* College of Pharmacy, School of Medicine, University of New Mexico, Albuquerque, New Mexico; Department of Cell Biology and Physiology, School of Medicine, University of New Mexico, Albuquerque, New Mexico
1 To whom correspondence should be addressed at College of Pharmacy, MSC09 5360, 1 University of New Mexico, Albuquerque, NM 87131. Fax: (505) 272-0704. E-mail: mkwalker{at}unm.edu.
Received May 19, 2005; accepted August 9, 2005
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ABSTRACT |
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Key Words: Aryl hydrocarbon receptor (AhR); reactive oxygen species; cardiac hypertrophy; endothelin-1; NAD(P)H oxidase.
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INTRODUCTION |
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Endothelin-1, a potent vasoactive and mitogenic peptide produced mainly by endothelial cells, is associated with initiation and progression of cardiac hypertrophy (Shubeita et al., 1990), mainly through ETA receptors on the myocardium. The role of ET-1 in cardiac hypertrophy has been confirmed through receptor antagonist studies, such that treatment results in attenuated progression of cardiac remodeling (Ehmke et al., 1999
). Reactive oxygen species (ROS) have been shown to be critical mediators of ET-1-induced growth-promoting signaling events involved in the hypertrophic pathways in vascular smooth muscle cells (Daou and Srivastava, 2004
) and cardiomyocytes (Hirotani et al., 2002
). The role of ROS in ET-1induced cardiac hypertrophy has been further confirmed through studies showing that ET-1mediated generation of ROS in cardiac hypertrophy can be inhibited by pretreatment with an antioxidant (Xu et al., 2004
). Although it is not yet understood how an increase in ROS may contribute to ET-1dependent cardiac hypertrophy, recent studies suggest that ROS modulates vascular tone in both arteries and veins, resulting in increased total peripheral resistance and elevated blood pressure (Thakali et al., 2005
). Additionally, ROS may induce cardiac hypertrophy by activating signal transduction pathways, such as mitogen-activating protein kinase (MAPK), which mediate the hypertrophic response of cardiomyocytes (Cheng et al., 2005
).
In mammalian tissues, ROS are formed under both physiological and pathological conditions. Myocardial ROS, specifically the superoxide anion (), have been implicated in a large number of diseases, with evidence from both experimental and clinical studies suggesting a causal role of oxidative stress in the pathogenesis of congestive heart failure and cardiac hypertrophy (Dhalla et al., 2000
; Sugden and Clerk, 1998
). The role of ROS in cardiac hypertrophy is further supported by the finding that antioxidant treatment inhibits cardiac myocyte hypertrophy in both neonatal (Nakamura et al., 1998
) and adult rat myocytes (Tanka et al., 2001
).
Although the sources of ROS in the hypertrophying heart have not yet been fully elucidated, phagocyte-like NAD(P)H oxidases have emerged as a major source of ROS generation in the cardiovascular system (Li et al., 2002). The NAD(P)H oxidase complex consists of a core heterodimer comprised of a phagocytic oxidase (p22phox) subunit and a glycoprotein (gp91phox) subunit (or homologs termed Nox1 or Nox4), and four regulatory subunits: p47phox, p67phox, p40phox, and rac1. NAD(P)H oxidases have been identified in cardiomyocytes (Xiao et al., 2002
), and subunits of the NAD(P)H oxidase complex have been found to be upregulated in the myocardium in states of cardiac pathology (Li et al., 2002
).
It has not been determined whether an increase in ROS is associated with the ET-1dependent cardiac hypertrophy in AhR null mice. Thus, we tested the novel hypothesis that increased ET-1 signaling via the ETA receptor induces ROS, specifically via NAD(P)H oxidase induction in the hearts of AhR null mice.
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MATERIALS AND METHODS |
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Superoxide analysis in heart.
