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Alterations in cytochrome P450-derived arachidonic acid metabolism during pressure overload-induced cardiac hypertrophy Ahmed A. El-Sherbeni, Ayman O.S. El-Kadi * Faculty of Pharmacy and Pharmaceutical Sciences, University of Alberta, Edmonton, Alberta, Canada T6G 2E1

A R T I C L E I N F O

A B S T R A C T

Article history: Received 6 October 2013 Accepted 22 November 2013 Available online xxx

Cardiac hypertrophy is a major risk factor for many serious heart diseases. Recent data demonstrated the role of cytochrome P450 (CYP)-derived arachidonic acid (AA) metabolites in cardiovascular pathophysiology. In the current study our aim was to determine the aberrations in CYP-mediated AA metabolism in the heart during cardiac hypertrophy. Pressure overload cardiac hypertrophy was induced in Sprague Dawley rats using the descending aortic constriction procedure. Five weeks postsurgery, the cardiac levels of AA metabolites were determined in hypertrophied and normal hearts. In addition, the formation rate of AA metabolites, as well as, CYP expression in cardiac microsomal fraction was also determined. AA metabolites were measured by liquid chromatography–electrospray ionization-mass spectroscopy, whereas, the expression of CYPs was determined by Western blot analysis. Non-parametric analysis was performed to examine the association between metabolites formation and CYP expressions. Our results showed that 5,6-, 8,9-, 11,12-, and 14,15-epoxyeicosatrienoic acids (EETs), and 5-, 12-, 15-, and 20-hydroxyeicosatetraenoic acids (HETEs) levels were increased, whereas, 19-HETE formation was decreased in hypertrophied hearts. The increase in EETs was linked to CYP2B2. On the other hand, CYP1B1 and CYP2J3 were involved in mid-chain HETE metabolism, whereas, CYP4A2/3 inhibition was involved in the decrease in 19-HETE formation in hypertrophied hearts. In conclusion, CYP1B1 played cardiotoxic role, whereas, CYP2B2, CYP2J3 and CYP4A2/3 played cardioprotective roles during pressure overload-induced cardiac hypertrophy. These CYP can be valid targets for the development of drugs to treat and prevent cardiac hypertrophy and heart failure. ß 2013 Elsevier Inc. All rights reserved.

Keywords: Cardiac hypertrophy Heart failure Arachidonic acid metabolism Cytochrome P450 Eicosanoids

1. Introduction Cardiac hypertrophy is a major risk factor for several heart diseases, most importantly heart failure and sudden death [1]. Cardiac hypertrophy begins as adaptive response; it initially meets the insult-induced hemodynamic overload [2]. However, prolonged hypertrophy is catastrophic, resulting in progressive deterioration in heart functions that eventually leads to heart failure [3]. Better understanding of cardiac hypertrophy can lead to better diagnosis, and treatment of heart failure. Heart failure is a serious health problem; about 6.6 million people in US are affected by heart failure, and the number is expected to rise to about 9.6 million by 2030 [4]. There are about 650,000 new cases every year, and mortality rate associated with heart failure is 50% within 5 years [4,5].

* Corresponding author. E-mail addresses: [email protected], [email protected] (Ayman O.S. El-Kadi).

Eicosanoids are the biological active metabolites of AA by cyclooxygenases, lipoxygenases, or cytochrome P450 enzymes (CYP) [6]. These metabolites have been recognized to occupy central role in pathophysiology of cardiovascular system, especially CYP-derived AA metabolites [7]. AA is oxidized, by CYP, to epoxy- and hydroxy-metabolites in the presence of oxygen and NADPH [8]. Several CYP enzymes are known to catalyze the metabolism of CYP-derived epoxy- and hydroxy-metabolites. The formation of epoxy-metabolites is mediated mostly by CYP2 family, notably, CYP2B, CYP2C and CYP2J enzymes [8]. They catalyze the oxidation of AA to 5,6-, 8,9-, 11,12-, and 14,15epoxyeicosatrienoic acids (EETs). Soluble epoxide hydrolase (sEH), then, catalyzes the hydration of EETs to the less-biologically-active dihydroxyeicosatrienoic acids (DHETs) [9]. The hydroxyl-metabolites of AA are divided into mid-chain and v/v1 hydroxyeicosatetraenoic acids (HETEs) [8]. CYP1B1, and to a lower extent CYP1As, have lipoxygenase-like activity of oxidizing AA to midchain HETEs, typified by 5-, 12-, and 15-HETE, through hydroperoxy-intermediates [10,11]. On the other hand, a heterogeneous group of CYP catalyzes the formation of terminal HETEs, most importantly, CYP1As for 16-, 17, 18- and 19-HETE [12], CYP2E1 and

