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(Hypertension. 1996;28:663-668.)
© 1996 American Heart Association, Inc.

Articles

Angiotensin II Type 2 Receptor Subtype Mediates Phospholipase A₂–Dependent Signaling in Rabbit Proximal Tubular Epithelial Cells

Leila S. Jacobs; Janice G. Douglas

the Departments of Medicine and of Physiology and Biophysics, Case Western Reserve University School of Medicine, and University Hospitals of Cleveland (Ohio).

	Abstract

Top Abstract Introduction Methods Results Discussion References

 
We investigated the ability of angiotensin II (Ang II) or the stable analogue [Sar¹]-Ang II to increase intracellular and extracellular free arachidonic acid in primary cultures of rabbit proximal tubular epithelial cells to better characterize the receptor subtype and orientation of phospholipase A₂ (PLA₂)–mediated signaling. Proximal tubular cells were labeled with [³H]arachidonic acid for 4 hours and then treated with Ang II or [Sar¹]-Ang II. Lipids were extracted from labeled cells, separated by thin-layer chromatography, and quantified by liquid scintillation counting. Ang II (10 µmol/L, 1 minute) stimulated an increase in intracellular free [³H]arachidonic acid from 21.0±2.0 to 32.2±2.8 disintegrations per minute/µg protein, an effect that was potentiated by EGTA. [Sar¹]-Ang II stimulated a time- and concentration-dependent increase in [³H]arachidonic acid release from labeled cells. Release of [³H]arachidonic acid was maximal at 10 µmol/L [Sar¹]-Ang II, with an EC₅₀ of approximately 3 µmol/L. Ang II receptor antagonists caused concentration-dependent inhibition of [Sar¹]-Ang II–stimulated [³H]arachidonic acid release with the following order of potency: CGP 42112=PD 123319>losartan. Furthermore, in proximal tubular epithelial cells grown on polyester membrane filters, the Ang II receptor that mediated arachidonic acid release was predominantly apical rather than basolateral. These observations are consistent with activation of a Ca²⁺-independent, apical PLA₂ isoform in epithelial cells through an Ang II type 2 receptor subtype.

Key Words: angiotensin II • arachidonic acids • kidney • lipids • losartan • receptors, angiotensin II • signal transduction

	Introduction

Top Abstract Introduction Methods Results Discussion References

 
Angiotensin II (Ang II) acts directly on proximal convoluted tubules to regulate sodium and water transport.¹ The signal transduction pathways through which Ang II exerts these and other functional effects have not been fully elucidated. Ang II–stimulated Na⁺-H⁺ antiport in rabbit renal brush border membrane vesicles has been suggested to involve PLA₂ activation mediated by a 42-kD G protein.² Concentration-dependent effects of luminal Ang II on proximal tubular fluid transport are associated with activation of brush border membrane PLA₂.³ PLA₂ catalyzes release of AA esterified in the sn-2 position of membrane phospholipids. AA can be metabolized by cyclooxygenase, lipoxygenase, or cytochrome P450 monooxygenase into a wide range of biologically active compounds. A cytochrome P450 AA epoxygenase has been purified from rabbit renal cortex,⁴ and rabbit cytochromes P450 2C1 and 2C2 have been identified as AA epoxygenases.⁵ Epoxyeicosatrienoic acids are endogenous constituents of rabbit kidney,⁶ and Ang II stimulates cytochrome P450 AA epoxygenase-dependent formation of epoxyeicosatrienoic acids in proximal tubules.⁷ Cytochrome P450–dependent AA metabolites exhibit diverse effects, including inhibition⁸ and stimulation⁹ of Na⁺,K⁺-ATPase, modulation of ion transport,¹⁰ activation of Na⁺-H⁺ exchange and mitogenesis,¹¹ regulation of vasoactivity^{12 13} and hemodynamic responses,¹⁴ and stimulation of increased [Ca²⁺]_i.¹⁵

Binding experiments have revealed the existence of at least three pharmacologically distinct angiotensin receptor subtypes in rabbit proximal tubular epithelial cells.¹⁶ Although many enzymes that metabolize AA are located intracellularly, most previous studies have focused on extracellular release of AA formed by PLA₂. In this study, our objective was to determine the cellular compartmentalization of PLA₂-generated AA and to identify the Ang II receptor subtype involved in PLA₂ activation in rabbit proximal tubular epithelial cells.

