Estrogen regulates histone deacetylases to prevent cardiac hypertrophy

Angiotensin II and E2/ERβ regulate class II HDAC production

We first determined whether the hypertrophic peptides AngII and ET-1, and E2 or an ERβ-selective agonist, dipropylnitrile (DPN), modulate HDAC4 and 5 expression. In cultured neonatal rat cardiomyocytes, AngII or ET-1 (100 nM) strongly inhibited mRNA expression of both HDACs, determined by quantitative PCR (qPCR). However, relevant concentrations of E2 and DPN comparably and significantly prevented these actions of AngII and ET-1 but had little effect in the absence of AngII (Figure 1A). Protein levels of HDAC4 and 5 were also suppressed by AngII or ET-1, and the estrogenic compounds nearly completely prevented inhibition of HDAC protein production caused by the hypertrophic peptides (Figure 1B). These results also support ERβ as mediating estrogen actions. To further implicate ERβ, we knocked down ERα or ERβ with small interfering RNA (siRNA; validation of the protein knockdowns from the siRNAs is shown in Supplemental Figure S1, A and B). Only the knockdown of ERβ prevented the reversal by E2 of AngII inhibition of HDAC proteins (Figure 1C, lanes 2, 3, 5, and 6). These results indicate that in our model, E2/ERβ serve as derepressors of the antihypertrophic class II HDACs rather than as stimulators of their production. This occurs in the setting of prohypertrophic peptides that are implicated in human cardiovascular diseases.

HDAC phosphorylation and subcellular localization are regulated by AngII and E2/ERβ

It is well established that AngII causes the phosphorylation of class II antihypertrophic HDACs at specific serine residues. As a result, these deacetylases are exported from the nucleus to cytoplasm, where they no longer serve antihypertrophic functions (Zhang et al., 2002; Backs et al., 2006; Haberland et al., 2009). For HDAC4, hypertrophic agent signaling through calcium and calcium/calmodulin-dependent protein kinase II (CaMKII) causes Ser-632 phosphorylation, resulting in nuclear exclusion of the deacetylase (Backs et al., 2006). Here we found that AngII and ET-1 each stimulated HDAC4 S632 phosphorylation in whole-cell lysates, but this was significantly prevented by E2 or DPN (Figure 2A). HDAC4 total protein levels were unchanged, as phosphorylation was determined at 15 min of cell incubation.

We also determined the phosphorylation of HDAC4 in the intact cell. Cardiomyocytes were incubated with AngII ± E2 or DPN and microscopically visualized by immunofluorescence. Using a phospho-S632 HDAC4 specific antibody, we found that AngII caused increased Ser-632 phosphorylation and localized the modified HDAC4 to the cytoplasmic/perinuclear compartment (Supplemental Figure S2A). In contrast, E2 and DPN each inhibited AngII-induced HDAC4 phosphorylation.

To further understand the effect of phosphorylation for HDAC4 cell localization, we carried out similar short-exposure experiments and determined phospho-HDAC4 levels in cytoplasmic and nuclear cell fractions by immunoblot (Figure 2B). Compared to control, in which most HDAC4 was not phosphorylated and was found in the nucleus, AngII stimulated the trafficking of phosphorylated HDAC4 to cytoplasm. Because nuclear (nonphosphorylated) HDAC4 protein was markedly reduced by AngII exposure, we propose that most HDAC4 undergoes this posttranslational modification in response to the hypertrophic peptide. Validation of subcellular fraction purity is shown in Supplemental Figure S3A. In contrast, E2 and DPN markedly reduced the amount of phosphorylated HDAC4 (Figure 2B) and derepressed protein production (Figure 1B), resulting in relocalization to and enhanced expression of this enzyme in the nucleus.

We also determined AngII and ERβ interactions affecting another class II deacetylase, HDAC5. Ser-259/498 phosphorylations within this deacetylase protein occur from protein kinase D (PKD) activation and result in HDAC nuclear-cytoplasmic trafficking (Backs and Olson, 2006). As seen in Figure 2C, AngII and ET-1 each caused enhanced phosphorylation of HDAC5 at Ser-498. Phosphorylation was markedly reduced from concomitant exposure of the cardiomyocytes to either E2 or DPN. From subcellular fractions, AngII also caused Ser-259 phosphorylation of HDAC5 and the relocation of the modified deacetylase from nucleus to cytoplasm compared with control cells (Figure 2D). However, coexposure of the cells to E2 or DPN significantly reversed these effects of AngII, resulting in more HDAC5 localization in the nuclear compartment. To further support these findings, we visualized whole cells by microscopy. As seen in Supplemental Figure S2B, AngII induced the phosphorylation of HDAC5, and the phospho-HDAC primarily resided in the perinuclear (cytoplasmic) area, but phosphorylation was prevented by E2/ERβ. Therefore, both class II HDACs respond to hypertrophic factors with the expected phosphorylations and relocalization to cytoplasm. As a novel mechanism, E2/ERβ prevents both the specific phosphorylations and resulting nuclear exclusion of both class II HDACs, consistent with the antihypertrophic functions of estrogen (Pedram et al., 2005, 2008).