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CYP4A2 for 19-HETE [8,13], and CYP4As and CYP4Fs for 20-HETE [14]. The effects of CYP-derived AA metabolites on cardiovascular system are multifaceted. For example, EETs exhibit potent vasodilatory, anti-platelet, anti-inflammatory, fibrinolytic, vascular smooth muscle anti-migratory, and angiogenic properties [9,15,16]. On the other hand, 20-HETE is reported to be a potent vasoconstrictor and pro-inflammatory and it stimulates vascular smooth muscle cell migration [17–19]. Other terminal HETEs, 16-, 17-, 18-, and 19-HETE, act as endogenous antagonists for 20-HETE [20,21]. With respect to mid-chain HETEs, they have been reported to have vasoconstrictive and pro-inflammatory action [22–25]. Furthermore, 12-HETE has been reported to induce cellular hypertrophy, fibrosis and matrix protein synthesis, whereas, 15HETE has been proposed to be implicated in heart failure by induction of cardiac fibrosis [26,27]. In the light of the aforementioned biological activity of CYPderived AA metabolites, depicting a comprehensive metabolic profile of these metabolites in normal and hypertrophied hearts is invaluable. Measuring free, unesterified, metabolites levels in heart tissue has been proved to be feasible with some AA metabolites, such as mid-chain HETEs. For the other undetectable metabolites, measuring their formation and degradation rates in vitro by the heart can give a clear insight about the endogenous levels of these metabolites. The aims of the current study were: (1) to determine the alterations in CYP-derived AA metabolites levels in hearts tissue associated with pressure overload-induced cardiac hypertrophy in Sprague Dawley rats, (2) to determine the alterations in the formation and degradation rates of CYP-derived AA metabolites in heart tissue, and (3) to determine CYP enzymes involved in the observed aberrant changes in metabolites levels. 2. Materials and methods 2.1. Materials AA, butylated hydroxytoluene, camphorsulfonic acid, ethylenediaminetetraacetic acid, nicotinamide adenine dinucleotide phosphate (NADPH), sucrose, and triethylamine were purchased from Sigma–Aldrich Chemical Co. (St. Louis, MO). AA metabolite standards: 5,6-, 8,9-, 11,12-, and 14,15-EET, 5,6-, 8,9-, 11,12-, and 14,15-DHET, 5-, 12-, 15-, 16-, 17-, 18-, 19-, and 20-HETE, and 12hydroperoxyeicosatetraenoic acid (12-HPETE) as well as the deuterated metabolites (internal standards) were purchased from Cayman Chemical (Ann Arbor, MI). 8-Hydroxy-11,12-epoxyeicosatrienoic acid was purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). High performance liquid chromatography (HPLC) grade acetonitrile, methanol, water, and ethyl acetate were purchased from EM Scientific (Gibbstawn, NJ). Potassium chloride, calcium chloride dihydrate, potassium dihydrogen orthophosphate, dipotassium hydrogen orthophosphate, and magnesium chloride hexahydrate were obtained from BDH (Toronto, ON, Canada). Acrylamide, N0 N0 -bis-methylene-acrylamide, ammonium persulphate, b-mercaptoethanol, glycine, nitrocellulose membrane (0.45 mm) and TEMED were purchased from Bio-Rad Laboratories (Hercules, CA, USA). Chemiluminescence Western blotting detection reagents were purchased from GE Healthcare Life Sciences, Piscataway, NJ, USA. Mouse anti-rat CYP1A1, rabbit anti-rat CYP1B1, rabbit anti-rat CYP2C11 and rabbit anti-human CYP4F2 primary antibodies were purchased from Abcam (Cambridge, UK). Rabbit anti-human CYP2Js primary antibody was a generous gift from Dr. Darryl Zeldin (National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, NC). Mouse anti-rat CYP2B1/2, goat antirat CYP2E1, mouse anti-rat CYP4A1/2/3, goat anti-rat GAPDH and secondary antibodies were purchased from Santa Cruz Biotech-

nology, Inc. (Santa Cruz, CA). CYP1B1- and CYP2J2-containing cell microsomes (Supersomes1) were obtained from Gentest (Woburn, MA), whereas cytochrome b5 was obtained from Oxford Biomedical Research (Oxford, MI). Oasis1HLB (30 mg, 30 mm) cartridges were purchased from Waters Corporation (Maliford, MA, USA). Other chemicals were purchased from Fisher Scientific Co. (Toronto, ON, Canada). 2.2. Pressure overload induction by descending aortic constriction (DAC) All animals were maintained and used in accordance with the animal protocol approved by the University of Alberta Health Sciences Animal Policy and Welfare Committee, Edmonton, Alberta, Canada. Male Sprague Dawley rats (Charles River Canada, St. Constant, QC, Canada) weighing 150–200 g were subjected to the descending aortic constriction for induction of cardiac hypertrophy as described previously [28]. Briefly, animals were anesthetized for surgery with isoflurane. The suprarenal abdominal aorta was exposed by a midline laparotomy. A blunt 21-gauge needle was used as a guide for tying off aorta by silk suture. Shamoperated rats were subjected to the same procedure except for the aortic constriction. 2.3. Echocardiography Five weeks post-surgery, sham and DAC animals were weighed and mildly anesthetized using 1.5% isoflurane. Subsequently, transthoracic echocardiography was performed using a Vevo 770 Imaging System (VisualSonics, Toronto, ON). Heart dimensions, namely the left ventricle posterior wall thickness, interventricular septal wall thickness, and left ventricle volume during diastole, were determined. Systolic and diastolic functions, namely the percentage of fractional shortening, percentage of ejection fraction, and Tei index, were measured. Tei index was calculated as the sum of the isovolumic relaxation and contraction time divided by the ejection time. 2.4. Tissue collection and microsomal preparation Animals were sacrificed under isoflurane anesthesia. Hearts were immediately frozen in liquid nitrogen and stored at 80 8C. The tibial length was measure by micro-CT (SkyScan NV, Kontich, Belgium). Microsomal fractions were prepared by differential centrifugation of homogenized tissues as described previously [29]. Briefly, organs were washed in ice-cold potassium chloride (1.15%, w/v). Subsequently, they cut into pieces, and homogenized in ice-cold 0.25 M sucrose solution sucrose (17%, w/v). After homogenization process, the tissues were separated by differential ultracentrifugation. The final pellet was re-suspended in cold sucrose and stored at 80 8C. The microsomal protein concentration was determined by the Lowry method using bovine serum albumin as a standard [30]. 2.5. RNA extraction and cDNA synthesis Total RNA from the frozen tissues was isolated using TRIzol reagent (Invitrogen) according to the manufacturer’s instructions, and quantified by measuring the absorbance at 260 nm. RNA quality was determined by measuring the 260/280 ratio. Thereafter, first-strand cDNA synthesis was performed by using the HighCapacity cDNA reverse transcription kit (Applied Biosystems) according to the manufacturer’s instructions. The final reaction mix was kept at 25 8C for 10 min, heated to 37 8C for 120 min, heated for 85 8C for 5 s, and finally cooled to 4 8C.