To assess in vivo levels and localization of cardiac AhR null and wild-type mice were treated with dihydroethidium (DHE, Molecular Probes, Eugene, OR). Dihydroethidium enters the cell and is oxidized primarily by
to yield fluorescent products, such as ethidium (Buxser et al., 1999
), which intercalates into DNA. Because ethidium fluoresces at a different wavelength (Ex = 495 nm; Em = 595 nm) than DHE (Ex = 365 nm; Em = 415 nm), ethidium fluorescence can be used to visualize localized
production (Chan et al., 1998
). However, the specificity of ethidium as a marker of
has recently been questioned. A new molecular species has been identified, which is believed to be the specific product of DHE oxidation by
(Zhao et al., 2003
). This new product has a different molecular weight than ethidium, and distinctive fluorescence characteristics (Ex = 480 nm; Em = 567 nm). Thus, in these experiments we used a custom-built fluorescence filter (Ex = 460500 nm; Em = 540585 nm) to characterize the production of the
specific derived DHE product, in addition to ethidium fluorescence. Dihydroethidium was dissolved in 0.1 M phosphate-buffered saline (PBS) containing 20% dimethyl sulfoxide to a final concentration of 0.5 mM. Mice were injected ip with 1 mg/ml. Dihydroethidium -treated animals were sacrificed 6 h after treatment, and after exsanguination, the hearts were dissected, fixed in 10% neutral-buffered formalin (vol/vol) for 24 h, and sectioned at 30 µm with a vibratome. Tissue sections were viewed using excitation at 510550 nm and emission at >580 nm for ethidium detection (Olympus BH2-RFCA), and digital images were acquired (Olympus MLH020550) and analyzed with Image ProPlus. The percentage of
production was quantified as the ratio of fluorescent area to total ventricular area.
BQ-123 dosing groups.
Two-month-old AhR wild-type and null male mice were randomly assigned to either BQ-123 (American Peptide Company, Vista, CA) or vehicle (saline) treatment groups, and treated for 28 or 58 days. The BQ-123 group received 100 nmol/kg/day (dose based on personal communication with Dr. Nancy Kanagy; a dose shown to effectively reduce blood pressure in AhR null mice, Lund et al., 2005) for 28 or 58 days via osmotic mini-pumps at a flow of 0.25 µl/h (model no. 2004, Alzet, Cupertino, CA). These treatment durations were chosen in an effort to prevent the pathology (both gross tissue and molecular) associated with the onset and progression of cardiac hypertrophy observed in adult AhR null mice. The 58-day treatment period was successful in preventing the onset of cardiac hypertrophy in AhR null mice (Lund et al., 2005
); thus the 28-day treatment was carried out to determine if a shorter treatment period would also be adequate. Animals were briefly anesthetized with avertin (2.5% ip at 0.015 ml/g), and mini-pumps were implanted subcutaneously. Animals were monitored daily for health status throughout the study period. No obvious side effects were noted in the BQ-123-treated animals throughout the study (no differences noted in weight, eating, drinking, or activity compared to untreated animals). One group of animals was sacrificed on day 28; plasma was collected for detection of 8-isoprostane, and hearts were collected for assessment of NAD(P)H oxidase activity. In the second group, the mini-pumps were replaced on day 29, and treatment continued until day 58. On day 58, mice were euthanized, body and heart weights were measured, and heart tissue was collected for thiobarbituric acid reactive substances (TBARS) and NAD(P)H oxidase subunit quantification.
Plasma and tissue collection.
AhR wild-type and null mice were anesthetized with ketamine/xylazine and euthanized by exsanguination. Blood was collected in a heparinized syringe (BD Vacutainer Systems, Franklin Lakes, NJ) through cardiac puncture, and immediately centrifuged (950 x g, 10 min, 4°C) to separate plasma. Plasma was stored at 70°C for 8-isoprostane analysis, until assayed. Additionally, the heart was dissected, weighed, and frozen in liquid nitrogen. Cardiac tissue was stored at 70°C until assayed.
8-Isoprostane analysis.
Plasma 8-isoprostane was measured in AhR null and wild-type animals using an 8-isoprostane competitive enzyme immunoassay (EIA; Caymen Chemicals, Ann Arbor, MI), following the protocol provided for determination of total (free and esterfied) 8-isoprostane. All samples were purified using an 8-isoprostane affinity column (Caymen Chemicals model no. 416358) prior to analysis. Samples were run in triplicate and results were averaged. Results were calculated and quantified by a blinded participant.
Thiobarbituric acid reactive substances (TBARS) analysis.
Cardiac tissue was resuspended by diluting 1:10 weight/volume in normal saline. Tissue was homogenized in Potter-Elvejhem glass homogenizer, and sonicated for 15 s at 40 V. A TBARS assay kit (OXItek, ZeptoMetrix Corp, Buffalo, NY) was used to measure TBARS levels in whole, uncentrifuged homogenates. Duplicate samples were read on a spectrophotometer (Beckman Instruments DU Series 600), and using a malondialdehyde (MDA) standard curve, and results were expressed as MDA equivalents.
Quantification of ventricular NAD(P)H oxidase-mediated O2 levels.
