Role of GPER in estrogen-dependent nitric oxide formation and vasodilation
Estrogens are potent regulators of vascular tone, yet underlying receptor- and ligand-specific signaling pathways remain poorly characterized. The primary physiological estrogen 17\xce\xb2-estradiol (E2), a non-selective agonist of classical nuclear estrogen receptors (ER\xce\xb1 and ER\xce\xb2) as well as the G protein-coupled estrogen receptor (GPER), stimulates formation of the vasodilator nitric oxide (NO) in endothelial cells. Here, we studied the contribution of GPER signaling in E2-dependent activation of endothelial NO formation and subsequent vasodilation. Employing E2 and the GPER-selective agonist G-1, we investigated eNOS phosphorylation and NO formation in human endothelial cells, and endothelium-dependent vasodilation in the aortae of wild-type and Gper-deficient mice. Both E2 and G-1 induced phosphorylation of eNOS at the activation site Ser1177 to similar extents. Endothelial NO production to E2 was comparable to that of G-1, and was substantially reduced after pharmacological inhibition of GPER. Similarly, the clinically used ER-targeting drugs 4OH-tamoxifen, raloxifene, and ICI182,780 (faslodex, fulvestrant\xe2\x84\xa2) induced NO formation in part via GPER. We identified c-Src, EGFR, PI3K and ERK signaling pathways to be involved in GPER-dependent NO formation. In line with activation of NO formation in cells, E2 and G-1 induced equally potent vasodilation in the aorta of wild-type mice. Gper deletion completely abrogated the vasodilator response to G-1, while reducing the response to E2 by ~50%. These findings indicate that a substantial portion of E2-induced endothelium-dependent vasodilation and NO formation is mediated by GPER. Thus, selective targeting of vascular GPER may be a suitable approach to activate the endothelial NO pathway, possibly leading to reduced vascular tone and inhibition of atherosclerotic vascular disease.
INTRODUCTION<!>Human endothelial cells<!>Quantitative real-time polymerase chain reaction (qPCR)<!>Fluorescence microscopy<!>Western blotting<!>Detection of NO<!>Animals<!>Vascular function studies<!>Materials<!>Statistical analyses<!>Expression and intracellular localization of GPER in human endothelial\ncells<!>GPER mediates phosphorylation of eNOS at Ser1177<!>E2 and G-1 mediate NO formation via GPER<!>GPER-dependent NO formation is induced by selective estrogen receptors\nmodulators and downregulators<!>GPER induces NO production in endothelial cells through c-Src, EGFR, PI3K and\nERK pathways<!>GPER contributes to endothelium-dependent arterial dilatation by E2<!>DISCUSSION
<p>Epidemiological studies show a lower incidence of coronary artery disease,
hypertension and stroke in premenopausal women compared to age-matched men; however,
these sex differences lessen with the onset of menopause [1, 2]. Such
observations suggest that the loss of ovarian sex hormones coincides with a loss of
protection against vascular diseases, implicating a beneficial functional role for
natural estrogen (predominantly 17β-estradiol, E2) in arterial health
[3–5]. In particular, endothelial cells, which form
the luminal cell monolayer of the vascular wall, have emerged as critical mediators
of estrogen's salutary effects in the cardiovascular system [3, 6,
7].</p><p>Endothelial cells release multiple vasoactive substances including the
endothelium-derived relaxing factor (EDRF) [8], which was identified as nitric oxide (NO) [9, 10]. The enzyme that catalyzes NO formation (utilizing L-arginine
and molecular oxygen as substrates) is NO synthase (NOS) [11, 12].
