., and E.R.P. prepared the figures and wrote the manuscript; all authors approved the final version of manuscript; M.R.M., M.B., and E.R.P. were involved in the conception and/or design of the research. Competing interests: M.R.M., G.S., M.B., and E.R.P. are inventors on a U.S. patent application for the therapeutic use of compounds targeting GPER. E.R.P. and J.B.A. are inventors on U.S. patent nos. 7,875,721 and 8,487,100 for GPER-selective ligands and imaging agents. All other authors declare that they have no competing interests.
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