Cardiac production was assessed in ventricular samples by the lucigenin chemiluminescence method. Chemiluminescence is produced by the reaction of lucigenen with
and only weakly with H2O2, but not with mucloperoxidase. To prevent autoxidation of lucigenin, a with a low concentration (5 µmol/l) of lucigenin was used, as previously described (Wu et al., 2004
), with the following modifications. Ventricular tissue (hearts cut transversely to include right and left ventricle) were cut in 10-mg blocks, rinsed in ice-cold PBS, and placed in cold saline on ice for 10 min. Lucigenin and PBS were added to recording tubes and incubated in the dark for 15 min. Background counts were then obtained by measuring chemiluminescence in a luminometer (Turner Designs TD-20/20 Luminometer) for 5 min (with a 1.5-min dark adjustment). Ventricular blocks were then added and measured for 5 min. To evaluate NAD(P)H oxidase activity, 100 µM NAD(P)H was then added to ventricle samples, and luminescence was measured for an additional 5 min. In some experiments, ventricle blocks were pre-incubated with 105 M apocynin (4-hydroxy-3-methoxy-acetophenone), an inhibitor of superoxide production by NAD(P)H oxidases, or Tiron (4,5-dihydroxyl-1,3-benzene-disulphonic acid, 10 mmol/l), a cell-permeable nonenzymatic scavenger of
for 30 min before reading. Background counts (with lucigenin) were subtracted from each value obtained from ventricular blocks. Lucigenin chemiluminescent counts were adjusted on the basis of dry weight of the ventricle blocks. Activity was expressed as relative light units (RLU)/per mg dry tissue weight/5 min.
Gp91phox, p47phox, p67phox mRNA analysis.
Total RNA was isolated from the left ventricle (LV) plus attached septum with Trizol (Sigma Chemical Co., St. Louis, MO). cDNA was synthesized from total RNA in a 60-µl final reaction volume containing 250 ng of sample RNA, 12.5 nM of 18S reverse transcriptase (RT) primer (Table 1), 0.005 µg oligo dT, 0.0004u RNAsin, 0.006u M-MLV RT enzyme, 25 mM dNTP, 12 µl 5x RT buffer, and sterile water to 60 µl volume. The mixture was heated at 42°C for 1 h and then cooled to 4°C. Real-time PCR was performed with gene-specific primers in an iCycler (Biorad, Hercules, CA). The following murine-specific primer sets were used at a concentration of 500 nM for the PCR reaction: gp91phox forward: CACCCATTCACACTGACCTCTG, gp91phox reverse: CTTATCACAGCCACAAGCATTGAA; p47phox forward: CTGCTGTTGAAGAGGACGAGATG, p47phox reverse: AGCCGGTGATATCCCCTTTCC; p67phox forward: CTCGCCAGAACACACTAAACTGA, p67phox reverse: TCCTTCATGCTTTCTTCGGACAG; m18S Forward: GCTTGCGTTGATTAAGTCCCTG, and m18S Reverse: AGTTCGACCGTCTTCTCAGC. Control reactions without reverse transcriptase or without RNA were run to verify the absence of contaminated DNA and primer-dimerization, respectively. Polymerase chain reaction amplification was carried out in a 25-µl reaction volume containing 0.25 ng of cDNA, 500 nM each forward and reverse primers, 12.5 µl iQ SYBR green Supermix (Biorad), and 9.5 µl sterile water. The PCR reactions were initiated with denaturation at 95°C for 60 s; followed by amplification with 40 cycles at 30 s, 95°C; annealing for 2 min at 54°C, and an extension at 72°C for 5 min. To confirm the presence of a single amplification product, PCR products were subjected to a melt curve analysis and were run on an agarose gel. Samples were run in triplicate and results were averaged. CT (change in threshold cycle) was calculated by subtracting the CT of the 18S control gene from the CT value of the gene of interest and mean normalized gene expression was calculated as previously described (Lund et al., 2003
). Results are expressed as normalized gene expression as percentage of controls.
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RESULTS |
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BQ-123-Treatment Reduces Plasma 8-Isoprostane Levels in AhR Null Mice
Isoprostanes are prostaglandin-like compounds that are produced by the random oxidation of tissue phospholipids by oxygen radicals. At least one of the isoprostanes, 8-isoprostane (8-epi PGF2), has been shown to have biological activity and is used as a marker of antioxidant deficiency and oxidative stress (Morrow et al., 1995). AhR null mice exhibit significantly increased levels of plasma 8-isoprostane, compared to AhR wild-type mice (Fig. 2). BQ-123-treatment significantly reduced plasma 8-isoprostane levels in AhR null mice, compared to untreated AhR null mice, to values that were not significantly different from controls at p < 0.05 (Fig. 2). There was no difference in plasma 8-isoprostane between BQ-123treated and untreated wild-type mice.