	Methods

Top Abstract Introduction Methods Results Discussion References

 
Materials
Ang II and [Sar¹]-Ang II were obtained from Peninsula Laboratories. CGP 42112 was obtained from CIBA-Geigy, Bachem Bioscience, or Bachem California. PD 123319 was donated by Parke-Davis. Losartan (DuP 753) was the generous gift of Dr Ronald D. Smith of DuPont-Merck. [³H]AA (180 to 240 Ci/mmol) and [methoxy-³H]inulin (233.5 mCi/g) were obtained from New England Nuclear Research Products. AA, fatty acid–free bovine serum albumin, EGTA, and pertussis toxin were obtained from Sigma Chemical Co. Phosphatidylcholine, phosphatidylserine, phosphatidylinositol, phosphatidylethanolamine, and phosphatidic acid were obtained from Matreya or Sigma. Chloroform, n-hexane, and isooctane were obtained from Burdick and Jackson. Methanol, ethyl acetate, and acetic acid were obtained from Fisher Scientific Co.

Cell Culture
Renal proximal tubular epithelial cells were isolated from male, barrier-reared Elite-New Zealand White rabbits (2 kg, Hazelton, Denver, Pa) as previously described.¹⁷ Procedures performed on rabbits were in accordance with the Institutional Animal Care and Use Committee guidelines. Rabbits were injected intravenously into an ear vein with 0.6 mL heparin sodium (20 000 U/mL) (Rugby Laboratories, Inc). After 10 minutes, rabbits were injected intravenously into an ear vein with 3 mL pentobarbital sodium (50 mg/mL) (Abbott Laboratories), and kidneys were immediately removed. Primary cultures of proximal tubular cells were passaged onto plastic 12-well cluster plates (Costar) in DMEM (GIBCO) and Ham's F-12 (GIBCO) (1:1, by volume) supplemented with 350 µg/mL L-glutamine, 100 U/mL penicillin, 100 µg/mL streptomycin, 600 µg/mL sodium bicarbonate, 0.5 µmol/L hydrocortisone, 5 µg/mL transferrin, 5 µg/mL insulin, and 5% fetal bovine serum (Hyclone Laboratories) and maintained in a humidified atmosphere of air supplemented with 5% CO₂. Proximal tubular cells (passage 1) were grown to confluence and used for experiments 3 days after plating. Cellular protein content was determined by the method of Lowry et al.¹⁸

For polarity experiments, proximal tubular cells were grown on polyester membrane filters (Transwell-clear chambers, 24-mm diameter, 0.4-µm pore size, Costar). Before experiments, we evaluated the integrity of representative monolayers by adding [³H]methoxyinulin to the apical medium and measuring the leak of [³H]methoxyinulin from the apical to the basolateral compartment by sampling the basolateral medium after incubation (1 hour, 37°C) and quantifying by liquid scintillation spectrometry. Monolayers excluded more than 95% of apically applied [³H]methoxyinulin from the basolateral compartment.

Labeling and Measurement of Phospholipids
For determination of the time course of [³H]AA incorporation into phospholipids, proximal tubular cells were labeled with [³H]AA (1 µCi/mL) in DMEM/F-12 containing 0.5% fetal bovine serum for 1, 2, 4, or 6 hours and washed three times with DMEM/F-12 containing 1 mg/mL bovine serum albumin for removal of unincorporated label; lipids were extracted by the method of Bligh and Dyer.¹⁹ The chloroform lipid extract was dried under vacuum centrifugation in a SpeedVac (Savant Instruments, Inc), resuspended in 50-µL chloroform, and applied to thin-layer chromatography plates (Whatman silica gel 150A). Phospholipid standards were also applied to the plates, both in the samples and in lanes to which samples were not added. For separation of phospholipids, thin-layer chromatography plates were developed in chloroform/methanol/acetic acid/water (75:50:10:6, by volume).²⁰ Lipids were visualized by placement of thin-layer chromatography plates into a chamber containing iodine. Radiolabeled phospholipids that comigrated with standards were scraped from the plate and quantified by liquid scintillation spectrometry.