Signaling mechanisms that regulate HDAC phosphorylation

HDAC4 is phosphorylated predominantly through CaMKII in response to hypertrophic factors (Backs et al., 2006), and so we determined whether E2/ERβ inhibited this kinase. Calcium is an early and important signal for CaMKII activation, and we measured calcium flux by modified Fura-2 fluorescence in the cardiomyocytes. AngII stimulated a rapid and sustained increase in calcium over the length of the experiment, significantly and comparably reduced by E2, DPN, or a second ERβ-specific agonist, β-LGND2 (Yepuru et al., 2010; Figure 3A). We postulated that E2/ERβ might cause inhibition of calcium flux through a Gαi-dependent mechanism. Estrogen stimulates the Gαi subunit (Kumar et al., 2007), and Gαi inhibits calcium flux (Zhao et al., 2010; Wang et al., 2011). We found that pertussis toxin, which inhibits estrogen activation of Gαi (Kumar et al., 2007), strongly reversed E2 inhibition of AngII-induced calcium flux (Figure 3B).

To implicate calcium in the regulation of CAMKII, we performed an in vitro kinase assay in which CaMKII was immunoprecipitated from cardiomyocytes under various conditions and incubated with HDAC4 substrate protein. Phosphorylation of HDAC4 at S632 (a CaMKII phosphorylation site) was then determined by specific immunoblotting. AngII caused enhanced calcium-dependent, CaMKII-induced S632 phosphorylation of HDAC4, since the calcium chelators 1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) and ethylene glycol tetraacetic acid (EGTA) suppressed this action (Figure 3C). Thus E2/ERβ inhibition of calcium, probably through Gαi signaling, accounts for the sex steroid receptor inhibiting CaMKII-induced HDAC4 phosphorylation and resulting nuclear exclusion (Figures 2A and 3).

Providing support for this idea, we performed additional HDAC4 S632 phosphorylation experiments. We first showed that AngII stimulation of HDAC4 S632 phosphorylation in the cardiomyocytes at 60 min was significantly prevented by coincubation with a soluble inhibitor of CaMKII, KN93 (Figure 4A). We also knocked down this kinase with specific siRNA (siRNA validation in Supplemental Figure S1C) and found a significant loss of HDAC4 S362 phosphorylation. These results strongly implicate CaMKII for AngII action in this respect. We then incubated the cardiomyocytes with AngII ± E2 or DPN, immunoprecipitated the CaMKII from each condition, and added equal protein amounts of the kinase to HDAC4 substrate protein for in vitro kinase activity determination. E2 or DPN comparably caused a significant inhibition of CaMKII-dependent, AngII-induced HDAC4 S362 phosphorylation (Figure 4B)

We also determined HDAC5 phosphorylation as regulated by prohypertrophic peptides and antihypertrophic steroids. Phosphorylation of HDAC5 at S259 occurs by hypertrophic factor signaling through protein kinase Cδ (PKCδ) and PKD. It has been reported that S498 phosphorylation results from CaMKII activity, but this has not been consistently found (Zhang et al., 2002; Backs et al., 2006; Backs and Olson, 2006; Kong et al., 2006). We therefore investigated the former pathway.

In cultured cardiomyocytes, AngII caused significantly enhanced activation of PKCδ, reflected as Tyr-311 phosphorylation, which was prevented by E2 or DPN (Figure 4C). AngII also caused activating phosphorylations of PKD (S744/748), which were comparably and significantly inhibited by estrogenic compounds (Figure 4D). Furthermore, PKCδ siRNA prevented PKD phosphorylations as induced by AngII, linking the two kinases (validation of specific PKC siRNA(s) in Supplemental Figure S1, D and E, where PKCβ serves as a negative control, and Figure 4C). However, calcium chelation with BAPTA did not affect the ability of AngII to stimulate PKCδ or PKD activity (Supplemental Figure S3B). As shown in Figure 4E, S259 HDAC5 phosphorylation was stimulated by AngII and inhibited by E2, DPN, or a PKD inhibitor (CID 755673; validated in Figure 4D). PKCδ or PKD1 knockdown with siRNA also significantly impaired AngII-induced HDAC5 phosphorylation (PKD protein inhibition by siRNA is shown in Supplemental Figure S1F). Our results indicate novel functions of E2/ERβ to inhibit the linked pathways from Gαi, calcium, and CaMKII to HDAC4 phosphorylation and from PKC to PKD, which is important for HDAC5 phosphorylation, with phosphorylations of both deacetylases resulting from hypertrophic peptide actions. These results indicate a new mechanism for endogenous steroids to prevent exclusion from the nucleus of class II HDAC(s), underlying the antihypertrophic functions of estrogen.

