Effect of Soluble Epoxide Hydrolase on the Modulation of Coronary Reactive Hyperemia: Role of Oxylipins and PPARγ

Coronary reactive hyperemia (CRH) is a physiological response to ischemic insult that prevents the potential harm associated with an interruption of blood supply. The relationship between the pharmacologic inhibition of soluble epoxide hydrolase (sEH) and CRH response to a brief ischemia is not known. sEH is involved in the main catabolic pathway of epoxyeicosatrienoic acids (EETs), which are converted into dihydroxyeicosatrienoic acids (DHETs). EETs protect against ischemia/reperfusion injury and have numerous beneficial physiological effects. We hypothesized that inhibition of sEH by t-AUCB enhances CRH in isolated mouse hearts through changing the oxylipin profiles, including an increase in EETs/DHETs ratio. Compared to controls, t-AUCB–treated mice had increased CRH, including repayment volume (RV), repayment duration, and repayment/debt ratio (p < 0.05). Treatment with t-AUCB significantly changed oxylipin profiles, including an increase in EET/DHET ratio, increase in EpOME/DiHOME ratio, increase in the levels of HODEs, decrease in the levels of mid-chain HETEs, and decrease in prostanoids (p < 0.05). Treatment with MS-PPOH (CYP epoxygenase inhibitor) reduced CRH, including RV (p < 0.05). Involvement of PPARγ in the modulation of CRH was demonstrated using a PPARγ-antagonist (T0070907) and a PPARγ-agonist (rosiglitazone). T0070907 reduced CRH (p < 0.05), whereas rosiglitazone enhanced CRH (p < 0.05) in isolated mouse hearts compared to the non-treated. These data demonstrate that sEH inhibition enhances, whereas CYP epoxygenases-inhibition attenuates CRH, PPARγ mediate CRH downstream of the CYP epoxygenases-EET pathway, and the changes in oxylipin profiles associated with sEH-inhibition collectively contributed to the enhanced CRH.


Introduction
All animal care and experimentation protocols were submitted to, approved, and carried out in accordance with the West Virginia University Institutional Animal Care and Use Committee and were in accordance with the principles and guidelines of the NIH's Guide for the Care and Use of Laboratory Animals. Both male and female mice (14-16 wks old) in equal ratio were used in our study. Mice were maintained in cages with a 12:12 h light-dark cycle and free access to standard chow and water.

Langendorff-Perfused Heart Preparation
There are two available Langendorff technique modes to perfuse the heart: 1) constant pressure mode and 2) constant flow mode. In our experiments, we selected the constant pressure mode. Because the two modes have important differences, the selection between the two is particularly important when assessing the role of coronary regulatory mechanisms in response to changing coronary flow (CF) conditions or pathologies, such as ischemia. The constant pressure mode is physiologically more relevant in experiments involving ischemia, like the model in our experiments; therefore, it was selected over the other mode [32].
Soluble epoxide hydrolase null (sEH -/-) and wild-type (WT; sEH +/+ ) mice (14-16 wks.) of both sexes (equal ratios) were euthanized with sodium pentobarbital (100 mg/kg body weight intra-peritoneally). Hearts were excised and immediately placed into heparinized (5 U/mL) ice-cold Krebs-Henseleit buffer containing (in mM) 119.0 NaCl, 11.0 glucose, 22.0 NaHCO 3 , 4.7 KCl, 1.2 KH 2 PO 4 , 1.2 MgSO 4 , 2.5 CaCl 2 , 2.0 pyruvate, and 0.5 EDTA. After removal of the lungs and tissue surrounding the heart, the aorta was rapidly cannulated with a 20-gauge, blunt-ended needle and continuously perfused with 37°C buffer continuously bubbled with [95% O 2 ]-[5% CO 2 ] at a constant perfusion pressure of 80 mmHg. The left atrium was excised, and a water-filled balloon made of plastic wrap was inserted into the left ventricle through the mitral valve. The balloon was connected to a pressure transducer for continuous measurement of left ventricular developed pressure (LVDP) and heart rate (HR). The heart was then immersed in a water-jacketed perfusate bath (37°C) and left to beat spontaneously. Left ventricular diastolic pressure was adjusted to 2-5 mmHg. A flow transducer was installed above the cannulated aorta for continuous measurement of CF with an ultrasonic flow probe (Transonic Systems, Ithaca, NY). A Power-Lab Chart data acquisition system (AD Instruments, Colorado Springs, CO) was used for data acquisition. Heart function was allowed to stabilize for 30-40 min before initiation of CRH. Only hearts whose CF increased by more than two fold after a 15-second total occlusion were included in the analysis. This demonstrated that the isolated heart preparation was intact to be included in the experiment. Hearts with persistent arrhythmias or LVDP <80 mmHg were excluded.