Please cite this article in press as: El-Sherbeni AA, El-Kadi AOS. Alterations in cytochrome P450-derived arachidonic acid metabolism during pressure overload-induced cardiac hypertrophy. Biochem Pharmacol (2013), http://dx.doi.org/10.1016/j.bcp.2013.11.015

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2.6. Quantification by real time-PCR Quantitative analysis of specific mRNA expression was performed by real time-PCR, by subjecting the resulting cDNA to PCR amplification using 96-well optical reaction plates in the ABI Prism 7500 System (Applied Biosystems). 25-mL reaction mix contained 0.1 mL of 10 mM forward primer and 0.1 mL of 10 mM reverse primer, 12.5 mL of SYBR Green Universal Mastermix, 11.05 mL of nuclease-free water, and 1.25 mL of cDNA sample. The primers used in the current study were chosen from previously published studies and are listed in Table 1. No-template controls were incorporated onto the same plate to test for the contamination of any assay reagents. Thermocycling conditions were initiated at 95 8C for 10 min, followed by 40 PCR cycles of denaturation at 95 8C for 15 s, and annealing/extension at 60 8C for 1 min. Dissociation curves were performed by the end of each cycle to confirm the specificity of the primers and the purity of the final PCR product. 2.7. The extraction of AA metabolites from heart tissue 200 mg of heart tissue was homogenized on ice with 1 mL 100 mM potassium phosphate buffer (pH 7.4). The homogenates were centrifuged at 10,000  g for 15 min at 0 8C. The separated supernatant was added to 1 mL of methanol containing the deuterated metabolites (internal standards), butylated hydroxytoluene and ethylenediaminetetraacetic acid. Samples were centrifuged again for 15 min at 10,000  g at 0 8C. AA metabolites were extracted from the resulted supernatant by solid-phase cartridge (Oasis1HLB). Conditioning and equilibration of solidphase cartridge were performed with methanol, ethyl acetate, 0.2% formic acid (v/v), and 10% methanol, in sequence. After sample application, cartridges were washed by 0.2% formic acid (v/v), and 10% methanol in 0.2% formic acid (v/v), in sequence. Finally, AA metabolites were eluted by 1% formic acid in acetonitrile (v/v) and ethyl acetate, in sequence. Thereafter, the samples were evaporated to dryness, reconstituted with 50 mL acetonitrile and subjected to analysis. Epoxy-metabolites were measured as the sum of each EET and its corresponding DHET.

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The microsomal incubations were performed without adding sEH inhibitor, since the inhibition of sEH may activate the other pathways of EETs degradation [9]. Therefore, EETs formation rates were calculated based on the sum of each EET and its corresponding DHETs. All AA metabolites were found to be stable in phosphate buffer except 5,6-EET due to its spontaneous transformation to 5,6-d-lactone. Therefore, the method of Fulton et al. [31] was used to accurately measure 5,6-EET. Briefly, incubations were performed as aforementioned with 50 mM AA, thereafter, 5,6-EET and 5,6-d-lactone were transformed to the quite stable 5,6-DHET by camphorsulfonic acid and triethylamine, in sequence. Incubations with recombinant CYP enzymes were also performed. Incubation mixture (200 mL) containing 3 mM magnesium chloride hexahydrate, NADPH-P450 reductase, cytochrome b5 (40 pmole), CYP enzyme and a substrate in 100 mM potassium phosphate buffer (pH 7.4). CYP1B1 (20 pmol) was incubated with AA (100 mM) and NADPH (2 mM) for 10 min. Also, CYP2J2 (20 pmol), which is the ortholog of the commercially unavailable rat CYP2J3, was incubated with 12-HPETE (4 mg/mL) for 10 min, with or without NADPH (2 mM). The reaction was terminated by the addition of 600 mL ice-cold acidified acetonitrile and extracted twice by 1 mL ethyl acetate, then dried and reconstituted in 50 mL acetonitrile. 2.9. sEH activity assay sEH activity was measured using Morisseau and Hammock method with modifications [32]. 14,15-EET was used as the natural substrate for sEH [32]. Briefly, the cytosolic fraction was diluted to 0.4 mg/mL with sodium phosphate buffer (76 mM, pH 7.4) supplemented with BSA (2.5 mg/mL). The assay was initiated by the addition of 14,15-EET (final concentration of 14,15-EET was 2 mg/ml) final volume of incubates was 200 mL. The mixture was incubated at 37 8C for 10 min. The reaction was terminated by the addition of 600 mL ice cold acetonitrile followed by the internal standard, the deuterated 14,15-EET. Metabolites were extracted by 1 mL ethyl acetate twice and dried using speed vacuum (Savant, Farmingdale, NY).

2.8. AA incubations 2.10. HPLC methods Preliminary incubations were performed to ensure that the formation of AA metabolites was linear with respect to incubation time and protein content under assay conditions. A total microsomal protein concentration of 1 mg/mL was incubated with 50, or 100 mM AA for 30 min. The total volume of microsomal incubates was 200 mL of incubation buffer (3 mM magnesium chloride hexahydrate dissolved in 100 mM potassium phosphate buffer, pH 7.4). Incubations were conducted at 37 8C in a shaking water bath (90 rpm) after a 5-min pre-equilibration period. The reaction was initiated by the addition of the cofactor NADPH (final concentration 2 mM) and terminated by the addition of 600 mL icecold acidified acetonitrile. AA metabolites were extracted twice by 1 mL ethyl acetate and dried using speed vacuum (Savant, Farmingdale, NY, USA), then reconstituted in 50 mL acetonitrile.