Functional roles of class II HDACs

An important transition during the development of cardiac hypertrophy is switching of the myosin heavy chain (MHC) isoform from α-MHC in the normal heart to β-MHC in the hypertrophied organ (Lompre et al., 1979). This change embodies reorganization of the ventricle and sarcomere. We previously showed that AngII stimulates this gene/protein isoform transition in the heart and that estrogen significantly prevents this hypertrophic factor action (Pedram et al., 2005, 2008). Estrogen signaling from the membrane receptor pool has been shown by others and us to modulate the epigenetic regulation of transcription, most often working in conjunction with the nuclear ER pool (Pedram et al., 2012). We therefore determined whether AngII induces acetylation of histone 3 at the βMHC promoter, an effect that likely results from the exclusion of class II HDACs in the nucleus (Figure 2, B and D), thereby contributing to the development of cardiac hypertrophy.

Using cultured cardiomyocytes and chromatin immunoprecipitation (ChIP), we found that AngII caused significant acetylation of histone 3 (H3) at the β-MHC promoter, an epigenetic mark that strongly correlates with induction of genes (Zhang et al., 2002; Suganuma et al., 2010; Figure 5A, top). H3 acetylation induced by AngII also correlated with increased βMHC mRNA expression stimulated by the hypertrophic peptide (Figure 5A, bottom). E2 and DPN significantly blocked all of these effects of AngII. These results are consistent with the ability of E2/ERβ to prevent the nuclear exclusion of HDAC4 and 5 shown here.

To determine in vivo relevance to our studies, we used material from investigations of cardiac hypertrophy that we recently reported (Pedram et al., 2013). Wild-type (WT) and ERβKO ovariectomized female mice were infused by osmotic minipump with AngII over 4 wk in the absence or presence of coinfused β-LGND2 or an E2 pellet inserted under the skin. The mice were then examined for multiple cardiac parameters. E2 or β-LGND2 substantially prevented AngII-induced hypertension, cardiac hypertrophy, and cardiac fibrosis (Pedram et al., 2013). Here, using DNA from the left ventricles of WT mice, we determined that AngII stimulated H3 acetylation at the β-MHC promoter by ChIP assay, which was significantly inhibited upon coinfusion of the mice with β-LGND2 or exposed to E2 (Figure 5B, top). However, estrogenic compound effects were active in the WT but not the ERβKO mouse, consistent with mediation of estrogen's antihypertrophic effects through ERβ. These results correlate with βMHC gene expression in the ventricles of WT (but not ERβKO) mice exposed to the same conditions (Figure 5B, bottom). Thus the epigenetic and genetic data are consistent and correlate with the regulation of class II HDAC function/phosphorylation and cell localization.

To further support the latter conclusions in vivo, we determined the phosphorylation of HDAC4 at S632 in ventricles from the various mice. As mentioned, phosphorylation at this residue leads to nuclear exclusion of HDAC4. In WT mice, AngII infusion resulted in an increased phosphorylation of HDAC4 S632, which was significantly prevented by E2 or β-LGND2 administration (Figure 5C). However, the estrogenic compounds opposed only phosphorylation stimulated by AngII in WT mice.

Modulation of HDAC2 by AngII and E2/ERβ

Class I HDACS such as HDAC2 have been strongly implicated as prohypertrophic enzymes, promoting the transcription of genes such as Nppa and Myh7 (Trivedi et al., 2007) that are associated with the cardiac phenotype (Backs and Olson, 2006; Trivedi et al., 2007). We determined whether AngII and E2/ERβ regulate this HDAC. In cultured cardiomyocytes, AngII significantly stimulated HDAC2 mRNA expression (Figure 6A) and protein (Figure 6B) at 24 h, each significantly inhibited by E2 or DPN. We also investigated the phosphorylation and cell localization of the deacetylase, since HDAC2 is an effective hypertrophy-promoting protein when it is phosphorylated in the nucleus of the cardiomyocyte (Kee et al., 2006; Trivedi et al., 2007).

As shown in the cell images of Supplemental Figure S2C, AngII stimulated the activating phosphorylation of HDAC2 at Ser-394 (S394) that is seen in the nucleus. In addition, E2 and DPN significantly diminished the phosphorylation of nuclear HDAC2. To confirm the cell images, we carried out immunoblotting of HDAC2 S394 phosphorylation in cytosolic and nuclear fractions. AngII and ET-1 significantly stimulated the nuclear HDAC phosphorylation that E2 and DPN inhibited (Figure 6C). Of interest, under all conditions, total HDAC2 remained in the nucleus. Thus E2/ERβ does not cause intracellular translocation of the prohypertrophic HDAC2 protein, in contrast to the steroid effects on class II HDACs.