Three NOS isoforms exist, with one isoform (NOS III) being predominantly expressed
in endothelial cells and hence referred to as endothelial NOS (eNOS) [11, 12]. eNOS-derived NO is a potent vasodilator, but also conveys
vasoprotection through multiple mechanisms such as inhibition of leukocyte adhesion
and migration, platelet aggregation and thrombosis, as well as mitigating
proliferation and migration of the underlying vascular smooth muscle cells
[12]. E2 has been shown
to stimulate NO formation in cultured human endothelial cells [13]. In particular, E2 binding to
estrogen receptors (ER) triggers signaling cascades that include activation of the
kinases c-Src [14], ERK
[15, 16], PI3K [17], and Akt [18], the latter eliciting phosphorylation of the
eNOS activation residue Ser1177 [19–22]. Studies
in murine aortae have confirmed the involvement of these pathways in
endothelium-dependent, NO-mediated vasodilation induced by E2 [19, 23,
24]. Different ER subtypes (the
holoreceptor ERÎ± its truncated isoform ER46, and possibly ERÎ²)
localized to the plasma membrane have been associated with these actions
[25–27].</p><p>In 1997, an orphan G protein-coupled receptor termed GPR30 was cloned from
human endothelial cells exposed to fluid shear stress [28], a prototypic physiologic stimulus of eNOS
activation [29]. Following
the demonstration of binding and signaling in response to E2 [17, 30], GPR30 was renamed G protein-coupled estrogen receptor (GPER)
[31]. Experiments
utilizing GPER-deficient (Gper−/−) mice
[32] and GPER-selective
ligands, the agonist G-1 [33, 34] and antagonists G15 and G36
[35, 36], have established the vascular actions of
this third estrogen receptor in vascular biology [37–45] as well as many other aspects of physiology [46–51]. We and others have previously shown that G-1 induces eNOS
activation in human endothelial cells [52–54], as well
as endothelium-dependent, NO-mediated vasodilation of multiple human and rodent
arteries [3, 53, 55, 56]. These findings established GPER as
an ER mediating eNOS activation; however, the extent to which GPER contributes to
the overall E2-dependent response and the activation of eNOS by multiple
ER-targeting drugs remain unresolved. Hence, we set out to determine the role of
GPER in eNOS phosphorylation and the mechanisms of NO production in human
endothelial cells, as well as vasodilation in the aorta of wild-type and
Gper−/− mice. To this end, we
utilized the GPER-selective agonist G-1 [33] and the non-selective ER agonist E2, as well as the
selective ER modulators (SERMs) 4OH-tamoxifen and raloxifene and the selective ER
downregulator (SERD) ICI182,780 (faslodex, fulvestrant™), which have been
shown to act as GPER agonists in other systems [57].</p><!><p>Telomerase-immortalized human umbilical vein endothelial (TIVE) cells
were a generous gift from Rolf Renne, PhD (University of Florida, Gainesville,
FL), and their derivation has been described previously [58]. Sex of the cells was determined by
fluorescence in situ hybridization (FISH) analysis (TriCore
Reference Laboratories, Albuquerque, NM). Primary single donor human umbilical
vein endothelial cells (HUVEC) were obtained from Lonza (cat # C2517AS).