Effect of t-AUCB on CRH Response
After stabilization for 30-40 min, isolated WT mouse hearts were subjected to 15 sec of total occlusion. Recordings of the first CRH (baseline CF, CF tracing, LVDP, HR, RV, PHF, and RD) were analyzed for each heart and averaged. t-AUCB was infused at a final concentration of 10 μM and 1% of CF rate for 15 min, after which another CRH was induced and the same parameters (baseline CF, CF tracing, LVDP, HR, RV, and RD) were recorded again and analyzed.
Effect of t-AUCB on CRH Response in sEH -/-Mice: To investigate the specificity of the sEH-inhibitor, t-AUCB, we evaluated its effect on CRH in sEH -/mouse hearts, where the sEH gene is globally deleted. We followed the same protocol described above. CRH was induced in sEH -/mice before and after infusing t-AUCB for 15 min. The two CRH responses were analyzed.

LC-MS/MS Oxylipin Analysis
Levels of oxylipins (5,8,11,(12)(13)(14)5,8,11,(12)(13)(14)12-and 15-HETE, 9,10-and 12,13-EpOME, 9,10-and 12,13-DiHOME, 9-and 13-HODE, 6-keto prostaglandin-F 1α [6K-PG-F 1α ], PG-F 2α , thromboxane B 2 [TxB 2 ], PGD 2 , and PGE 2 ) were determined in pre-and post-CRH heart perfusates of WT and t-AUCBtreated WT mice through liquid chromatography, tandem mass spectroscopy (LC-MS/MS) as described previously [36]. Heart perfusates were collected after the first 30 min of stabilization and right after reperfusion for 2.5 min. Hearts were immersed in 5 mL of warm (37°C) Krebs-Henseleit buffer with 5 μL of 10 μM t-AUCB to block further EET breakdown by sEH. Heart perfusates were collected two times before ischemia (baseline) and pooled together as one sample and two times after ischemia and pooled together as another sample for LC-MS/MS analysis. Samples were stored at -80°C until processing. Samples were spiked with 30 ng PGE2-d4, 10,11-DiHN, and 10,11-EpHep (Cayman) as internal standards, mixed with 0.1 vol of 1% acetic acid in 50% methanol, and extracted by serial passage through Oasis HLB C18 3mL columns (Waters, Milford, MA, USA). Columns were washed twice with 0.1% acetic acid in 5% methanol and eluted with methanol into glass tubes containing 6 μL of 30% glycerol in methanol. The methanol was then evaporated under a stream of nitrogen gas, and the dried tubes were frozen and stored at -80°C until analysis. Online liquid chromatography of extracted samples was performed with an Agilent 1200 Series capillary HPLC (Agilent Technologies, Santa Clara, CA, USA). Separations were achieved using a Halo C18 column (2.7 mm, 10062.1 mm; MAC-MOD Analytical, Chadds Ford, PA), which was held at 50°C. Mobile phase A was 85:15:0.1 water: acetonitrile: acetic acid. Mobile phase B was 70:30:0.1 acetonitrile: methanol: acetic acid. Flow rate was 400 μL/min; Gradient elution was used. Mobile phase percentage B and flow rate were varied as follows: 20% B at 0 min, ramp from 0 to 5 min to 40% B, ramp from 5 to 7 min to 55% B, ramp from 7 to 13 min to 64% B. From 13 to 19 min the column was flushed with 100% B at a flow rate of 550 μL/min. Samples were solvated in 50 μl of 30% ethanol. The injection volume was 10 μL. Samples were analyzed in triplicate. Analyses were performed on an MDS Sciex API 3000 equipped with a TurboIonSpray source (Applied Biosystems). Turbo desolvation gas was heated to 425°C at a flow rate of 6 L/min. Negative ion electrospray ionization tandem mass spectrometry with multiple reaction monitoring was used for detection.