Creatine kinase (CK2) phosphorylates HDAC2 at S394, resulting in enhanced activity of the enzyme (Eom et al., 2011). CK2 has been reported to be constitutively active in some cells, but we clearly find that in cardiomyocytes, AngII and ET-1 each causes an increase in cytoplasmic CK2 activity, determined as precipitated kinase from cytosolic fraction activity against a substrate protein in vitro (Figure 6D). Furthermore, a kinase family member, CK1, has also been considered to be constitutively active, but recently was found to be regulated by Wnt signaling (Cruciat et al., 2013). The activation of CK2 by AngII was unaffected by calcium chelation with BAPTA (Supplemental Figure S3C). E2 and DPN significantly inhibited AngII and ET-1 activation of the kinase, which was also blocked by the soluble CK2 inhibitor tetrabromobenzotriole (TBB; Kramerov et al., 2011; Figure 6D). These findings correlate with the ability of TBB, E2, and DPN to also block AngII- and ET-1–induced phosphorylation of HDAC2 at S394 at 1-h cell exposure (Figure 6E), implicating CK2 as a hypertrophic factor modulating HDAC2 activity (Kramerov et al., 2011).

We then determined the subcellular phosphorylation and localization of CK2 protein in the cardiomyocytes. AngII stimulated the expression of CK2 protein in the nucleus of the whole cell, which was substantially reduced by E2, DPN, or the CK inhibitors TBB and apigenin (also validating the function of the latter; Supplemental Figure S2D). To determine whether AngII promoted the trafficking of the CK2 protein to the nucleus, we fractionated the cardiomyocytes after 2-h exposure to peptide and steroids. AngII stimulated the trafficking of this protein kinase from cytoplasm to the nucleus compared with the control condition (Figure 6F). This is consistent with the enhanced phosphorylation of nuclear HDAC2 by CK2 upon exposure of the cells to AngII (Figure 6E). The increased trafficking of this kinase to the nucleus was inhibited by E2, DPN, and, of interest, the CK2 inhibitors TBB and apigenin. The latter observation suggests the novel concept that activation of CK2 by AngII, as we show in cytoplasm (Figure 6D), may promote nuclear localization of the kinase and subsequent phosphorylation of HDAC2. It is also possible that inhibition of CK2 activity by estrogenic compounds, TBB, or apigenin promotes nuclear exclusion of the kinase (Figure 6F). Thus it is CK2 and not HDAC2 that traffics in response to AngII but is inhibited by E2/ERβ. There is little knowledge on 1) mechanistically how any substance activates CK2, 2) the structural requirements and protein–protein interactions for CK2 to traffic in or out of the nucleus, and 3) the upstream signaling by membrane E2/ERβ that affects all this, thus illustrating the need for future extensive investigation.

To investigate the functional roles of HDAC2, we incubated the cardiomyocytes with AngII ± estrogenic compounds and determined enhanced protein synthesis, a hallmark of myocyte hypertrophy (Pedram et al., 2005). AngII significantly stimulated leucine incorporation, which was inhibited by E2, DPN, or β-LGND2 (Figure 7A). To implicate HDAC2, the same study was done in cardiomyocytes transfected with control or HDAC2 siRNA or a dominant-negative (DN) HDAC2 plasmid (pcDNA3-HDAC2 S394A) 24 h before AngII exposure. HDAC2 siRNA resulted in a substantial decrease in the endogenous protein (Supplemental Figure S1G) and significantly prevented AngII stimulation of protein synthesis in the cells (Figure 7B). Expressing the DN mutant HDAC2 in cardiomyocytes (Eom et al., 2011; validated in Supplemental Figure S4) also strongly inhibited AngII-stimulated leucine incorporation (Figure 7B). Cell area, another parameter of hypertrophy, was also determined. AngII significantly increased cardiomyocyte area, which was significantly prevented by E2, DPN, or β-LGND2 (Figure 7C). Furthermore, this effect of AngII was also reversed by siRNA to HDAC2 or expression of the dominant-negative deacetylase. These results indicate that HDAC2 plays a significant role in the ability of AngII to stimulate in vitro cardiomyocyte hypertrophy, which is opposed by estrogenic compounds acting through ERβ.

To validate some of these key findings in vivo, we determined CK2 and HDAC2 activity in WT and ERβKO mouse ventricles. As shown in Figure 8A, AngII stimulated and E2 and β-LGND2 prevented CK2 activation, the latter only in WT mice. For HDAC2 S394 phosphorylation, AngII stimulated this phosphorylation in all mice, but E2 and β-LGND2 prevented this only in WT mouse ventricles (Figure 8B). HDAC2 protein abundance was also stimulated by AngII in all mice but singularly inhibited by E2 and β-LGND2 in WT mouse hearts (Figure 8C).