TIVE and HUVEC cells were cultured in M199 basal media supplemented with
20% FBS, 100 Î¼g/mL bovine neural-derived endothelial growth
factor and antibiotics (50 U/mL penicillin, 50 Î¼g/mL streptomycin). All
tissue culture vessels were coated for 30–60 min at room temperature
with 0.1% sterile gelatin in sterile milliQ-filtered water prior to
seeding cells. The expression pattern of endothelial cell-specific markers
remains unchanged from passages 2 to 12 in this cell line [58], and TIVE cells were used up to
passage 9 for experiments. HUVEC were used below passage 4.</p><!><p>Gene expression of GPER and eNOS in TIVE cells was analyzed at passages
3, 6, 9 and 12 and in HUVEC at passage 3. Total cellular RNA was extracted using
Trizol Reagent and eluted using Qiagen RNeasy Mini kit (Qiagen, Valencia, CA)
according the manufacturer's instructions. RNA (400 ng) was reverse
transcribed using the First Strand cDNA synthesis kit (Applied Biosystems,
Carlsbad, CA). SYBR green-based detection of amplified gene-specific cDNA
fragments was performed on a 7500 FAST real-time PCR system (Applied
Biosystems). Primer sequences are provided in Table 1. GAPDH served as a house-keeping control.</p><!><p>Cells were seeded onto gelatin-coated coverslips (approximately 30,000
cells per coverslip in a 24 well cell culture plate) for 48 hours and fixed in
PBS containing 4% paraformaldehyde for 15 min at room temperature. For
examining GPER localization, cells were treated with either permeabilizing (PBS
containing 3% BSA and 0.1% Triton X-100) or non-permeabilizing
(PBS containing 3% BSA) blocking buffer for 1 hour at room temperature,
and incubated with a rabbit anti-mouse GPER antibody targeting a sequence within
the second extracellular loop (acetyl-FADVREVQWLEVTLGFIC, 1:10,000)
[18] or negative
control pre-immune rabbit serum overnight at 4°C. Slides were then
washed 3 times with permeabilizing (0.1% Triton X-100) or
non-permeabilizing PBS, incubated with goat anti-rabbit IgG conjugated to Alexa
Fluor 488 (1:500) for 1 h at room temperature, washed three times with PBS, and
mounted in Vectashield supplemented with DAPI (200 ng/mL). To evaluate
ERÎ± and GPER staining in TIVE and HUVEC. cells were permeabilized and
incubated with 1:50 mouse IgG antibody targeting ERÎ± (Santa Cruz sc8002
clone F10) overnight at 4°C, washed three times with PBS-T and incubated
with 1:500 rabbit anti-mouse Alexa Fluor 488. Cells were co-stained using a GPER
antibody targeting the N-terminus [17] at 1:10,000 and 1:500 anti-rabbit Alexa Fluor 568. DAPI
was used as a nuclear counterstain described above. Fluorescence signals were
visualized using a Zeiss LSM510 Meta or Zeiss 720 confocal fluorescent
microscope.</p><!><p>TIVE cells (200,000 cells per well) were seeded and grown to
70–80% confluence in 6-well plates and serum starved overnight.
The next morning, cells were treated with the non-selective estrogen receptor
agonist E2 (100 nM), the GPER-selective agonist G-1 (1–100 nM) or
vehicle (DMSO 0.01 %) for up to 15 min [52], briefly washed with ice cold PBS and
lysed on ice with 50 Î¼L of ice cold NP-40 lysis buffer supplemented with
1% SDS, 5 Î¼M NaVO4 and 5 Î¼M NaF to preserve
phosphorylated proteins. Cell debris was pelleted at 12,000 ×
g at 4 °C and the supernatant of soluble proteins
was collected, aliquoted and stored at −80°C. Protein
concentrations were determined by Coomassie (Bradford) assay (Pierce, Rockfield,
IL). For each sample, 20 Î¼g of total protein was resolved by 10%
SDS PAGE and blotted to PVDF membrane. Blots were blocked for 1 hour at room
temperature in TBS-T 0.01% supplemented with 3% newborn calf
serum and incubated overnight at 4°C with mouse anti-human pSer1177-eNOS
(1:500, antibody 612392, BD Biosciences, Sparks, MD) or mouse anti-human