Effect of MS-PPOH (CYP epoxygenase inhibitor) on CRH Response
Isolated WT mice hearts were stabilized for 30-40 min, followed by 15 sec of total occlusion. Recordings of the first CRH (baseline CF, CF tracing, LVDP, HR, RV, PHF and RD) were analyzed for each heart and averaged. MS-PPOH was infused at a final concentration of 1.0 μM for 15 minutes, after which the second CRH was induced. CRHs before and after MS-PPOH infusion were analyzed and compared.

Effect of T0070907 (PPARγ antagonist) on CRH Response
Isolated WT mouse hearts were stabilized for 30-40 min followed by 15 sec of total occlusion. Recordings of the first CRH (baseline CF, CF tracing, LVDP, HR, RV, PHF and RD) were analyzed for each heart and averaged for each group, as mentioned previously. T0070907 was infused at a final concentration of 10 μM and 1% of CF rate for 15 min, after which another CRH was induced and the same parameters (baseline CF, CF tracing, LVDP, HR, RV, and RD) were recorded again and analyzed.

Effect of t-AUCB and T0070907 on CRH Response
In this experiment, we investigated a possible link between the role of PPARγ in CRH and pharmacological inhibition of sEH by t-AUCB. We evaluated the effects of selective sEH-inhibition on CRH followed by PPARγ-antagonist. Baseline CRH was induced in WT mice. t-AUCB was infused at a final concentration of 10 μM for 15 minutes, after which a second CRH was induced. T0070907 was infused while t-AUCB was still being infused, at a final concentration of 10 μM for another 15 min, and a third CRH was induced. The three CRH responses were analyzed and compared.

Effect of Rosiglitazone on CRH Response
After stabilization, WT mouse hearts were subjected to 15 sec of total occlusion. As described above, baseline CRH was induced in each mouse heart. Rosiglitazone was infused at a final concentration of 10 μM for 15 min, followed by another CRH. CRHs before and after rosiglitazone infusion were analyzed and compared.

Statistical and Data Analyses
Flow debt (baseline flow rate multiplied by occlusion duration) and RV (the integral of hyperemic area above the baseline flow) were calculated using "the integral relative to baseline" function in the data pad of Lab-Chart 7.0 software. Since absolute coronary flow rates change proportionally with heart mass, the RV and flow debt are presented as ml/g wet heart weight, and baseline and peak flow rate data are presented as (mL.min -1 .g wet heart weight -1 ). Values are means ± standard error; n represents the number of animals. For data analysis, two-tailed paired t-tests were used for paired data analysis, repeated measures ANOVAs were used for populations measured 3 times, and two-way ANOVAs were used to compare data between groups. Differences were considered statistically significant when p < 0.05.