Table 1 Primers sequences used for real-time PCRs. Gene

Forward primer

Reverse primer

ANP BNP a-MHC b-MHC EPHX2 GAPDH

GGAGCCTGCGAAGGTCAA CAGAAGCTGCTGGAGCTGATAAG ACAGAGTGCTTCGTGCCTGAT CGCTCAGTCATGGCGGAT GATTCTCATCAAGTGGCTGAAGAC CAAGGTCATCCATGACAACTTTG

TATCTTCGGTACCGGAAGCTGT TGTAGGGCCTTGGTCCTTTG CGAATTTCGGAGGGTTCTGC GCCCCAAATGCAGCCAT GGACACGCCACTGGCTAAAT GGGCCATCCACAGTCTTCTG

AA metabolites were analyzed using liquid chromatography– electrospray ionization-mass spectrometry (LC–ESI-MS) (Waters Micromass ZQ 4000 spectrometer). The method used was based on previously published papers with modifications [33–35]. The mass spectrometer was run in negative ionization mode with single ion monitoring. The nebulizer gas was acquired from an in house high purity nitrogen source. The temperature of the source was set at 150 8C, and the capillary and cone voltage were 3.51 kV and 25 V, respectively. The samples (20 mL) were separated on reverse phase C18 column (Kromasil, 250 mm  2.1 mm) at 30 8C. The mobile phase consisted of water/acetonitrile with 0.005% acetic acid and delivered using a linear gradient method at a flow rate of 200 mL/ min, as follows: 50–80% acetonitrile in 35 min, 80–100% acetonitrile in 5 min, and 100% for 6 min. The concentrations of the AA metabolites were calculated by comparing their corresponding analyte/reference response ratio based on peak area to calibration standards. For the CYP2J2-12-HPETE incubation, samples were resolved on reverse phase C18 column (Alltima HP, 250 mm  2.1 mm) at 25 8C. Mobile phase consisted of acetonitrile:water:formic acid (55:45:0.003) at flow rate of 0.3 mL/min. Detection was performed by UV at 210 nm. Column eluate corresponding to peaks of interest was collected for further analysis with MS. The source temperature was 150 8C, and cone voltage was 25 V.

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2.11. Western blot analysis Western blot analysis was performed using a previously described method [36]. Briefly, 20 mg of microsomal preparations were separated by 10% sodium dodecyl sulfate-polyacrylamide gel, and then electrophoretically transferred to nitrocellulose membrane. Protein blots were then blocked overnight at 4 8C in blocking solution containing 0.15 M sodium chloride, 3 mM potassium chloride, 25 mM Tris-base (TBS), 5% skim milk, 2% bovine serum albumin and 0.5% Tween-20. The blots were then incubated with a primary antibody for a 2 h incubation period (24 h for CYP2B1/2 antibody). Thereafter, blots were incubated with a peroxidaseconjugated secondary antibody for 1 h at room temperature. The bands were visualized using the enhanced chemiluminescence method according to the manufacturer’s instructions (GE Healthcare Life Sciences, Piscataway, NJ, USA). The intensity of the protein bands were quantified, relative to the signals obtained for GAPDH, using ImageJ software (National Institutes of Health, Bethesda, MD, http://rsb.info.nih.gov/ij). 2.12. Statistical analysis Statistical analysis was performed by SigmaPlot software, version 11.0. Data are presented as mean  S.E.M. Student’s t test or one-way analysis of variance followed by a Tukey’s post hoc test was performed in all data sets. All result was considered statistically significant where p < 0.05. Spearman correlation coefficients (r) were used for evaluating the possible association between AA metabolites formation and CYP enzymes expression.

3. Results 3.1. The effect of DAC on the development of cardiac hypertrophy Five weeks post-surgery, DAC animals showed a 28% increase in left ventricle weight to tibial length ratio compared to sham (Fig. 1A). Echocardigraphic analysis showed a significant thickening of the left ventricle posterior wall thickness and interventricular septal wall thickness by 31% and 28%, respectively in DAC hearts compared to sham (Fig. 1A). On the other hand, left ventricle volume showed a significant decrease by 14% in DAC hearts compared to sham (Fig. 1A). Regarding heart functions, the percentage of fractional shortening and the percentage of ejection fraction did not change, while, tie index showed a significant increase by 14% in DAC hearts compared to sham (Fig. 1B). Additionally, atrial natriuretic peptide (ANP), brain natriuretic peptide (BNP) and b/a-myosin heavy chain (b/a-MHC) mRNA were significantly increased by 166%, 678% and 3658%, respectively, in DAC hearts compared to sham (Fig. 1C). Our results showed that compensated cardiac hypertrophy was developed 5 weeks post-surgery.

Fig. 1. Assessment of cardiac hypertrophy. Five-week post-surgery, rats were weighed and mildly anesthetized for echocardiography to measure: heart dimensions parameters (A), typified by left ventricular weight to tibial length (LV wt/tl), left ventricle posterior wall thickness (LVPW), interventricular septal wall thickness (IVS), and left ventricle volume (LVV) during diastole, and heart functions (B), typified by percentage of fractional shortening (%FS), ejection fraction (%EF), and Tei index. Thereafter, hearts were harvested and total RNA was isolated from hearts of sham and DAC animals. Gene expressions were determined by realtime PCR for hypertrophic markers (C), typified by ANP, BNP and b/a myosin heavy chain. Results are expressed as mean  S.E.M. *p < 0.05.

3.2. Effect of cardiac hypertrophy on the levels of AA metabolites in heart tissue The HPLC method provided good resolution for all formed metabolites, 4 EETs, 4 DHETs, 3 mid-chain and 5 v/v1 HETEs, except the coelution of 16- and 17-HETE. The retention time, and ions monitored are shown in Table 2. Both EETs and DHETs quantification has been a challenging task due to their low levels in heart tissue. We measured EETs levels indirectly after their conversion to DHET, therefore, the measured levels were, in fact, the sum of EET and its corresponding DHET levels. Thus, it reflected the level of endogenous epoxy-metabolites of AA in heart tissue. There was a significant increase in cardiac 5,6-, 8,9-, 11,12- and

14,15-EET + DHET levels by 89%, 76%, 62% and 69%, respectively, in DAC animals compared to sham (Fig. 2A). Mid-chain HETEs, 5-, 12-, and 15-HETE, were found to be higher in heart tissue compared to EETs or DHETs. In DAC heart there was a significant increase in 5-, 12-, and 15-HETE levels by 52%, 100% and 31%, respectively, compared to sham (Fig. 2B). With respect to v/v1 HETEs, the endogenous concentrations of 16-, 17-, 18-, and 19-HETE could not be detected in either DAC or sham hearts (Fig. 2C). While for 20-HETE, its cardiac level was comparable to EETs and DHETs, and there was a significant increase in its level in heart tissue of DAC animals by 341% compared to sham (Fig. 2C).