β-actin (1:10,000, antibody MAB1501, Millipore, Darmstadt, Germany)
antibodies. Blots were then washed, incubated with secondary HRP-conjugated
antibodies (1:5000) for 1 hour at room temperature, developed with PicoWest
Chemiluminescence detection (ThermoScientific) for one minute at room
temperature. Blots were imaged on Kodak X ray film and quantified using ImageJ
densitometry analysis software (National Institutes of Health).</p><!><p>NO was determined by detection of the stable NO metabolites
NO2/NO3 (Nitric Oxide Detection Kit, ab65328, Abcam,
Boston, MA). TIVE cells (300,000) were seeded onto 60 mm dishes. At 70%
confluence, cells were serum starved in HEPES-buffered physiologic saline
solution (HEPES-PSS, composition in mM: 134 NaCl, 6 KCl, 1 MgCl2, 2
CaCl2, 0.026 EDTA, 10 glucose, and 10 HEPES; pH 7.4) for three
hours prior to experiments. Cells were treated with either the non-selective ER
agonist E2 (100 nM), the GPER-selective agonist G-1 (100 nM), the SERMs
raloxifene (100 nM) and 4OH-tamoxifen (100 nM), the SERD ICI182,780 (100 nM),
acetylcholine (100 nM), or vehicle (DMSO 0.01%) for 15 min in HEPES-PSS
at 37°C. TIVE cells were lysed on ice with 80 μL Griess assay
sample buffer. Per the manufacturer's recommendation, we further lysed
each sample by pulling it through a 27.5 gauge needle, briefly pelleted cell
debris (3 min in at 13,000 rpm at 4°C) and collected the supernatants
for Griess assay. Samples were either stored overnight at −80°C
or immediately loaded into a 96-well plate, treated with nitrate reductase and
nitrate reductase cofactor, and incubated at room temperature for 90 min to
convert nitrates. Sequential addition of Griess reagents 1 and 2 was used to
develop azo purple for 5 min at room temperature. Spectrophotometric detection
of azo purple absorbance at 540 nm was performed on a Synergy H1 multimode
microplate reader (BioTek, Winooski, VT) at room temperature. Three readings per
plate were taken. Background signal was subtracted and net azo signal was
normalized to total protein (determined by Coomassie blue readings at 533 nm)
for each replicate. Samples were analyzed in triplicate for each experiment, and
at least three experiments were performed for each condition. As indicated,
selected experiments were repeated in the presence of the GPER-selective
inhibitor G36 (1 Î¼M), the EGFR inhibitor AG1478 (1 Î¼M), the
c-Src inhibitor PP2 (1 Î¼M), the PI3K inhibitor LY294002 (1 Î¼M),
or the ERK inhibitor PD98059 (1 μM) for 30 min at 37°C prior to
stimulation with E2 or G-1.</p><!><p>Gper−/− mice (Procter
& Gamble, Cincinnati, OH, provided by Jan S. Rosenbaum) were generated
and backcrossed onto the C57BL/6J background as described [52]. Male
Gper−/− and wild-type mice
(Harlan Laboratories, Indianapolis, IN) were bred and housed at the University
of New Mexico Animal Resources Facility under controlled temperature of
22–23°C on a 12 hour light-dark cycle. Mice were given
ad libitum access to water and a rodent diet devoid of
alfalfa or soybean meal to minimize the presence of natural phytoestrogens
(Teklad 2020SX, Harlan Laboratories, Madison, WI). All procedures were approved
by the University of New Mexico Institutional Animal Care and Use Committee and
in accordance with the National Institutes of Health Guide for the Care and Use
of Laboratory Animals.</p><!><p>At 3–5 months of age,
Gper−/− and wild-type mice were
sacrificed by intraperitoneal injection of 2 mg/g sodium pentobarbital. The
thoracic aorta was immediately excised, transferred to ice-cold PSS (in mM:
129.8 NaCl, 5.4 KCl, 0.83 MgSO4, 0.43 NaH2PO4,
19 NaHCO3, 1.8 CaCl2, and 5.5 glucose; pH 7.4), and
cleaned of fat and connective tissue to exclude perivascular adipose-dependent
contractile effects [59].