Measurement of AA
For examination of the effect of Ang II or [Sar¹]-Ang II on intracellular and extracellular free [³H]AA, proximal tubular cells were labeled with [³H]AA (1 µCi/mL) in DMEM/F-12 containing 0.5% fetal bovine serum for 4 hours and washed three times with DMEM/F-12 containing 1 mg/mL bovine serum albumin, and Ang II or [Sar¹]-Ang II in DMEM/F-12 containing 1 mg/mL bovine serum albumin was added for the indicated time. For measurement of intracellular [³H]AA, incubations were terminated by placement of cells on ice and addition of ice-cold methanol. Cells were scraped from the plate and transferred to tubes containing chloroform. Lipids were extracted, visualized, and quantified as described above for phospholipids. Before scraping, thin-layer chromatography plates were scanned with a linear analyzer (Berthold). For separation of AA, thin-layer chromatography plates were developed in the organic (upper) phase of ethyl acetate/isooctane/acetic acid/water (45:25:10:50, by volume)²¹ or, for optimal resolution of AA and its metabolites, in the organic phase of n-hexane/ethyl acetate/isooctane/acetic acid/water (1:1:0.91:0.37:0.80, by volume).⁴ For measurement of extracellular [³H]AA, the extracellular medium was centrifuged (12 000 rpm, 2 minutes) for removal of nonadherent cells and quantified by liquid scintillation spectrometry.

	Results

Top Abstract Introduction Methods Results Discussion References

 
Incorporation of AA into Phospholipids
We designed the experiment shown in Fig 1 to determine the optimal labeling time for [³H]AA. The time course indicates that [³H]AA preferentially incorporates into phosphatidylcholine during 1 to 4 hours of labeling. Incorporation of [³H]AA into phosphatidylserine, phosphatidylinositol, phosphatidylethanolamine, and phosphatidic acid had a significantly slower onset but exceeded incorporation into phosphatidylcholine at 6 hours (Fig 1). The solvent system used for separation of phospholipids only partially resolves phosphatidylserine from phosphatidylinositol and phosphatidylethanolamine from phosphatidic acid,²⁰ such that values for phosphatidylserine/phosphatidylinositol and phosphatidylethanolamine/phosphatidic acid are presented together. Phosphatidylcholine is the most abundant phospholipid in kidney cortical tubules²² and is a major substrate for PLA₂. Moreover, a phospholipid pool undergoing rapid turnover may be more biologically active than an equilibrated phospholipid pool. Therefore, rabbit proximal tubular epithelial cells were labeled with [³H]AA for 4 hours, a time point at which AA incorporation into phosphatidylcholine was maximal.

View larger version (15K): [in this window] [in a new window]   Figure 1. Incorporation of [3H]AA into rabbit proximal tubular epithelial cell phospholipids as a function of time. Values are mean±SE (n=3, one experiment). PS indicates phosphatidylserine; PI, phosphatidylinositol; PE, phosphatidylethanolamine; PA, phosphatidic acid; and PC, phosphatidylcholine.

Intracellular Free AA
Ang II (10 µmol/L, 1 minute) stimulated an increase in intracellular free [³H]AA from 21.0±2.0 to 32.2±2.8 disintegrations per minute/µg protein. Extracellular EGTA (3 mmol/L, 1 minute) potentiated the effect of Ang II, increasing intracellular free [³H]AA from 22.4±0.9 to 40.0±2.9 dpm/µg protein (Fig 2). These data were confirmed by scanning thin-layer chromatography plates with a linear analyzer before scraping. Since the stable analogue [Sar¹]-Ang II stimulated a larger increase in intracellular and extracellular free [³H]AA than Ang II, we used it in subsequent experiments to clarify temporal and dose-response relationships.

View larger version (33K): [in this window] [in a new window]   Figure 2. Effect of Ang II (10 µmol/L, 1 minute), in the absence or presence of extracellular EGTA (3 mmol/L, 1 minute), on intracellular free [3H]AA in rabbit proximal tubular epithelial cells. Values are mean±SE (n=3, two experiments). Significance of Ang II–stimulated increase over control was determined by Student's unpaired t test (*P<.01, **P<.001).

Time Dependence of AA Release
The time course of [Sar¹]-Ang II (10 µmol/L)–stimulated [³H]AA release into the extracellular medium of labeled proximal tubular cells exhibited a rapid onset (1 minute) and a plateau at approximately 10 to 20 minutes (Fig 3). The maximal [Sar¹]-Ang II–stimulated increase (508% of control) in [³H]AA release occurred at 5 minutes.

View larger version (16K): [in this window] [in a new window]   Figure 3. Time dependence of 10 µmol/L [Sar1]-Ang II–stimulated [3H]AA release from rabbit proximal tubular epithelial cells. Values are mean±SE (n=3, one experiment).