Discussion
Pharmacologic inhibition of sEH in WT mice enhanced CRH in an isolated heart model after brief ischemia. sEH inhibition was also associated with changes in various oxylipin profiles. The relationship among inhibition of CYP epoxygenases, inhibition of sEH, oxylipins, and PPARγ in the modulation of CRH in isolated mouse hearts is not known. Therefore, this study was designed to investigate the role of sEH, oxylipins, and PPARγ in the modulation of CRH using isolated WT mouse hearts. Our data demonstrated that: 1) Inhibition of sEH by t-AUCB enhanced CRH; 2) Treatment with t-AUCB significantly changed oxylipin profiles, including   Soluble Epoxide Hydrolase and Coronary Reactive Hyperemia an increased EET/DHET ratio, increased EpOME/DiHOME ratio, increased HODEs, decreased mid-chain HETEs, and decreased prostanoids; 3) Inhibition of CYP epoxygenases (by MS-PPOH) attenuated CRH; 4) The PPARγ antagonist T0070907 decreased CRH, and 5) the PPARγ agonist rosiglitazone increased CRH.
Inhibition of sEH by t-AUCB significantly enhanced CRH after brief ischemia compared to non-treated WT mice. The association between compromised CRH and some cardiovascular pathologies [26][27][28] confirms the significance of CRH. Further, the t-AUCB-enhanced CRH was accompanied by a 1.0-(at baseline) to 1.7-(post-ischemia) fold increase in the 14,15-EET/ DHET ratio, primarily driven by a decrease in 14,15-DHET. Although, a trend of increase in 14,15-EET level was observed in t-AUCB-treated vs. non-treated WT mice, the difference was not statistically significant. This could be attributed to several factors including the acute use of t-AUCB for a short period, the short half-life of EETs compared to DHETs, and the short duration (15 seconds) of ischemic insult. Another dihydroxy-EET metabolite (11, decreased in t-AUCB-treated mouse hearts, further confirming the efficacy of the sEH-inhibitor, t-AUCB. The specificity of t-AUCB as a sEH inhibitor was tested using sEH knockout (sEH -/-) mice, where no difference in CRH between the t-AUCB-treated sEH -/versus nontreated sEH -/mice was observed. The rapid hydration of EETs [37] may explain the challenge of detecting them. We were able to detect 14,15-EET and two diols (11,(12)(13)(14) in heart perfusate samples. As mentioned earlier, EETs are formed by the epoxidation of arachidonic acid (AA) by CYP epoxygenases and possess well-established beneficial cardiovascular effects [7,[38][39][40][41], including protection against ischemia/reperfusion injury [5], and vasodilation in many vascular beds such as the intestines [42], kidney preglomerular vasculature [43], and brain [44]. Inhibition of sEH, the main catabolic enzyme of EETs, has been widely used to increase the level of EETs and to investigate their effects [11]. Acute inhibition of sEH, through injection of a sEH inhibitor, lowered blood pressure in spontaneously hypertensive rats (SHR) [45], whereas chronic inhibition of sEH lowered angiotensin-II-induced hypertension [46]. Besides lowering blood pressure, sEH inhibition ameliorated renal damage in angiotensin-dependent, salt-sensitive hypertension rats [47]. In the current study, the brief ischemia had no effect on EET or DHET levels. This could be due to the short duration of ischemia (15 seconds). In contrast, DHETs (EETs metabolites) were elevated in the mouse heart perfusate in response to longer (20-min) ischemia [6]. Another interesting finding in the current study was that no significant difference was found in CRH between male and female mice in either t-AUCB-treated or non-treated groups (data not presented). Based on our data, the enhancement of CRH associated with sEH inhibition in WT mice may be partially due to an increase in the EET/DHET ratio.
Mid-chain (5-, 11-, 12-and 15-) HETE levels decreased in response to ischemia and to t-AUCB treatment at baseline and post-ischemia. Mid-chain HETEs are produced through allylic oxidation of AA by lipoxygenase (LOX) [48]. They were shown to have chemotaxis effects, change vascular tone, and induce the production of vascular endothelial growth factors [16][17][18][19]. Also, the increased formation of mid-chain HETEs was involved in cardiovascular dysfunction [49][50][51][52]. Unlike the vasodilatory effect of EETs in the kidneys [43], 12-HETE caused vasoconstriction in small renal arteries [53]. In human umbilical artery smooth muscle cells, 11-HETE inhibited vascular smooth muscle cell proliferation [54], whereas in the isolated Guinea Pig lungs, 5-and 15-HETEs produced vasoconstriction and edema [55]. Also, the generation of mid-chain HETEs is increased in essential hypertension [56] suggesting that they could be involved in its pathogenesis. These reports point to opposite effects of EETs and HETEs in vascular biology. Maayah et al. reported that mid-chain HETEs blocked the synthesis Fig 