Vessels were cut into 3 mm long rings, and carefully mounted onto two 200
Î¼m pins of a Mulvany-Halpern myograph (620M Multi Wire Myograph System,
Danish Myo Technology, Aarhus, Denmark) to record isometric tension using a
PowerLab 8/35 data acquisition system and LabChart Pro software (AD Instruments,
Colorado Springs, CO). Vascular function experiments were performed as
previously described [59]. Briefly, functional integrity of the vascular smooth muscle
was assessed by repeatedly exposing vessels to KCl (PSS with equimolar
substitution of 60 mM potassium for sodium). The presence of an intact
endothelium was confirmed by precontracting vascular rings with phenylephrine (1
Î¼M), followed by exposure to acetylcholine (1 Î¼M) to induce
endothelium-dependent, NO-mediated relaxation. A relaxation response of
>80% was considered to represent an intact and functional
endothelium. Rings were then precontracted with prostaglandin
F2α to 30–40% of the KCl-induced
contraction, and the vasorelaxation response to E2 (3 Î¼M) or G-1 (3
Î¼M) was recorded for 50 min as described [38]. Ethanol at a final concentration of
0.1% served as solvent control. Experiments were conducted in the
presence of meclofenamate (pretreated with 1 Î¼M for 30 min) to exclude
effects of endothelial vasoconstrictor prostanoids [38]. Relaxation is expressed as the
percentage of precontraction to prostaglandin F2α.</p><!><p>Prostaglandin F2α, meclofenamate, AG1478, PP2,
LY294002, and PD98059 were from Cayman Chemical (Ann Arbor, MI). M199 medium was
from Gibco (ThermoFisher). G-1 and G36 were synthesized as described
[33, 36] and provided by Jeffrey Arterburn (New
Mexico State University, Las Cruces, NM). All other drugs were from
Sigma-Aldrich (St. Louis, MO).</p><!><p>When comparing multiple groups, data were analyzed using one-way or
two-way analysis of variance (ANOVA) with repeated measures as appropriate
followed by Bonferroni's post-hoc test. When comparing two groups, the
unpaired Student's t-test was used. All analyses were
performed using GraphPad Prism version 5.0 for Macintosh (GraphPad Software, San
Diego, CA). Values are expressed as mean of independent experiments, and error
bars represent SEM. n equals the number of animals or
independent experiments. A p value <0.05 was considered
statistically significant.</p><!><p>To study the role of GPER in stimulated endothelial NO release, we
employed a well-characterized hTERT-immortalized human
endothelial cell line, TIVE cells [58]. The cells were derived from male fetal umbilical cord
as demonstrated by FISH analysis for X and Y chromosome centromeres (Figure 1A). As GPER was originally cloned
from endothelial cells [28], we confirmed expression of its mRNA, which was unaffected
by limited serial passaging (Figure 1B).
However, since eNOS gene expression decreased between passages 9 and 12 in TIVE
cells (Figure 1B), all experiments were
performed in cells passaged less than 9 times. RNA expression of ERÎ±,
eNOS, and GPER in TIVE cells was found to be similar to primary HUVECs (Figure 1C). GPER protein staining employing
an antibody that recognizes a putative "extracellular loop"
[18] detected GPER in
TIVE cells only under permeabilizing (and not non-permeabilizing) conditions,
indicating that the majority of the protein is localized intracellularly in TIVE
cells (Figure 1D, E). Comparing GPER
staining to ERÎ± staining in both TIVE cells and HUVECs, the majority of
ERÎ± localized to the nucleus in both cell types, whereas GPER was
localized to membranes in the cytosol, with strong perinuclear localization
(Figure 1F, G). We therefore conclude
that GPER is predominantly localized to intracellular membranes in human
endothelial cells under steady-state conditions, similar to previous
observations in other cell types [17, 52], and that
GPER and ERÎ± are each similarly expressed in HUVEC and TIVE cells.</p><!><p>We next examined the effects of the non-selective ER agonist E2 and the
GPER-selective agonist G-1 [33] on eNOS activation by determining the phosphorylation status
of the critical activation residue Ser1177 in human endothelial cells. We
observed a G-1-mediated, dose-dependent increase in eNOS phosphorylation with a
3-fold increase in activation at 100 nM (p<0.05 vs.