Concentration Dependence of AA Release
[Sar¹]-Ang II stimulated a concentration-dependent increase in [³H]AA release (5 minutes) into the extracellular medium of labeled proximal tubular cells (Fig 4). Release of [³H]AA was maximal at 10 µmol/L, with an EC₅₀ of approximately 3 µmol/L.

View larger version (13K): [in this window] [in a new window]   Figure 4. Concentration dependence of [Sar1]-Ang II–stimulated [3H]AA release in 5 minutes from rabbit proximal tubular epithelial cells. Values are mean±SE (n=3, representative of five experiments).

Effect of Inhibitors on AA Release
We designed the experiment shown in Fig 5 to characterize the Ang II receptor subtype involved in Ang II–stimulated PLA₂ activity. [Sar¹]-Ang II (1 µmol/L) and inhibitor were added simultaneously for 5 minutes. The AT₂-selective antagonists CGP 42112 and PD 123319 produced concentration-dependent inhibition of [Sar¹]-Ang II–stimulated [³H]AA release from proximal tubular cells. The inability of PD 123319 to completely inhibit [Sar¹]-Ang II–stimulated [³H]AA release may be attributable to a partial agonistic effect of PD 123319 previously described in renal tissue. Indicative of their partial agonistic effects, low concentrations (1 to 10 µmol/L) of either CGP 42112 or PD 123319 potentiated the effect of [Sar¹]-Ang II on [³H]AA release. In contrast, the AT₁-selective antagonist losartan did not inhibit [Sar¹]-Ang II–stimulated [³H]AA release (Fig 5). Potentiation of [Sar¹]-Ang II–stimulated [³H]AA release by 100 µmol/L losartan suggests AT₁ receptor involvement. However, the potentiation was not significant (P=.08) when evaluated by ANOVA with Tukey's adjustment for multiple pairwise comparisons. Ang II antagonists inhibited [Sar¹]-Ang II–stimulated [³H]AA release with the following order of potency: CGP 42112=PD 123319>losartan, indicating that an AT₂ receptor subtype mediates PLA₂ activation.

View larger version (18K): [in this window] [in a new window]   Figure 5. Concentration-dependent inhibition by CGP 42112 (circles) and PD 123319 (small rectangles) of [Sar1]-Ang II–stimulated [3H]AA release from rabbit proximal tubular epithelial cells. Contrast the lack of inhibition by losartan (squares). [Sar1]-Ang II (1 µmol/L) and inhibitor were added simultaneously for 5 minutes. Values are mean±SE (n=3, representative of three experiments).

Polarity of AA Release
We designed the experiment shown in Fig 6 to determine the polarity of Ang II–stimulated PLA₂ activity and AA release. Apical addition of [Sar¹]-Ang II (10 µmol/L, 5 minutes) stimulated increases of 232.0% and 195.3% in apical and basolateral release of [³H]AA, respectively, into the extracellular medium of proximal tubular cells grown on polyester membrane filters that maintained negligible paracellular transport. In contrast, basolateral addition of [Sar¹]-Ang II (10 µmol/L, 5 minutes) stimulated significantly smaller increases of 136.8% and 129.3% in apical and basolateral release of [³H]AA, respectively (Fig 6).

View larger version (24K): [in this window] [in a new window]   Figure 6. Effect of apical versus basolateral addition of [Sar1]-Ang II (10 µmol/L, 5 minutes) on [3H]AA release from rabbit proximal tubular epithelial cells grown on polyester membrane filters. Apical control was defined as apical [3H]AA release (5 minutes) in the absence of [Sar1]-Ang II. Basolateral control was defined as basolateral [3H]AA release (5 minutes) in the absence of [Sar1]-Ang II. Apical and basolateral controls differed because [3H]AA release occurs across the entire apical surface, whereas [3H]AA release from the basolateral surface occurs only through the polyester membrane filter pores. Values are mean±SE (n=3, two experiments). Solid bars indicate apical release; shaded bars, basolateral release.