Transcriptional targets for HDAC2

HDAC2 promotes hypertrophy by activating transcription factors and hypertrophic genes and inhibiting antihypertrophic genes. We determined the ability of AngII and E2/ERβ to regulate representative genes in cardiomyocytes. GATA4 is an important transcription factor (TF) and is activated by hypertrophic factor signaling to promote cardiac hypertrophy (Zhou et al., 2012). AngII stimulated the increased mRNA and protein expression of GATA4 in cultured cardiomyocytes, which was prevented by E2 or DPN (Figure 9A). Furthermore, the transcriptional activity of this TF was stimulated by AngII, determined from transfection of a GATA4-luciferase reporter into cardiomyocytes (Figure 9B). GATA4 activity was prevented by HDAC2 siRNA or from coincubation of the myocytes with E2 or DPN. We then determined the response of a hypertrophic gene, ACTA1 (Trivedi et al., 2007), and found that AngII stimulated the expression of this mRNA, dependent upon HDAC2; E2 and DPN also significantly inhibited this action of AngII (Figure 9C). Thus AngII stimulated GATA4 production and activity, the latter dependent upon HDAC2, which was inhibited by E2 or DPN. ACTA1 was shown to be a target for GATA4 (Bisping et al., 2012), and we sequentially linked AngII, HDAC2, and GATA4 to ACTA1 expression.

As an inhibitor of cardiac hypertrophy, the Inpp5f gene codes for a phosphatase that represses AKT activity in cardiomyocytes (Trivedi et al., 2007). AngII inhibited the expression of the Inpp5f mRNA in cardiomyocytes in an HDAC2-dependent manner, but this was derepressed by coincubation of the cardiomyocytes with E2 or DPN (Figure 9D). AngII signals through AKT to cause an inhibitory phosphorylation of the glycogen synthase kinase 3β (GSK3β) at Ser-9, lifting the repressive phosphorylation by GSK3β of the GATA4 transcription factor and thereby promoting cardiac hypertrophy (Morisco et al., 2001; Antos et al., 2002). Here we implicate HDAC2 in the ability of AngII to repress the Inpp5f mRNA, providing a mechanism by which AngII stimulates AKT to inhibit GSK3β activity and therefore inducing GATA4-mediated hypertrophic gene expression, such as ACTA1 (Figure 9C). We recently showed that ERβ inhibits AngII-induced AKT in cardiomyocytes, in part by derepressing Inpp5F (Pedram et al., 2013). Here we suggest the ability of E2 and DPN to inhibit HDAC2 as a novel mechanism to limit AngII hypertrophic effects that result from GATA4 activation and Inpp5F suppression (Figure 9, B and D).

Cell culture and reagents

Neonatal rat cardiomyocytes were isolated as described (Pedram et al., 2005) and cultured in DMEM/F12 supplemented with 10% fetal bovine serum, 1× insulin-transferrin-selenium (Sigma, St. Louis, MO) antibiotic and antimycotic, and 10 μg/ml fibronectin (to aid adherence). Angiotensin II, endothelin-1, E2, BAPTA, EGTA, and pertussis toxin were all obtained from Sigma, and DPN was from Tocris Pharmaceuticals (Minneapolis, MN). KN93 (CaMKII inhibitor) was from EMD-Millipore (Billerica, MA), and apigenin, TBB, and CID 755673 were from Tocris. HDAC2, 4, and 5 and CaMKII total and phospho-antibodies and MHC isoform antibodies were from Abcam (Cambridge, MA). Phospho and/or total protein antibodies to CK2, PKCδ, and PKCβ were from Santa Cruz Biotechnology (Santa Cruz, CA), and PKD and phospho-PKD antibodies were from Cell Signaling (Danvers, MA). Control, ERα, ERβ, PKC, and PKD siRNAs were from Qiagen (Germantown, MD) or Santa Cruz Biotechnology. All primers were from Invitrogen (Carlsbad, CA). Dominant-negative HDAC2 (pcDNA3-HDAC2 S394A) and WT-HDAC2–encoding plasmids were kind gifts from Chinmay Trivedi, University of Massachusetts Medical School (Worcester, MA).