vehicle, n=4–7), which was similar to eNOS
phosphorylation stimulated by 100 nM E2 (Figure
2).</p><!><p>We next sought to determine whether phosphorylation of eNOS by E2 and
G-1 translated into increased NO production. Relative to basal NO, non-selective
ER activation by E2 rapidly stimulated NO formation by 5-fold
(p<0.01 vs. vehicle,
n=4–6, Figure
3), with selective activation of GPER by G-1 yielding approximately
73% of the E2-induced response (p<0.01 vs.
vehicle, n=4–6, Figure 3). Pretreating cells with the GPER-selective antagonist G36
[36] reduced NO
production in response to E2 by 58%, and completely blocked the
G-1-induced response (both p<0.01,
n=4, Figure 3). By
comparison, activation of the NO pathway by the muscarinic M3
receptor agonist acetylcholine showed no difference in NO formation compared
with E2 (124±13% vs. 100±11%,
n=6–8, p=n.s.), and
was unaffected by GPER inhibition (n=5, Figure 3). These findings indicate that GPER alone can
mediate a substantial portion of the E2-induced eNOS activation response and NO
formation in human endothelial cells.</p><!><p>SERMs such as 4OH-tamoxifen and raloxifene as well as SERDs such as
ICI182,780 (faslodex, fulvestrant™) are classically thought to target
ERÎ±/ERÎ², yet more recent data identified these compounds as GPER
agonists [15, 57]. SERMs/SERDs are used in the treatment of
estrogen-sensitive cancer and osteoporosis, inducing both pro-estrogenic and
anti-estrogenic effects depending on the tissue and cell type [60]. We hypothesized that
SERMs/SERDs might also induce NO production through GPER activation in
endothelial cells. 4OH-tamoxifen, raloxifene and ICI182,780 stimulated NO
production to a similar extent compared to E2 (Figure 3). Acute inhibition of GPER using G36 reduced NO production
to 4OH-tamoxifen by 56% (n=3,
p<0.05), to raloxifene by 31%
(n=3, p=0.086), and to ICI
182,780 by 33% (n=3,
p<0.05, Figure 3).
These data establish that GPER contributes to endothelial NO formation induced
by SERMs and SERDs and extend our knowledge regarding the pharmacological
properties of these clinically approved drugs.</p><!><p>We next sought to determine the signaling pathway(s) involved in
GPER-mediated NO formation in human endothelial cells using inhibitors targeting
multiple pathways previously associated with GPER signaling, including c-Src,
trans-activation of the epidermal growth factor receptor (EGFR), PI3K and ERK1/2
[40]. c-Src has been
shown to directly phosphorylate eNOS at the activation residue Tyr83
[22, 61]. In endothelial cells, inhibition of
c-Src by PP2 reduced NO production in response to the GPER-selective agonist G-1
by 63%, and to the non-selective ER agonist E2 by 54% (both
n=4–6, p<0.01,
Figure 4A). Similarly, inhibition of
EGFR with AG1478 diminished NO production in response to G-1 by 62%, and
to E2 by 84% (both n=3–4,
p<0.01, Figure
4B). EGFR transactivation initiates a number of downstream secondary
messengers such as PI3K/Akt, which directly mediates eNOS phosphorylation at
ser1177 [21]. Indeed,
PI3K inhibition, by pretreatment of endothelial cells with LY294002, reduced NO
production in response to G-1 by 46%, and to E2 by 54%
(n=4, p<0.05, Figure 4C). An alternative signaling target
of GPER is the kinase ERK1/2, shown to activate eNOS at phosphorylation sites
Thr602 and Thr604 [62].