Effect of Pertussis Toxin
For determination of whether the AT₂ receptor is coupled to PLA₂ through a pertussis toxin–sensitive G protein, proximal tubular cells were treated with pertussis toxin (100 ng/mL, 18 hours) before they were labeled with [³H]AA. Values for extracellular release of [³H]AA were as follows: control=53.6±3.6 dpm/µg protein, pertussis toxin=56.3±1.2, [Sar¹]-Ang II (10 µmol/L, 5 minutes)=728.2±57.4, and [Sar¹]-Ang II (10 µmol/L, 5 minutes) plus pertussis toxin=700.4±34.1 (mean±SE, n=3, one experiment). The difference between [Sar¹]-Ang II and [Sar¹]-Ang II plus pertussis toxin was not significant (P=.7060, Student's unpaired t test). These data suggest that the AT₂ receptor is not linked to PLA₂ through a pertussis toxin–sensitive G protein.

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
The functional role of the AT₂ receptor remains uncertain. Current knowledge about it is summarized in a recent editorial.²³ AT₂ receptors mediate cGMP production in rat kidney.²⁴ They mediate inhibition of rat coronary endothelial cell proliferation through an unidentified signaling pathway.²⁵ The present study demonstrates an additional role for AT₂ receptors linked to PLA₂-mediated AA release. Previous studies from this laboratory demonstrated that AA and its cytochrome P450–dependent metabolites mediate Ang II–induced natriuresis in proximal tubular epithelium. However, the receptor subtype mediating this response was not determined. Furthermore, it is possible that this signaling pathway might mediate the antimitogenesis linked to the AT₂ receptor.

The reaction catalyzed by PLA₂ is the rate-limiting step in the formation of numerous biologically active metabolites derived from AA. [Sar¹]-Ang II has proved to be a useful agonist with which to study activation of PLA₂. [Sar¹]-Ang II is a stable analogue of Ang II that has sarcosine (N-methylglycine) substituted for aspartic acid in position 1.²⁶ [Sar¹]-Ang II and Ang II act on the same receptor,²⁶ but [Sar¹]-Ang II is more potent^{26 27} and has a higher equilibrium association constant than Ang II.²⁷ [Sar¹]-Ang II stimulated a time-dependent (Fig 3) and concentration-dependent (Fig 4) increase in [³H]AA release into the extracellular medium of rabbit proximal tubular epithelial cells.

Ang II stimulated an increase in intracellular free [³H]AA in proximal tubular cells that was potentiated by EGTA (Fig 2), suggesting that Ang II may activate a Ca²⁺-independent PLA₂ isoform distinct from the Ca²⁺-dependent PLA₂ frequently linked to vasoactive agonists. A cytosolic Ca²⁺-independent, plasmalogen-selective PLA₂ activity has been associated with hypoxic injury to rabbit proximal tubules.²⁸ AA may affect cellular function directly or be metabolized into epoxyeicosatrienoic acids or other biologically active eicosanoids. Intracellular AA directly activates K⁺ channels in neonatal rat cardiac atrial cells²⁹ and toad stomach smooth muscle cells³⁰ and activates Ca²⁺ channels in guinea pig ventricular myocytes.³¹ AA also stimulates an increase in [Ca²⁺]_i that is not completely inhibited by ketoconazole,¹⁵ modulates Ca²⁺ release from the sarcoplasmic reticulum in rat ventricular myocytes,³² inhibits myosin light chain phosphatase,³³ and mediates PLA₂-dependent effects on endosome fusion and endocytosis.³⁴ Thus, intracellular AA release represents a physiologically relevant observation as the AA is available for regulation of ion channels or further metabolism by intracellular enzymes.

The fact that Ang II activates PLA₂ in epithelial cells in the high nanomolar to micromolar range appears appropriate for apical PLA₂ regulation because the K_i of the Ang II receptor in rabbit brush border membranes (the predominant site mediating Ang II–linked signaling described herein) is 782 nmol/L.¹⁶ Furthermore, luminal Ang II concentrations are approximately 10 to 40 nmol/L,^{35 36} which is 1000-fold higher than plasma concentrations. Of note is the fact that this is the first report of a complete dose-response relationship of Ang II–stimulated PLA₂-mediated AA release.