Measurement of protein synthesis, protein secretion, and cell area

After 24 h in media without serum, the cells were treated with 100 nM AngII in the presence or absence of 10 nM E2 or ERβ agonists. New protein synthesis, a marker of hypertrophy, was determined as described (Pedram et al., 2005). The cells were cultured as described with the addition of 1 μCi/ml [³H]leucine for 48 h. In some conditions, siRNA (control or to HDAC2) or a dominant-negative HDAC2 was expressed for 24 h before AngII exposure. After 48 h, the cells were rinsed in DMEM/F12, and lysed with TRIzol. The protein and RNA were extracted, the protein fraction counted in a scintillation counter, and the mean ± SEM of the counts plotted. Differences between treatments were compared for combined data from three experiments and the results analyzed by analysis of variance (ANOVA) plus Schefe's test (p < 0.05 was significant). Cell surface area was determined under the various conditions after culturing on cover slips and imaging by confocal microscopy. This was done at 40–50% cell confluency. Surface area was quantified by imaging to the complete boundary of 30 individual cells/condition using ImageJ software (National Institutes of Health, Bethesda, MD). From this, a mean ± SEM value was calculated for surface area in each condition from each experiment (Pedram et al., 2005). The planar aspect and deviations of individual cells in the z-axis were determined and adjusted.

Animal studies

All studies were approved by the Animal Care and Research and Development Committees at the Department of Veterans Affairs Medical Center, Long Beach, CA. The 10- to 12-wk-old female C57/BJ6 mice, intact or ovariectomized, were obtained from Harlan/Sprague Dawley, housed in 12-h on/off lighting, and fed rodent chow devoid of soy or most plant products. AngII (1.1 mg/kg per day) in saline or saline alone–filled osmotic minipumps (Alzet, Cupertino, CA; DURECT, Cupertino, CA) or AngII plus 100 μl of β-LGND2 (0.5 mg; Yepuru et al., 2010) was provided by 21-d infusion after subcutaneous insertion under inhaled chlorofluorane anesthesia. In some mice, an E2 pellet (0.5 mg, 21-d release pellets; Innovative Research of America, Sarasota, FL) or placebo pellet was inserted under the skin, but these mice did not receive β-LGND2. This pellet is well documented to produce physiological levels of E2 in the serum of mice (Pedram et al., 2010).

At 21 d, the hearts were removed and weighed, and the ratio of heart to total body weight was determined. RNA, DNA, and protein were extracted and the heart sectioned for further studies. For comparison, ovariectomized female wild-type and ERβ gene–deleted mice were obtained from Ken Korach, National Institute of Environmental Health Sciences (Research Triangle Park, NC) and Harlan Laboratories (Hayward, CA). These mice were subjected to the same conditions as described, and all mice were identically housed and fed the same chow.

Cellular calcium/Fura assay

Calcium activity reflected both extracellular calcium influx through membrane channels and intracellular calcium mobilization from stores and was determined as follows. Calcium was measured in neonatal cardiomyocyte cells by loading with Fluo-4 NW (Molecular Probes, Eugene, OR). The cells were cultured in 96-well microplates (poly-d-lysine [PDL] coated) to subconfluence (40,000–50,000 cells/well), and then cells were synchronized overnight in medium lacking fetal bovine serum and phenol red. The medium was removed to eliminate sources of baseline fluorescence, particularly esterase activity. Fluo-4 NW (100 μl) in loading solution was carefully added to each well, and the cells were incubated at 37°C for 60 min. The microplate was then transferred into a 37°C prewarmed Nowastar spectrofluorometer (BMG Labtech, Offenburg, Germany). The instrument was preset and loaded with test solutions at 2× concentration, with subsequent injection of 50 μl of control or AngII ± estrogenic compounds, followed immediately by 250 cycles of reading over 140 s. In one condition, pertussis toxin (100 mg/ml) was added at the same time as AngII + E2. Each condition was done in triplicate, and the study was repeated a second time. Fluorescence was measured at excitation of 494 nm and emission at 516 nm.