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Table 2 The retention times, and ions monitored for the CYP-derived AA metabolites analyzed. Metabolite

Retention time (min)

m/z

14,15-DHET d11-14,15-DHET 11,12-DHET 8,9-DHET 5,6-DHET 19-HETE 20-HETE 18-HETE 16/17-HETE 15-HETE d8-15-HETE 12-HETE 5-HETE 14,15-EET d11-14,15-EET 11,12-EET 8,9-EET 5,6-EET

16.8 16.4 18.4 19.6 21.2 21 21.6 22.6 23.5 26 25.4 28.3 30 32.7 32.4 34.6 35.5 36.2

319 348 319 319 319 319 319 319 319 319 327 319 319 319 330 319 319 319

3.3. Effect of cardiac hypertrophy on the formation and degradation of EETs With respect to EETs formation rate, there was a 100%, 179%, 179% and 303% increase in 5,6-, 8,9-, 11,12-, and 14,15-EET formation rate, respectively, in DAC hearts compared to sham (Fig. 3A). Moreover, there was an alteration in the regioselectivity of epoxide group insertion. Regioselectivity expressed as 14,15EET:11,12-EET:8,9-EET ratio was 1.2:1.2:1 for normal and 1.7:1.2:1 for DAC hearts (Fig. 3A). The observed increase in EETs formation rate was in line with the observed increase in the levels of EETs + DHETs in heart tissue during cardiac hypertrophy. In order to determine whether or not cardiac hypertrophy also altered the degradation of EETs to DHETs, sEH activity, and expression were measured. sEH is the dominant enzyme responsible for catalyzing the transformation of EETs to DHETs. Interestingly, the activity of sEH significantly decreased by 30% in DAC hearts compared to sham (Fig. 3B). Additionally, EPHX2 gene expression was significantly decreased in DAC heart by 98% compared to sham (Fig. 3C). 3.4. Effect of cardiac hypertrophy on the formation of HETEs Lipoxygenases are also known to catalyze the formation of mid-chain HETEs, therefore, the formation of mid-chain HETEs, 5-, 12-, and 15-HETE was determined in microsomal incubates with and without CYP activation by NADPH. Despite lipoxygenases are mainly cytosolic, lipoxygenase activity was previously reported in microsomal fraction [37]. In consistence with the published data, the heart microsomal fraction was able to form only the 12-HETE without NADPH. By adding NADPH, the rate of 12-HETE formation was significantly increased, whereas, 5- and 15-HETE formations were still below the detection limit. 12-HETE formation mediated by the microsomal fraction of sham hearts was 4.48 and 3.64 pmole/min/mg protein with and without NADPH, respectively. For DAC hearts, 12-HETE formation was 11.41 and 6.64 pmole/min/mg protein with and without NADPH, respectively. In DAC hearts, there was a significant increase in NADPHdependent 12-HETE formation rate by 470% compared to sham (Fig. 4A). In contrast, several v/v1 HETEs were formed by heart microsomes after CYP activation by NADPH. Except for 18-HETE, the formation of 16/17-, 19-, and 20-HETE could be detected in DAC, and sham hearts (Fig. 4B). Albeit, 16/17-HETE formation rate

Fig. 2. Effect of cardiac hypertrophy on cardiac AA metabolites levels. Heart tissue (200 mg) was homogenized on ice with 1 mL 100 mM potassium phosphate buffer (pH 7.4). The homogenates were centrifuged at 10,000  g for 15 min at 0 8C. The separated supernatant was added to 1 mL of methanol containing the deuterated metabolites (internal standards), butylated hydroxytoluene, and ethylenediaminetetraacetic acid. The samples were centrifuged again for 15 min at 10,000  g at 0 8C. AA metabolites were extracted from the resulted supernatant by solid-phase cartridge (Oasis1HLB). Samples were evaporated to dryness, then, reconstituted and the sum of each EETs and its corresponding DHET (A), mid-chain HETEs (B), and v/v1 HETEs (C) were measured using LC–ESI-MS as described under Section 2. Results are expressed as mean  S.E.M. *p < 0.05.

showed no significant changes, 19-HETE formation rate decreased significantly by 41% in DAC hearts compared to sham (Fig. 4B). On the other hand, 20-HETE formation rate was slightly higher but did not reach the statistical significance (p = 0.06), in DAC hearts compared to sham (Fig. 4B). 3.5. Effect of cardiac hypertrophy on CYP protein expression In order to determine main CYP enzymes involved in the observed alteration in AA metabolism, two-step process was performed. First, the effect of DAC-induced cardiac hypertrophy on CYP protein expression was determined by Western blot analysis. Second, we tested the correlation between CYP expression and the measured AA metabolites formation rates, as previously described [38,39].

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Fig. 3. Effect of cardiac hypertrophy on the formation and degradation of EETs in the heart. Heart microsomal fraction was incubated with 50 mM arachidonic acid to determine EETs formation (A). Also, heart cytosol was incubated with 2 mg/ml 14,15-EET to determine sEH activity, i.e. EETs degradation (B). sEH activity was determined by measuring the formation rate of 14,15-DHET. The reaction was lasted for 30 min or 10 min for EETs formation and degradation experiment, respectively. The reaction was terminated by the addition of ice-cold acidified acetonitrile. Metabolites were extracted by ethyl acetate and dried. Reconstituted metabolites were injected into LC–ESI-MS as described under Section 2. Total RNA was isolated from hearts of sham and DAC animals for the determination of EPHX2 gene expression (C) by real-time PCR. Results are expressed as mean  S.E.M. *p < 0.05.