Ang II receptor antagonists inhibited [Sar¹]-Ang II–stimulated [³H]AA release with an order of potency of CGP 42112=PD 123319>losartan (Fig 5), suggesting that activation of PLA₂ occurs through an AT₂ subtype. Ang II receptors with the same order of potency for Ang II antagonists have been identified in brush border but not basolateral membranes from rabbit proximal tubular epithelial cells.¹⁶ Moreover, Ang II stimulates PLA₂ through an AT₂ receptor subtype in neonatal rat ventricular myocytes,³⁷ and binding sites sensitive to PD 123177,³⁸ PD 123319,^{39 40} and PD 121981⁴¹ that probably represent AT₂ subtypes have been detected in rat kidney^{38 39 40} and rhesus monkey kidney cortex.⁴¹ The AT₂ receptor subtype in rabbit proximal tubular epithelial cells described herein may be analogous to previously described renal receptors in other species.^{38 39 40 41}

The antagonist action of CGP 42112 in the present study is in contrast to its reported agonist actions in some other systems. The agonist versus antagonist properties of CGP 42112 depend on tissue or cell type and the response being monitored. For example, the AT₂ receptor–selective peptide CGP 42112 produces significant stimulation of neuronal potassium current after superfusion.⁴²

Ang II–induced proximal tubular fluid transport and natriuresis are mediated by apical PLA₂ activity.^{3 43} Apical addition of [Sar¹]-Ang II stimulated increases of 232.0% and 195.3% in apical and basolateral release of [³H]AA, respectively, into the extracellular medium of polarized proximal tubular cells grown on polyester membrane filters (Fig 6). Basolateral addition of [Sar¹]-Ang II stimulated significantly smaller increases of 136.8% and 129.3% in apical and basolateral release of [³H]AA, respectively (Fig 6). These data indicate that Ang II–stimulated PLA₂ activity is preferentially localized to the apical surface and release of AA occurs from both the apical and basolateral surfaces. In contrast, basolateral Ang II receptors have been linked to adenylyl cyclase.⁴⁴

It remains to be determined whether renal Ang II receptors are linked to PLA₂ by a G protein. The ß- and -subunits of transducin stimulate PLA₂ in bovine rod outer segments,⁴⁵ and ß- and -subunits activate muscarinic potassium channels through stimulation of PLA₂ in neonatal rat atrial membranes.⁴⁶ Despite the fact that some pharmacological studies reported an absence of G protein coupling to AT₂ receptors, cloning studies revealed a G protein–coupled receptor consensus sequence in AT₂ receptors.^{47 48} This laboratory has reported that GTPS inhibits ¹²⁵I–[Sar¹]-Ang II binding to basolateral membranes in a concentration-dependent manner but does not inhibit binding to brush border membranes.¹⁶ Hence, pharmacological studies suggest that the apical Ang II receptor is not G protein coupled. Of interest is the fact that pertussis toxin does not inhibit [Sar¹]-Ang II–stimulated AA release in a dose and time shown to be effective in these cells. Although these data suggest that apical Ang II receptors are not linked to PLA₂ by a pertussis toxin–sensitive G protein, coupling to a pertussis toxin–insensitive G protein is not precluded.

A model of Ang II–stimulated PLA₂ activity in rabbit proximal tubular epithelial cells is illustrated in Fig 7. In summary, apical addition of Ang II activates PLA₂ independent of extracellular Ca²⁺ and increases intracellular AA through an AT₂ receptor subtype in rabbit proximal tubular epithelial cells.

View larger version (19K): [in this window] [in a new window]   Figure 7. Schematic model of Ang II–stimulated PLA2 activity in rabbit proximal tubular epithelial cells. EETs indicates epoxyeicosatrienoic acids.

	Selected Abbreviations and Acronyms

 

AA = arachidonic acid Ang II = angiotensin II AT1, AT2 = angiotensin type 1, type 2 DMEM = Dulbecco's modified Eagle's medium PLA2 = phospholipase A2

	Acknowledgments

 
This research was supported by Hypertension Program Project Grant HL-41618 from the National Heart, Lung, and Blood Institute to J.G. Douglas. L.S. Jacobs was supported by Hypertension Training Grant HL-07714 from the National Heart, Lung, and Blood Institute. We thank Judith Preston for excellence in cell isolation and culture. We thank Dr George R. Dubyak for providing a SpeedVac, Dr Vincent M. Monnier for providing a thin-layer chromatography plate linear analyzer, and Dr Steven J. Bowlin for performing the ANOVA.

	Footnotes

 
Reprint requests to Janice G. Douglas, MD, Division of Hypertension, Case Western Reserve University, School of Medicine W165, 10900 Euclid Ave, Cleveland, OH 44106-4982.

Received March 25, 1996; first decision April 26, 1996; first decision June 20, 1996;

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