CK2 kinase assays

Synchronized cardiomyocytes were exposed to various treatments (e.g., ANG II ± E2) for 60 min. Cells for each condition (∼1 million) were then washed with DMEM/F12 medium and lysed in buffer (50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 50 mM NaF, 5 mM EDTA, 0.5% Triton X-100, 40 mM β-glycerophosphate, 200 mM sodium orthovanadate, 40 mM p-nitrophenol phosphate, 100 mM phenyl­methylsulfonyl fluoride (PMSF), and protease inhibitor cocktail [Sigma]). The lysates were centrifuged at 14,000 rpm for 10 min, and the supernatants were exposed to 50 μl of suspended protein A agarose beads in microcentrifuge tubes and rotated for 30 min at 4°C. The cell lysate/protein A bead complex was centrifuged at 6000 rpm for 30 s at 4°C. The supernatant was transferred to a new tube and stored on ice. For the CK2 activity assay, 10 μl of CK2 antibody (dilution 1:50; Santa Cruz Biotechnology) was conjugated to 50 μl of protein A beads for 2 h at room temperature, and the bead complex was washed. Then 1 ml of precleared whole-cell extract from each experimental condition was added to the CK2 antibody/protein A bead complex in lysis buffer (which also serves as the immune-precipitation buffer) and was rotated end over end overnight at 4°C. Beads were washed once with lysis buffer and twice with HEPES buffer (25 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid [HEPES], 10 mM Mg acetate). To the lysates/antibody/bead complexes, 40 μl of stock mixture, made up as 20 μl of 3× kinase buffer (25 mM HEPES, pH 7.5, 10 mM Mg acetate, 2 mM dithiothreitol, 40 mM ATP), and 10 μl of H₂O was added to 2 μg of CK2 substrate peptide (RRRADDSDDDDD; Signal Chem, Richmond, Canada) in 9 μl of water with 1 μl of [γ-³²P]ATP. Each tube was vortexed and incubated at 30°C for 30 min, and the reaction was stopped by adding 200 μg of casein and 100 μl of 4% trichloroacetic acid. Aliquots (50 μl) of supernatants were spotted on P81 phosphocellulose paper (Whatman) and washed with 0.5% phosphoric acid, and the radioactivity of the samples was counted in 5 ml of scintillation liquid. For in vivo determination of CK2 activity, the kinase protein was immunoprecipitated from the ventricular protein lysates of treated WT and ERβKO female mice, and the assay was carried out as described.

PKC, PKD, and CaMKII assays

PKCδ activity was determined at 30 min of incubation as phosphorylation of the protein isoform at the active site, Tyr-311. Cells were exposed to various conditions, and the cells were lysed, separated by SDS–PAGE, transferred to nitrocellulose, and immunoblotted with antibodies to Tyr-311 or antibodies to determine total PKC protein (Santa Cruz Biotechnology). PKD activity at 30 min was determined as Ser-744/748 phosphorylation by immunoblots from cells exposed to the various conditions, using phospho and total antibodies from Cell Signaling. CaMKII activity was determined at 60 min by immunoblot of phosphorylated Ser-632 of HDAC4 from the cardiomyocytes and by an in vitro tube assay, using [γ-³²P]ATP, immunoprecipitated CaMKII, and exogenous recombinant human HDAC4 substrate peptide (amino acids 612–1084; Abcam) to reflect HDAC4 Ser-467/632 phosphorylation by CaMKII. All studies were done three times for the combined data.

Quantitative real-time PCR

Total RNA was extracted using the Qiagen RNeasy Mini Kit following the manufacturer's protocol. All the samples were treated with DNAse-free (Ambion). RNA purity and concentrations were measured by UV spectrophotometry (A₂₆₀ and A₂₈₀). cDNA was synthesized using approximately 500 ng of RNA and Oligo dT primers with the Improm-II reverse transcription system (Promega). Quantitative real-time PCR (qRT-PCR) was used to examine the relative expression of HDAC2, 4, 5, βMHC, Inpp5f, GATA4, and ACTA1, and expression was normalized using the housekeeper glyceraldehyde-3-phosphate dehydrogenase (GAPDH). Primers were designed using Primer3 (http://frodo.wi.mit.edu/) and were blasted to check specificity (primers: HDAC2 forward [F], 5′-ttggcctttctgagctgatt-3′; HDAC2 reverse [R], 5′-agagggtctctgccactgaa-3′; HDAC4 F, 5′-gctctcccagctctccagca-3′; HDAC4 R, 5′-gttgtgagctgctgctgcaccgt-3′;HDAC5 F, 5′-tgagaggcaggcccttcagt-3′; HDAC5 R, 5′-cctccagtgccactcccaac-3′; GATA4 F, 5′-ggctcccagagattcttcct-3′; GATA4 R, 5′-ctctgctacggccagtaagg-3′; β-MHC F, 5′-accaagcagccacgccagta3′; β-MHC R, 5′-tgctttgcctttgcccttgt-3′; Inpp5f F, 5′-aacttgggaaaggcctgg-3′; Inpp5f R, 5′-catggagctgcggatctt-3′; α–smooth muscle actin (ACTA1), 5′-tcgcgaccttactgactacctg-3′, 5′-gcttctctttgatgtcgcgc-3′; GAPDH F, 5′-ccacagtcc atgccatca-3′; GAPDH R, 5′-ggatgaccttgcccacag-3′). Primers were designed to have an annealing temperature of 55°C and to amplify regions of approximately 150 base pairs. PCR amplicon sizes were confirmed by agarose gel electrophoresis before qRT-PCR analysis. For qRT-PCR analysis 500 ng of cDNA was used in a 50-μl reaction consisting of 25 μl of SYBR Green ER qPCR Supermix (Invitrogen), 1 μl of 10 μM forward/reverse primer stocks, and nuclease-free water. Thermocycling was carried out using the iCycler (Bio-Rad) with a melting curve temperature of 60°C. Relative mRNA levels were calculated using the C_t method (Livak and Schmittgen, 2001).