Fig. 4. Effect of cardiac hypertrophy on the formation of HETEs in the heart. Heart microsomal fraction was incubated with 100 mM arachidonic acid. The reaction was lasted for 30 min, then, metabolites were extracted by ethyl acetate. The samples were evaporated to dryness, then, reconstituted and mid-chain HETEs (A), as well as, v/v1 HETEs (B) were measured using LC–ESI-MS as described under Section 2. 18-HETE standard curve was used to interpret the signals of coeluted metabolites, 16- and 17-HETE, since standard curves for all HETEs were nearly identical. Results are expressed as mean  S.E.M. *p < 0.05.

12-HETE suggesting that CYP1B1 was involved in 12-HETE formation; r value was 0.73 (Fig. 6). Furthermore, CYP4As was found to correlate significantly with 19-HETE (r = 0.61) (Fig. 6). Interestingly, CYP2J3 expression exhibited significant inverse correlation with 12-HETE, r value was 0.67 (Fig. 6). 3.6. Role of CYP1B1 and CYP2J2 in mid-chain HETEs metabolism

Regarding CYP expression in the cardiac microsomal fraction, several CYP enzymes, namely CYP1As, CYP1B1, CYP2Bs, CYP2C11, CYP2E1, CYP2J3, and CYP4As, were measured using Western blot analysis (Fig. 5). For CYP1 family, CYP1As expression was significantly decreased by 76%, whereas CYP1B1 expression was significantly induced by 100% in DAC hearts compared to sham (Fig. 5). With regard to CYP2 family, CYP2Bs expression significantly increased by 72% in DAC hearts compared to sham (Fig. 5), whereas, CYP2J3 expression was significantly decreased by 31% in DAC hearts compared to sham (Fig. 5). CYP2C11, and CYP2E1 expressions were not significantly altered by cardiac hypertrophy (Fig. 5). For CYP4 family, CYP4As expression was decreased by 65% in DAC hearts compared to sham, while the expression of CYP4Fs was not altered (Fig. 5). The correlation analysis based on the fact that if a CYP enzyme is predominantly involved in the formation of a specific AA metabolite, we will observe that the higher the expression of this enzyme, the higher the formation rate of its metabolites, and vice versa. Using non-parametric correlation analysis, CYP2Bs was found to be correlated significantly with 5,6-, 8,9-, 11,12- and 14,15-EET; r values were 0.61, 0.71, 0.78 and 0.72, respectively (Fig. 6). Significant correlation was observed between CYP1B1 and

In order to gain insight about the role of CYP1B1, and CYP2J3 in mid-chain HETEs formation, human recombinant CYP1B1, and CYP2J2 were examined. CYP1B1 was found to mediate the metabolism of AA to 5-, 12-, and 15-HETE at rates comparable to that of 12-HETE formation by the cardiac microsomal fraction (Fig. 7). On the other hand, CYP2J2 did not catalyze the degradation of mid-chain HETEs, 5-, 12-, and 15-HETE. We hypothesized that CYP2J2 decreases the formation of 12-HETE by consuming its precursor, 12-HPETE. To test this hypothesis, we incubated 12HPETE with CYP2J2. Our results showed that CYP2J2 did catalyze the degradation of 12-HPETE to hepoxilins, which are hydroxyepoxyeicosatrienoic acids, based on its retention time and its mass spectrum (Fig. 8). Only after the addition of NADPH, hepoxilins peak was detected in CYP2J2-12-HPETE chromatogram (Fig. 8A). The CYP2J2-mediated formation of hepoxilins was accompanied by a consumption of 12-HPETE (Fig. 8B). Mass spectrum showed base peak corresponds to hepoxilin molecular ion, [MH], at m/z = 335 and two fragment ions peaks at m/ z = 317 and 273 correspond to [MH–H2O] and [MH–H2O– CO2], respectively (Fig. 8C). This is in consistent with the reported mass spectrum of hepoxilins [40].

Please cite this article in press as: El-Sherbeni AA, El-Kadi AOS. Alterations in cytochrome P450-derived arachidonic acid metabolism during pressure overload-induced cardiac hypertrophy. Biochem Pharmacol (2013), http://dx.doi.org/10.1016/j.bcp.2013.11.015

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Fig. 7. Formation of mid-chain HETEs by recombinant CYP1B1. The reconstituted system consisted of 20 pmol CYP1B1, and 100 mM arachidonic acid with 40 pmol cytochrome b5. Metabolites were extracted by ethyl acetate and dried. Reconstituted metabolites were injected into LC–ESI-MS as described under Section 2. Results are expressed as mean  S.E.M. (n = 3). *p < 0.05.

Fig. 5. Effect of cardiac hypertrophy on CYP protein expression in the heart. Microsomal protein was isolated from the heart and separated on a 10% sodium dodecyl sulfate-polyacrylamide gel. CYP1As, CYP1B1, CYP2Bs, CYP2C11, CYP2E1, CYP2J3, CYP4As, CYP4Fs, and GAPDH proteins were detected by the enhance chemiluminescence method as described under Section 2. The graph represents the amount of protein normalized to the loading control (mean  S.E.M.), and expressed as a percentage of the control. *p < 0.05.

4. Discussion In the current study, we constricted the descending aorta to induce compensated cardiac hypertrophy in Sprague Dawley rats. Progression of cardiac hypertrophy was gradual over 5 weeks as

assessed by multiple echocardiography (data not shown), with no sign of the development of heart failure. The absence of systolic heart functions deterioration indicated that the heart was able to compensate the pressure overload via cardiac remodeling [3]. Several cardiac hypertrophic models have been used to examine the alteration in the formation rates of some CYP-derived AA metabolites [41–44]. In the current study, alterations in the cardiac levels of almost all CYP-derived AA metabolites as well as their formation rates were investigated during cardiac hypertrophy. Additionally, the cardiac hypertrophy model (DAC) model, used in the current study has several advantages over the previously used models; notably, no exogenous agents, such as isoproterenol or doxorubicin, were given to induce hypertrophy. These exogenous agents may have their own effect on CYP-mediated AA metabolism. Moreover, cardiac hypertrophy is developed over relatively longer period of time, thus, DAC model is more clinically relevant [45]. The alterations in CYP-mediated AA metabolism during cardiac hypertrophy were determined by measuring the primary

Fig. 6. Correlation between CYP protein expression levels and the formation rate of CYP-derived AA metabolites in individual heart samples. Spearman correlation coefficients (r) were used for evaluating the possible association. Tendency of correlation is indicated by ascending or descending lines, however, the correlation is not necessarily linear because spearman correlation analysis is a non-parametric method. Correlations shown were statistically significant according to p-value (p < 0.05).