Inhibition of ERK1/2, by pretreatment with PD98059, reduced NO production in
response to G-1 by 59%, and to E2 by 48% (both
n=4, p<0.05, Figure 4D), identifying ERK as an additional
component of GPER-dependent NO formation in human endothelial cells. Taken
together, these data indicate that both non-selective ER (E2) and selective GPER
(G-1) activation of NO formation is mediated by c-Src, EGFR, PI3K and ERK
pathways.</p><!><p>Given the similar levels of E2- and G-1-induced activation of eNOS in
endothelial cell, we next studied the contribution of GPER to E2- and
G-1-induced, endothelium-dependent vasodilation in the aorta of wild-type and
Gper−/− mice. This vascular
bed displays dilation to E2 largely mediated by endothelial NO production
[63–66]. In WT mice, both the non-selective ER
agonist E2 and the GPER-selective agonist G-1 induced a time-dependent
vasodilation (60±10% and 76±7%,
n=3–8, both
p<0.001 vs. solvent, Figure 5A). Interestingly, E2 induced a rapid relaxation within 10
minutes that was more potent compared to G-1 (31±8% vs.
8±5% at 10 min, p<0.05), whereas the
sustained vasodilator response to E2 and G-1 was similar. Deletion of
Gper reduced E2-induced vasodilation by 48% (from
60±10% to 31±6%,
p<0.001, Figure
5A), indicating that a substantial portion of the overall vasodilator
response is mediated by GPER. As expected, the GPER-selective agonist G-1 did
not induce vasodilation in aortae from
Gper−/− mice, demonstrating
the specificity of G-1 towards GPER (Figure
5A). Furthermore, endothelium-dependent, NO-mediated vasodilation
induced by acetylcholine was similar in WT and
Gper−/− mice, excluding
inherent differences in endothelial function between the two genotypes
(92±2% vs. 90±3%,
n=4–8, p=n.s., Figure 5B). Together, these findings indicate
that E2 induces potent, endothelium-dependent vasodilation that is to a
significant extent mediated by GPER.</p><!><p>The present study demonstrates the following insights: (i) that activation
of GPER by SERMs and SERDs, as well as E2, is substantially involved in
eNOS-dependent NO production, (ii) that NO production by E2 and G-1 require the
activation of c-Src and EGFR as well as PI3K and ERK1/2, (iii) approximately
50% of the endothelium-dependent vasodilator response to E2 is dependent
upon GPER and (iv) the GPER-selective agonist G-1 does not induce relaxation in
Gper-deficient mice. Previous studies are consistent with our
findings that eNOS activation by GPER involves the PI3K/Akt and ERK1/2 pathways and
that vasodilation in response to G-1 involves a c-Src/EGFR-mediated pathway
[52, 54, 55].
Although multiple forms of ERÎ± have also been associated with E2-mediated
eNOS activation [25], our
current findings point towards an essential contribution of GPER to the overall
endothelium-dependent, NO-mediated vascular response to E2 and reveal new insights
regarding GPER in mediating the effects of clinically relevant ER-targeting
therapeutics. GPER agonists are known to activate multiple signaling pathways that
are also involved in E2-mediated NO production, including c-Src, EGFR
transactivation, PI3K, and ERK1/2 [21, 22, 61, 62].