12. A schematic diagram comparing the oxylipin changes observed in response to a brief ischemia and their possible impact on coronary reactive hyperemia (CRH) between WT and t-AUCBtreated WT mice. Treatment with t-AUCB enhanced CRH possibly through increased EET/DHET ratio, increased 13-HODE, increased EpOME/DiHOME ratio, decreased mid-chain HETEs, and PPARγ activation. Soluble Epoxide Hydrolase and Coronary Reactive Hyperemia of EETs and increased their conversion to DHETs in RL-14 cells [57]. Moreover, although sEH is not directly involved in the generation or breakdown of mid-chain HETEs, sEH was found to be essential for mid-chain HETE-mediated induction of cellular hypertrophy [57]. Therefore, not only do EETs and mid-chain HETEs have opposite effects, they seem to affect the level of each other. In contrast to our findings, Li et al. reported that the plasma levels of 5-, 11-, 12-, and 15-HETE were not changed by sEH-inhibition in WT mice [11]. However, their results were based on plasma samples, whereas ours were based on isolated heart perfusate samples and demonstrated significant decrease in the level of the same HETEs in response to sEH inhibition. Therefore, in this study, it is possible that the decrease in mid-chain HETEs induced by sEHi (t-AUCB) has also contributed to blocking the enhanced conversion of EETs to DHETS by HETEs. This in turn would contribute to the enhanced CRH through an increase in EET/DHET ratio. Based on these results, the decrease in mid-chain HETEs in both t-AUCB-treated and non-treated mice in response to ischemia suggests that these metabolites are down regulated in response to ischemic episodes. This is an interesting finding because HETEs were the only oxylipins analyzed in this paper whose levels were affected by the brief ischemia. Also, the further decrease of these HETEs by t-AUCB-treatment may have played a role in the enhanced CRH observed in t-AUCB-treated hearts.
Linoleic acid (LA) epoxidation metabolites, such as 9,10-, and 12,13-EpOMEs, and their corresponding sEH-metabolized DiHOMEs were detected in the heart perfusate samples. As expected, inhibition of sEH by t-AUCB was further confirmed by an increased EpOME/ DiHOME ratio. This resulted from increased 9,10-,12,13-EpOMEs (albeit not statistically significant), and decreased 9,10-, 12,13-DiHOMEs in t-AUCB-treated versus non-treated mice. The physiological significance of EpOMEs and DiHOMEs remains poorly understood, and the evidence is somewhat contradictory, with a few studies suggesting toxic effects, while others indicating beneficial effects of these bioactive metabolites. EpOMEs and DiHOMEs were reported to increase oxidative stress in vascular endothelial cells [58]; DiHOMEs were toxic to renal proximal tubular cells [59]; and intravenously injected 9,10-EpOME had cardiodepressive effects in dogs [60]. The main caveat of these studies, which reported that EpOMEs and DiHOMEs have toxic effects, is that very high concentrations (100-500 μM) were used [48]. Contrary to these reports, lower, more physiological concentrations of EpOMEs and DiHOMEs did not have toxic effects and rather had beneficial effects [61]. Mitchell et al reported that LA and its oxidative metabolites (EpOMEs and DiHOMEs) did not have toxic effects during acute exposure in Langendorff-perfused rat hearts [61]. Pretreatment with 12,13-EpOME protected primary cultures of rabbit renal proximal tubular cells against hypoxia/reoxygenation injury [62]. Also, the increased EpOME/DiHOME ratio, as induced by sEH inhibition using AUDA, improved renal recovery in response to ischemia/reperfusion injury in C57BL/6 mice [14]. Based on these findings and on the described biologic role of LA epoxides, the increase in EpOME/DiHOME ratio in t-AUCB-treated mouse heart perfusates may have contributed to enhancing CRH.
Linoleic acid (LA) is also metabolized through hydroxylation by CYP epoxygenases to form hydroxyl-LA metabolites known as hydroxyoctadecadienoic acids (HODEs) [48]. The detected two HODE isomers, 9-, and 13-HODE, were increased in t-AUCB-treated WT. Like EpOMEs, the physiologic functions of HODEs are still being investigated [48]. 13-HODE is suggested to have an anti-inflammatory role in inflammatory diseases through its effect as a PPARγ-agonist [63][64][65][66][67]. Also, 13-HODE, through increasing prostacyclin (PGI 2 ) biosynthesis, was involved in splenic and coronary artery relaxation in smooth muscle cells in Mongrel dogs [15]. 9-HODE, unlike 13-HODE, was described as pro-inflammatory in an experimental wound-healing model in rats [68,69]. In contrast to 13-HODE, which can be produced from LA by the action of 12/15-LOX, 9-HODE production by LOX has not been reported in humans; rather, mouse 8-LOX metabolizes LA to 9-HODE [68]. In contrast to our data, Luria et al. reported no change in 9-and 13-HODE in sEH -/compared to sEH +/+ [10]. This discrepancy may be explained by the difference in the source of the metabolites: we used heart perfusate samples, whereas Luria et al [10] used urinary samples. Also, the reported metabolic and functional effects of sEH -/ -(genetic deletion of sEH) and pharmacologic inhibition of sEH are not necessarily the same [11]. Therefore, and based on our results, the increase in 13-HODE in t-AUCB-treated mouse heart perfusates might have played a role in enhancing CRH, whereas the role of 9-HODE is not yet known based on its, thus far, described biologic functions.