Subcellular fractionation

The subcellular fractionation was performed using the Subcellular Protein Fractionation Kit (78840; Pierce). The extraction buffer for cytoplasmic isolation (500 μl) was added to cell pellets. Cells were incubated at 4°C for 20 min while rotating. Then the homogenates were centrifuged (500 × g) at 4°C for 5 min. The supernatants (cytoplasmic extracts) were transferred to new Eppendorf tubes. The membrane extraction buffer (500 μl) was added to cell pellets, followed by vortexing and incubation at 4°C for 10 min. The homogenates were centrifuged (5600 × g) at 4°C (5 min), and the supernatants (membrane extracts) were transferred to new Eppendorf tubes. The nuclear extraction buffer (250 μl) was added to cell pellets, followed by vortexing and incubation at 4°C for 30 min. The homogenates were centrifuged (7300 × g) at 4°C for 5 min, supernatants (nuclear extracts) were transferred to new Eppendorf tubes, and protein concentrations were estimated by the Bradford assay. Western blotting was performed by loading 20 μg of each cytoplasmic and nuclear extract. Lack of contamination of the cell fractions was accomplished as previously described (Pedram et al., 2006).

Immunofluorescence microscopy

Cardiomyocyte cells (3 × 10⁵) were placed on PDL-coated glass-bottom dishes (MatTek, Ashland, MA). Twenty-four hours later, at a confluency of 70–80%, cells were treated with various agents, washed three times with phosphate-buffered saline (PBS), and fixed with 4% fresh paraformaldehyde in PBS for 10 min at room temperature. After further washing with PBS, the cells were permeabilized with 0.2% Triton Χ-100 in PBS for 5 min and washed with PBS. Before antibody incubation, cells were blocked with serum from the same species as the secondary antibody or 2% bovine serum albumin (BSA) in PBS at room temperature for 30 min. After washing with PBS, cells were incubated overnight at 4°C with primary antibody (dilution 1:50 in 0.5% BSA-PBS). The cells were then incubated for 1 h with fluorescein isothiocyanate–conjugated secondary antibody (dilution, 1:500; Vector Laboratories) at room temperature. Cells were washed three times with PBS, and samples were covered with coverslips and mounting medium and analyzed by immunofluorescence microscopy (Nikon).

Chromatin immunoprecipitation assay

ChIP assays were performed using the Active Motif ChIP-IT kit. For each experimental condition, four confluent 10-cm plates of neonatal cardiomyocytes cells (10⁷cells/dish) were used. After treatments for 24 h, cells were washed with ice-cold PBS and cross-linked for 5 min with 1% formaldehyde, and the reaction was stopped by adding glycine (125 mM). The cells were scraped and washed with cold PBS containing proteinase inhibitor cocktail (Sigma) and 1 mM PMSF. Enzymatic fragmentation of chromatin was performed using the Enzymatic Shearing kit (Active Motif) according to the manufacturer's instructions. For the ChIP assays, 50 μg of digested chromatin was precleared for 1 h with protein A–Sepharose beads. After centrifugation, supernatants were incubated with 10 μg of acetylated histone H3 antibody or rabbit immunoglobulin G (IgG; Cell Signaling) or no antibody (mock condition) and incubated overnight at 4°C under rotation. The immune-precipitated complexes were bound to protein A–Sepharose beads during 3 h at 4°C and washed successively in a lysis buffer (10 mM Tris/HCl, 1 mM EDTA, 1% Triton X-100, 0.1% SDS, 140 mM NaCl, 0.1% sodium deoxycholate, and 1 mM PMSF, pH 7.4), LiCl buffer (1 mM Tris/HCl, 250 mM LiCl, 0.5% Nonidet P40, 0.5% sodium deoxycholate, and 1 mM EDTA, pH 7.4), and Tris–EDTA buffer (1 mM EDTA and 10 mM Tris/HCl, pH 7.4). Chromatin complexes were eluted by three sequential incubations at room temperature (twice for 15 min and once for 45 min) in 90 μl of elution buffer (100 mM NaHCO₃ and 1% [wt/vol] SDS). Proteins were then eliminated by using 200 μg of proteinase K (Promega) in the presence of 10% (wt/vol) SDS by overnight incubation at 37°C. Chromatin DNA was extracted with phenol/chloroform, followed by ethanol precipitation. Samples were dissolved in 80 μl of water. The PCR primers used for the analysis of the rat βMHC promoter (directed against residues between −317 and −255) were 5′-atagcaacagcgaggctctttctg-3′, rat β-MHC, forward; and 5′-ggagctcacctaccagacaga-3′, rat β-MHC, reverse; they were provided by Qiagen. PCR amplification products were analyzed on ethidium bromide–stained 2.5% (wt/vol ) agarose gels.