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DISCUSSION |
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The development of cardiac hypertrophy in AhR null mice is well documented (Fernandez-Salguero et al., 1997; Lund et al., 2003
; Thackaberry et al., 2002
; Vasquez et al., 2003
), and a previous study has shown that ET-1 is the primary mediator of the cardiac hypertrophy (Lund et al., 2005
). Endothelin-1 is associated with generation of ROS in cardiomyocytes (Hirotani et al., 2002
), which are believed to mediate ET-1induced cardiac hypertrophy. This premise has been confirmed by study findings which show ET-1mediated generation of ROS in cardiac hypertrophy can be ameliorated with antioxidant therapy (Xu et al., 2004
).
Reactive oxygen species, including hydrogen peroxide (H2O2), and hydroxyl radicals (OH), have been implicated in the pathologic etiology and progression of cardiac hypertrophy (Dhalla et al., 2000
). Thus, to determine whether cardiac ROS are associated with the hypertrophic response in AhR null mice, DHE was used to visualize the presence and localization of
in vivo.
-derived specific fluorescence show elevated
levels in the ventricles of AhR null mice, compared to AhR wild-type mice. Interestingly, the epicardium of both AhR null and wild type exhibited high levels of fluorescence that may result from oxygen-rich circulation through coronary arteries present in the epicardium.
We next investigated whether the cardiac hypertrophy and elevated ROS observed in AhR null mice were mediated by ET-1 via ETA receptors. Endothelin-1 is believed to exert its growth-promoting effects on cardiac myocytes primarily through activation of ETA receptors, a conclusion supported by study findings that show that ETA receptor antagonism attenuates cardiac hypertrophy in experimental models (Ehmke et al., 1999; Ito et al., 1994
). Thus, 2-month-old AhR null mice were treated for 28 days with the ETA receptor antagonist BQ-123. In agreement with previously cited studies (Ehmke et al., 1999
; Ito et al., 1994
), we found that ETA blockade significantly decreases heart weight and the heart weight-to-body weight ratio in AhR null mice.
To characterize the ability of ETA blockade to reduce ROS associated with cardiac hypertrophy in AhR null mice, we used two independent measures of ROS generation: cardiac TBARS, a nonspecific assessment of ROS production, and plasma 8-isoprostane, a systemic product that forms as the result of oxidation of phospholipids by ROS (Roberts and Morrow, 2000). Cardiac TBARS are significantly elevated in AhR null mice, and the increase over that measured in AhR wild-type mice is similar to inductions previously reported in both animal and human models of cardiac hypertrophy (Luo et al., 2002
; Motoyama et al., 2001
). Additionally, plasma 8-isoprostane levels are also significantly increased in AhR null mice, compared to AhR wild-type mice; and similar inductions have also been reported in other models of cardiovascular pathology, such as hypertension (Patterson et al., 2005
). ETA blockade significantly reduces both cardiac TBARS and plasma 8-isoprostane in AhR null mice; however, levels of both of these indicators of ROS remain elevated compared to AhR wild-type mice, suggesting that induction of ROS in AhR null mice is only partially mediated by ET-1 activation of ETA receptors. One possible explanation for the inability of ETA blockade to completely reduce TBARS and 8-isoprostane levels is that plasma Ang II remains significantly elevated in AhR null mice, but not in wild-type mice, after chronic ETA antagonist therapy (Lund et al., 2005
). Angiotensin II has been shown to increase cardiac ROS production (Nakagami et al., 2003
) and thus, elevated levels of Ang II in AhR null mice may represent an additional contributor to the production of cardiac and plasma ROS. Future studies involving both ETA and Ang II receptor antagonism will help to define the roles of these two vasoactive peptides in ROS production in AhR null mice.