Treatment of WT mice hearts with t-AUCB significantly decreased prostanoid levels, including PGs and TxB 2 . PG-G 2 and PG-H 2 , AA metabolites formed by COX isoforms (1 and 2), are converted to the 4 main bioactive PGs (PG-D 2 , PG-E 2 , PG-I 2 , PG-F 2α ) and thromboxanes (TxA 2 and TxB 2 ) [70,71]. Most PGs have pro-inflammatory effects. For example, PG-E 2 augments arterial dilation and increases microvascular permeability [71]. However, PG-E 2 was found to have an anti-inflammatory role as well by up-regulating cAMP and inducing secretion of the anti-inflammatory IL-10 [72]. Similarly, PG-D 2 attenuated inflammation in experimental models of pleuritis and colitis [71]. 6-keto-PG-F 1α , the non-active hydrolysis product of prostacyclin (PG-I 2 ), and TxB 2 , the inactive degradation product of TxA 2 [71], were also decreased by t-AUCB. PG-F 2α is elevated in patients with chronic inflammatory diseases [71]. In support of our findings, sEH-inhibition by AUDA-BE reversed lipopolysaccharide (LPS)induced increase in PG-D 2 , PG-E 2 , 6-keto-PG-F 1α , and TxB 2 [13]. Also, sEH-inhibition by AUDA-BE, through increasing the concentrations of EETs, inhibited NF-κB translocation [12]. Combination of sEH-inhibition by AUDA-BE and COX-inhibition had an additive effect and further decreased PG-D 2 and PG-E 2 [12]. Therefore, the associated decrease in the measured PGs could be attributed to sEH-inhibition by t-AUCB. However, the impact of these changes on the t-AUCB-enhanced CRH is not yet clear.
As mentioned earlier, EETs are generated from AA by cytochrome P450 (CYP) epoxygenase enzymes, primarily CYP2C and CYP2J. We targeted the CYP epoxygenase-EET pathway using pharmacologic inhibition of CYP-epoxygenases to further support our findings of the involvement of CYP-epoxygenases in modulating CRH. sEH-inhibition increases the level of EETs by blocking their breakdown; we investigated the effect of blocking the synthesis of EETs on CRH using a CYP epoxygenases-inhibitor. Evidently, inhibiting CYP epoxygenases would negatively impact EET synthesis and, therefore, their effects. P450 enzymes are highest in coronary arteries and in small arterioles [2]. MS-PPOH was used as an EET-synthesis inhibitor that blocked the role of EETs as mediators of insulin-mediated augmentation of skeletal muscle perfusion [73]. Our data indicate that MS-PPOH significantly decreased CRH after a brief ischemia. These findings suggest that inhibition of CYP epoxygenases by MS-PPOH decreased CRH possibly due to inhibition of EETs synthesis. Further studies to investigate the impact of MS-PPOH on oxylipin profiles are needed to understand the metabolic changes associated with the inhibition of CYP epoxygenases.
This study also showed that CRH was decreased by T0070907, a PPARγ-antagonist, whereas rosiglitazone, a PPARγ-agonist, increased CRH. Moreover, T0070907 attenuated t-AUCB-enhanced CRH in WT mouse hearts. Studies indicated that EET effects could be mediated by PPARγ receptors [20][21][22][23]. Liu et al. suggested that selective sEH inhibition potentiates the anti-inflammatory effect in endothelial cells by increasing the retention of EETs so that PPARγ activation is prolonged [20]. Our results suggest that PPARγ receptors are involved in modulating CRH. Also, the attenuated t-AUCB-enhanced CRH in WT mice (by T0070907) suggests that PPARγ may mediate CRH downstream of CYP epoxygenases-EET pathway.
In summary, the findings of this study suggest that the CYP epoxygenases-EET pathway may have a role in mediating CRH possibly through PPARγ activation, as noted by sEH-inhibition, which enhanced CRH, and CYP epoxygenases-inhibition, which decreased CRH. Also, sEH-inhibition is accompanied by changes in AA oxylipins (increased EET/DHET ratio, decreased mid-chain HETEs, and decreased prostanoids) and LA oxylipins (increased EpOME/DiHOME ratio, and increased HODEs), which might have collectively accounted for the observed enhancement of CRH. Therefore, we conclude that sEH-inhibition enhances, whereas CYP epoxygenases-inhibition attenuates CRH, PPARγ activation mediates CRH downstream of the CYP epoxygenases-EET pathway, and the changes in oxylipin profiles associated with sEH-inhibition collectively account for the enhanced CRH.