Our results show that inhibition of these signaling pathways reduced NO formation in
response to both nonselective ER activation (E2) as well as GPER-selective
activation (G-1), consistent with recent studies of GPER-mediated activation of eNOS
[54]. These findings
extend previous observations that plasma membrane-associated subpopulations of
ERÎ± are capable of mediating E2-induced eNOS activation via c-Src/PI3K
[22], revealing an
alternative pathway via GPER.</p><p>This study enhances our understanding of the role of GPER in mediating the
effects of clinically used SERMs 4OH-tamoxifen and raloxifene, as well as the SERD
ICI182,780 in endothelial generation of NO. Since these compounds have previously
been shown to bind and activate GPER [14, 15, 17, 30],
our results reinforce the notion that SERMs or SERDs may have additional effects
outside of modifying ERÎ±/ERÎ² signaling. SERMs impart pharmacologic
effects by activation or inhibition of classic ERÎ±/ERÎ²; however,
SERM classification is specifically defined by their effects on ERÎ±,
regardless of their effects on GPER signaling. Conversely, ICI182,780, commonly
known for its inhibitory effects on ERÎ±/ERÎ², has been shown to
induce vasodilation in porcine epicardial arteries, an observation compatible with
GPER-dependent mechanisms [38]. Interestingly, a common clinical side-effect of ICI182,780 is
arterial hypotension [67],
possibly resulting from GPER-mediated endothelial NO formation. Furthermore, there
is a reduced incidence of adverse clinical events related to coronary artery disease
in randomized trials of SERMs in the treatment of breast cancer and osteoporosis
[60, 68, 69],
potentially resulting from GPER-mediated increases in NO formation that contribute
to the inhibition of atherosclerosis [52]. Thus, the discovery that clinically approved drugs are able
to activate GPER may require reconsideration of their therapeutic rational.</p><p>Vasodilator effects to E2 in men were first described in 1939 [70] and have since been confirmed
experimentally in numerous studies, with estrogen being particularly involved in the
regulation of endothelium-dependent, NO-mediated modulation of vascular tone. In
intact arteries, ERÎ± may be a principal mediator of rapid E2-induced,
NO-mediated, endothelium-dependent vasodilation [71], although more recent evidence suggests that
selective pharmacological activation of GPER may potently recapitulate these effects
[38, 72]. In line with previous reports [56, 73], we observed no difference between vasodilatory responses to E2
(non-selective ER activation) and G-1 (GPER-selective agonist). Furthermore, using a
genetically modified mouse model, we found ~50% reduction of the
relaxation response to E2 in GPER-deficient aortae. Collectively, these results
indicate that GPER is capable of mediating a substantial portion of E2-induced
vasodilatory responses independent of ERÎ±.</p><p>Although it may seem intuitive that the benefits of estrogen receptor
targeting would be greater in females, males may also benefit from ER-targeted
therapeutics. In this study, we utilized endothelial cells from male fetal umbilical
tissue and arteries from male mice, reinforcing previous observations indicating
that E2 potently affects vascular function in males [74]. The findings of the present study also
extend previous observations of a potent regulatory role of GPER on vasomotor tone
in males [53, 59, 75].
Given the absence of feminizing effects of G-1 [36, 76],
there may be pharmacologic utility in GPER-selective agonism via compounds such as
G-1 in promoting cardiovascular health in men as well as women.</p><p>The substantial contribution of GPER in E2-mediated eNOS activation further
supports the development of GPER-targeted compounds that may yield salutary vascular
effects by promoting eNOS activity, increasing NO bioavailability and reducing
vascular tone. Although large-scale clinical trials on hormone replacement therapy
with conjugated equine estrogens and medroxyprogesterone acetate in postmenopausal
women reported adverse effects on cardiovascular risk [77, 78],
subsequent re-analyses revealed decreased risk if therapy was initiated soon after
menopause [79, 80]. GPER activation has been shown to induce the
greatest vasodilation responses in the arteries of postmenopausal women
[53], suggesting that
older women may benefit from GPER-targeted therapy. In support of this, our group
has shown that G-1 treatment in mice partially abrogated diet-induced
atherosclerosis [52], a
disease that is closely linked to reduced NO generation by the endothelium
[12]. Thus, pharmacologic
modulation of GPER activity with small molecules such as G-1 could aid in promoting
or maintaining cardiovascular function in post-menopausal women, given the lack of
feminizing side effects of GPER-selective compounds on uterine growth observed in
response to E2 [52]. We
conclude therefore that GPER may mediate multiple protective cardiovascular effects
by promoting eNOS activation and NO production, and that this mechanism may
contribute to the salutary vascular effects of estrogen signaling.</p>