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Fig. 8. Degradation of 12-HETE by recombinant CYP2J2. The reconstituted system consisted of 20 pmol CYP2J2 and 4 mg/mL 12-HPETE with and without NADPH as described under Section 2. The chromatograms (A) and areas for 12-HPETE and hepoxilins peaks (B) with and without NADPH were determined by resolving the metabolites with reverse phase HPLC C18 column (Alltima HP, 250 mm  2.1 mm) at 25 8C. Mobile phase consisted of 55% acetonitrile:water:formic acid (55:45:0.003) at flow rate of 0.3 mL/ min. Detection was performed by UV at 210 nm. Mass spectrum (C) was determined by collecting column eluate corresponding to hepoxilins peak for further analysis with ESI coupled with single quadrupole MS. The source temperature was 150 8C, and cone voltage was 25 V. Results are expressed as mean  S.E.M. (n = 3). *p < 0.05.

oxygenation products of the AA CYP monoxygenase. The cardiac levels and formation rates of 16 metabolites, 4 EETs, 4 DHETs, 3 midchain and 5 v/v1 HETEs, were quantified. HPLC method used in the current study allowed the simultaneous measurement of all metabolites. Good resolution was obtained for the tested metabolites except the coelution of 16- and 17-HETE. 18-HETE standard curve was used to interpret the signals of coeluted metabolites, since standard curves for all HETEs were nearly identical. The instability problem of 5,6-EET led to a general notion that 5,6-EET role is underestimated [46,47]. Therefore, a previously published method was used to solve the instability problem [31], thereby, 5,6-EET alteration was accurately evaluated for the first time during cardiac hypertrophy. The scope of the current study was limited to CYPderived metabolites, therefore, cyclooxygenases and lipoxygenases expression and activity were not measured. Pressure overload-induced cardiac hypertrophy significantly increased EETs levels in the heart tissue. This increase in cardiac EETs levels was also confirmed in vitro by measuring their formation, and degradation rates by cardiac microsomal and cytosolic fractions, respectively. The induction of EETs in hypertrophied hearts was not reported before. In fact, isoproterenol-, doxorubicin-, benzo(a)pyrene-,

and arsenic-induced cardiac hypertrophy were found to inhibit EETs formation [41–44]. Several factors may explain why pressure overload model of cardiac hypertrophy causes an increase in EETs levels. First, the stretching of cardiomyocytes by pressure overload itself could directly increase EETs levels. It has been previously reported that shear stress in smooth muscles was able to increase the levels of EETs [48,49]. Second, the longer duration of time needed to develop cardiac hypertrophy in DAC model may alter the response on EETs. The previous models developed acute cardiac hypertrophy within hours or few days, in contrast, DAC induced cardiac hypertrophy over a longer period of time (several weeks). In this context, it has been previously reported that there was an increase in EETs plasma levels, and a decrease of sEH activity in patients with chronic cardiovascular disease compared to healthy volunteers which is in agreement with our results [50]. Our results may explain the failure of sEH inhibitors in the prevention or treatment of pressure overload-induced cardiac hypertrophy [51], despite the success of these inhibitors in acute models of cardiac hypertrophy [44,52]. Evidently, EETs have cardioprotective effects against cardiac hypertrophy [9]. Therefore, we believe that EETs induction is an adaptive response to pressure overload insult.

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With respect to v/v1 HETEs, only the 20-HETE level was found to increase in heart tissue during cardiac hypertrophy, whereas, other v/v1 HETEs, 16-, 17-, 18- and 19-HETE were undetectable. Also, the formation of 18-HETE was not detected in cardiac microsomal incubates, therefore, we can conclude that Sprague Dawley heart tissue does not produce substantial amounts of 18-HETE. The formation rate of 16/17-HETE was not altered during cardiac hypertrophy, consequently, it is expected that their levels in heart tissue would not be altered. For the 19-HETE, there was a significant decrease in its formation rate during cardiac hypertrophy, hence, an equivalent decrease in its cardiac level is expected. In summary, there was a significant increase in the cardiotoxic 20-HETE and a decrease in its endogenous antagonist, 19-HETE, during pressure overload-induced cardiac hypertrophy. The observed increase in 20-HETE is in agreement with the previously reported data for cardiac hypertrophy regardless of the model of cardiac hypertrophy used [41,44,52,53]. Despite the significant increase in the endogenous levels of mid-chain HETEs in heart tissue during cardiac hypertrophy, only the increase in 12-HETE could be linked to CYP enzymes. However, the role of CYP in regulating mid-chain HETEs levels in heart tissue requires further investigation. It is worth to note that the percent increase in 12-HETE level in heart tissue was about 1/5 of the percent increase in the formation of 12-HETE by heart microsomal fraction. To our knowledge, this is the first time to report that pressure overload-induced cardiac hypertrophy is associated with an increase in the cardiac mid-chain HETEs levels. However, several previous studies did report the detrimental role of midchain HETEs in the development of cardiac hypertrophy and heart failure [27,54–56]. As aforementioned above, cardiac levels of CYP-derived AA metabolites were altered during cardiac hypertrophy. Apparently, these metabolites have a role in the pathogenesis and progression of cardiac hypertrophy, most probably by inducing inflammation and fibrosis in the heart. In fact, inflammation and fibrosis represent key events in cardiac hypertrophy. Evidently, the expression of inflammatory markers has been shown to precede any manifestation of cardiac hypertrophy [57]. Also, several reports demonstrated the association between the magnitude of the expression of inflammatory markers and the deterioration of heart functions [58–60]. Furthermore, the induction and prevention of cardiac hypertrophy were reported to be achievable by altering inflammatory mediators in vivo and in vitro [57,61–63]. In contrast to inflammation, fibrosis is not associated with the initial stages of cardiac hypertrophy, but involved in the deterioration of heart functions [64,65]. The stiffness of the heart muscles due to fibrosis leads to increase relative workload per myocyte, resulting in further damage [64]. Thereby, progressive deterioration in the heart functions is occurred during the transition from compensated to decompensated cardiac hypertrophy that eventually leads to heart failure. Collectively, we suggest that the increase in cardiac levels of mid-chain and 20-HETEs mediate, at least in part, the pathogenesis and the progression of cardiac hypertrophy. In addition, the cardiotoxic effects of 20-HETE should be more pronounced due to the decrease in the cardiac levels of 19-HETE, which is the endogenous antagonist for 20-HETE [20,21]. To identify novel targets for the prevention and treatment of cardiac hypertrophy and heart failure, we attempted to identify the CYP enzymes involved in the observed alteration in CYP-mediated AA metabolism. We reported elsewhere that EETs formation in normal hearts microsomes reflects the regioselectivity of CYP2C11 [66]. In the current study, we found that regioselectivity was altered for 14,15-EET:11,12-EET:8,9–EET ratio from 1.20:1.21:1 to 1.73:1.20:1 during cardiac hypertrophy. CYP2C11 has a characteristic feature of producing comparable amounts of 11,12- and 14,15-EET [8]. Only CYP2J3 and CYP2B2 are characterized by the