In an effort to determine the source of elevated ROS observed in the myocardium in AhR null mice, we assessed NAD(P)H oxidase activity. The NAD(P)H oxidases of the cardiovascular system are membrane-associated enzymes that catalyze the 1-electron reduction of oxygen using NADH or NAD(P)H as the electron donor. In vitro studies suggest that ET-1 induces NAD(P)H oxidases in several cell types (Duerrschmidt et al., 2000; Fei et al., 2000
), including cardiac myocytes (Tanka et al., 2001
). Our results show that NAD(P)H oxidase-generated ROS, as measured by lucigenin chemiluminescence, is increased in hearts of AhR null mice. Furthermore, the origin of ROS appears to be mediated by NAD(P)H oxidase, because pretreatment with apocynin, an inhibitor of superoxide production by NAD(P)H oxidases, suppresses the increased lucigenin activity to control levels. Finally, this induction of NAD(P)H oxidase appears to be mediated through ET-1ETA receptor activation because BQ-123 significantly reduces NAD(P)H oxidase activity. These data indicate that cardiac hypertrophy in AhR null mice is associated with increased NAD(P)H oxidase induction, resulting in elevation of cardiac ROS levels.
These findings are further confirmed through analysis of expression of cardiac NAD(P)H oxidase components, gp91phox, p67phox, and p47phox. The gp91phox, along with the p22phox, are integral membrane proteins; whereas the p67phox and p47phox subunits are located in the cytosol. Upon activation, p47phox is phosphorylated and translocated with p67phox to the gp91phoxp22phox core oxidase, resulting in production. Expression of each of these subunits has been confirmed in cardiac myocytes (Xiao et al., 2002
), and NAD(P)H oxidase can be regulated over the long term by upregulation of transcription of the oxidase subunits (Lessègue and Clempus, 2003
; Touyz et al., 2002
). Real-time PCR analysis of mRNA expression of these NAD(P)H oxidase subunits shows that all are significantly elevated in AhR null mice, when compared to the wild type. BQ-123 treatment results in a decrease in expression of gp91phox, p67phox, and p47phox in AhR null mice. Such findings suggest that NADPH oxidase expression is regulated through an ET-1ETAdependent mechanism in hearts of AhR null mice. Whereas ET-1 has previously been shown to increase NAD(P)H oxidase activity in endothelial cells (Duerrschmidt et al., 2000
), and to slightly increase activity in vascular smooth muscle cells (Touyz et al., 2004
), to our knowledge these findings are the first to report that ETA blockade results in decreased expression of NAD(P)H oxidase subunits (gp91phox, p67phox, and p47phox), suggesting that ET-1 mediates NAD(P)H oxidase activity in cardiac tissue. These findings also suggest that any Ang-II derived ROS present in hearts of AhR null mice are not NAD(P)H oxidase derived, because NAD(P)H oxidase activity is normalized through BQ-123 treatment.
Previous studies have shown that both Ang II and ET-1 contribute to cardiac hypertrophy observed in AhR null mice (Lund et al., 2003, 2005
) and both of these cardiac mitogenic peptides are believed to mediate their growth-promoting effects on the heart by increased production of ROS (Tanka et al., 2001
). It has been proposed that ROS may mediate the cardiac hypertrophic response by acting as a regulator of gene expression (Kunsch and Medford, 1999
), either through direct activation of G proteins (Chiloeches et al., 1999
; Nishida et al., 2002
) or by altering activity of other growth-promoting signaling pathways, such as MAPKs (Kyaw et al., 2002
). Thus, it is tempting to speculate that induction of ROS by ET-1 and/or Ang II mediates the development and progression of cardiac hypertrophy in AhR null mice. Although the present study demonstrates that ET-1 increases cardiac ROS via ETA receptors in AhR null mice, more experiments are needed to delineate the causative role of ROS in the development and progression of the cardiac hypertrophy. The results reported here suggest a role for
in the progression of cardiac hypertrophy in AhR null mice; however, we have not yet determined whether other
- derived ROS, such as hydrogen peroxide (H2O2), may also contribute to the pathology of the myocardium.
In conclusion, we have shown that AhR null mice exhibit cardiac hypertrophy, which is associated with increased production of cardiac ROS. Furthermore, chronic treatment with an ETA receptor antagonist, BQ-123, results in a significant reduction of cardiac hypertrophy and NADPH oxidasederived production in hearts of AhR null mice. Such results suggest that ET-1 mediates the increase in cardiac NAD(P)H oxidaseinduced
production in AhR null mice through an ETA receptor-signaling pathway. The mechanisms by which ROS may contribute to the progression of cardiac hypertrophy in AhR null mice remains to be determined, and it can likely be clarified through studies involving antioxidant therapy. Given that recent studies have identified endogenous ligands of the AhR (Song et al., 2002
; reviewed in Denison and Nagy, 2003
), future studies that further elucidate the role of AhR in cardiovascular physiology may provide signaling pathways that serve as mediators of the induction or progression of cardiac hypertrophy.
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ACKNOWLEDGMENTS |
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