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formation of 14,15-EET as the major EETs [8,67]. Therefore, it was expected to observe an induction in CYP2J3 and/or CYP2B2 expression in hypertrophied hearts. In the current study, CYP2J3 enzyme was downregulated, while CYP2Bs was induced. Moreover, using non-parametric correlation analysis, CYP2Bs expression was significantly associated with EET formation rate. These data strongly suggest that the increase in EETs formation was mainly due to CYP2B2 induction. CYP1As, CYP2E1 and CYP4A2 were reported to produce 19HETE [8,12,13]. In the current study, the decrease in 19-HETE was only correlated with CYP4As expression. CYP4A subfamily includes three enzymes, CYP4A1 which hydroxylates AA at C20 mainly [8], and CYP4A2, as well as, CYP4A3 which produce 19-HETE [68]. Therefore, our findings suggest that the decrease in CYP4A2 and/or CYP4A3 expression was responsible for the observed decrease in 19-HETE formation. With respect to mid-chain HETEs, correlation analysis showed that CYP1B1 was involved in 12-HETE formation, whereas, CYP2J3 was involved in 12-HETE degradation. The recombinant CYP1B1 catalyzed the formation of 5-, 12- and 15-HETE similar to what has been previously reported [10]. Hence, CYP1B1 is responsible, in part, for 12-HETE endogenous formation. For CYP2J3, it has been previously reported that CYP2J2 exhibits hydroperoxidase isomerase activity and can metabolize 15-HPETE to hepoxilins before being transformed to mid-chain HETEs [40]. Here, we reported that the metabolism of 12-HPETE to hepoxilins can also occur via CYP2J2. One of the beneficial roles of CYP2Js in the heart is mediating the degradation of the cardiotoxic 12- and 15-HETE in favor of the formation of the cardioprotective metabolite, hepoxilins. In the current study, pressure overload-induced cardiac hypertrophy showed alterations in CYP-mediated AA metabolic profile. These alterations were characterized, and subsequently, CYP enzymes involved in these alterations were identified. Our findings suggest that in cardiac hypertrophy CYP1B1 has a detrimental effect on the heart by mediating the formation of the cardiotoxic mid-chain HETEs. On the other hand, CYP2B2, CYP2J3 and CYP4A2/3 play cardioprotective roles. CYP2B2 plays a major role in the formation of the cardioprotective EETs during cardiac hypertrophy, whereas, CYP4A2/3 mediates the formation of the cardioprotective metabolite, 19-HETE. Interestingly, CYP2J3 mediates the decrease in the cardiotoxic mid-chain HETEs levels, and the formation of the cardioprotective hepoxilins, in addition to its reported contribution to the formation of cardioprotective metabolite, EETs. Hence, our study may lead to a new biochemical targets for the development of drugs to treat and prevent cardiac hypertrophy and heart failure. Acknowledgements This work was supported by a grant from the Canadian Institutes of Health Research (CIHR) [Grant MOP 106665]. AAE is the recipient of Egyptian Government Scholarship. The authors are grateful to Anwar Anwar-Mohamed for his helpful comments. The authors would like to thank Dr. Vishwa Somayaji for his excellent technical assistance with liquid chromatography–electron spray ionization-mass spectrometry. References [1] Frey N, Olson EN. Cardiac hypertrophy: the good, the bad, and the ugly. Annu Rev Physiol 2003;65:45–79. [2] Nagata K, Liao R, Eberli FR, Satoh N, Chevalier B, Apstein CS, et al. Early changes in excitation–contraction coupling: transition from compensated hypertrophy to failure in Dahl salt-sensitive rat myocytes. Cardiovasc Res 1998;37:467–77. [3] Sasayama S, Kihara Y, Matsumori A, Hasegawa K. Transition from compensated to decompensated cardiac hypertrophy. Heart Fail Rev 1999;4.

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Please cite this article in press as: El-Sherbeni AA, El-Kadi AOS. Alterations in cytochrome P450-derived arachidonic acid metabolism during pressure overload-induced cardiac hypertrophy. Biochem Pharmacol (2013), http://dx.doi.org/10.1016/j.bcp.2013.11.015

Alterations in cytochrome P450-derived arachidonic acid metabolism during pressure overload-induced cardiac hypertrophy.

Cardiac hypertrophy is a major risk factor for many serious heart diseases. Recent data demonstrated the role of cytochrome P450 (CYP)-derived arachid...
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