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Am J Physiol Cell Physiol 294: C1407-C1418, 2008. First published April 2, 2008; doi:10.1152/ajpcell.00384.2007
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VASCULAR BIOLOGY

Asymmetric dimethylarginine inhibits HSP90 activity in pulmonary arterial endothelial cells: role of mitochondrial dysfunction

Neetu Sud,1,* Sandra M. Wells,2,* Shruti Sharma,1 Dean A. Wiseman,1 Jason Wilham,2 and Stephen M. Black1

1Vascular Biology Center, Medical College of Georgia, Augusta, Georgia; and 2Department of Biomedical and Pharmaceutical Sciences, The University of Montana, Missoula, Montana

Submitted 24 August 2007 ; accepted in final form 16 March 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Increased asymmetric dimethylarginine (ADMA) levels have been implicated in the pathogenesis of a number of conditions affecting the cardiovascular system. However, the mechanism(s) by which ADMA exerts its effect has not been adequately elucidated. Thus the purpose of this study was to determine the effect of increased ADMA on nitric oxide (NO) signaling and to begin to elucidate the mechanism by which ADMA acts. Our initial data demonstrated that ADMA increased NO synthase (NOS) uncoupling in both recombinant human endothelial NO synthase (eNOS) and pulmonary arterial endothelial cells (PAEC). Furthermore, we found that this endothelial NOS (eNOS) uncoupling increased 3-nitrotyrosine levels preferentially in the mitochondria of PAEC due to a redistribution of eNOS from the plasma membrane to the mitochondria. This increase in nitration in the mitochondria was found to induce mitochondrial dysfunction as determined by increased mitochondrial-derived reactive oxygen species and decreased generation of ATP. Finally, we found that the decrease in ATP resulted in a reduction in the chaperone activity of HSP90 resulting in a decrease in its interaction with eNOS. In conclusion increased levels of ADMA causes mitochondrial dysfunction and a loss of heat shock protein-90 chaperone activity secondary to an uncoupling of eNOS. Mitochondrial dysfunction may be an understudied component of the endothelial dysfunction associated with various cardiovascular disease states.

peroxynitrite; oxidative stress; mitochondria; protein-protein interactions

THERE IS INCREASING histological and physiological evidence that endothelial injury and the resulting aberration in the balance of its regulatory mechanisms play a major role in the development of vascular dysfunction. Endothelial dysfunction is a hallmark of many diseases of the vasculature, including hypertension (both pulmonary and systemic), atherosclerosis, and diabetes (25, 26). The most important consequence of endothelial dysfunction is the observed decrease in the ability of the endothelium to mediate vasodilation (61). Although the mechanisms involved in the development of endothelial dysfunction remain incompletely understood, recent studies have suggested that impaired vasodilation in a variety of cardiovascular diseases is linked to the inhibition of nitric oxide (NO) generation by the amino acid asymmetric dimethylarginine (ADMA) (4, 6–8, 59). ADMA is an endogenous competitive inhibitor of NO synthase (NOS) (4, 6–8, 59). ADMA is constantly produced during normal protein turnover in many tissues, including vascular endothelial cells, and is derived from hydrolysis of methylated proteins (4, 6–8, 59). It appears that both synthesis and degradation of ADMA is regulated in an active manner, and that loss of regulation of either of these pathways may result in elevated levels of ADMA (4, 6–8, 59). Increased ADMA levels have been implicated in the pathogenesis of a variety of conditions affecting the cardiovascular system (4, 6–8, 59) and, more recently, also in pulmonary hypertension (2, 37). In addition, several prospective and cross-sectional studies have identified ADMA as a marker of increased cardiovascular risk (4, 6–8, 59).

It has been previously demonstrated that endothelial NOS (eNOS) can interact with a 90-kDa heat shock protein (HSP90). HSP90 is a member of a molecular chaperone family of proteins that act to modulate protein functions within the cell. HSP90 appears to increase eNOS activity by facilitating the calmodulin-induced displacement of caveolin 1 from eNOS (24). This effect can be inhibited with the HSP90 inhibitor geldamycin (21). Several studies show that disruption of HSP90-eNOS interactions attenuates NO production (12, 21, 24, 33, 55) and leads to eNOS uncoupling (46, 47). HSP90 is ATP dependent and the ATPase site of the chaperone is responsible for the autophosphorylation required to enable HSP90 to interact with client proteins (53, 54), suggesting that mitochondrial dysfunction could decrease HSP90 chaperone activity through decreases in cellular ATP levels. Thus the purpose of this study was to investigate the effect of ADMA on NO and superoxide signaling in pulmonary arterial endothelial cells (PAEC) and to determine whether increases in the cellular levels of ADMA leads to the development of mitochondrial dysfunction, and if so, whether this is also associated with a decrease in ATP levels and the disruption of in eNOS-HSP90 interactions.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Purification of recombinant human eNOS. The poly-His-pCWeNOS vector (a gift from Paul Ortiz de Montellano, UCSF) was transformed in to the protease-deficient Escherichia coli strain BL21 (DE3) pLysS (Novagen). Cells were grown in Luria broth with 1% glycerol containing 200 µg/ml ampicillin and 40 µg/ml chloramphenicol. Cultures were grown at 28°C until an OD600 of 0.8 was reached. Approximately 1 h before, the heme precursor {delta}-aminolevulinic acid (0.5 mM final concentration) was added. Cells were then induced by adding isopropyl-β-D-thiogalactopyranoside (0.8 mM final concentration); 0.5 mM ATP and 3 µM riboflavin were also added and the cells were then grown at 22°C for an additional 48 h in dark. Cells were then harvested by centrifugation (15 min at 4,000 g at 4°C). The cell pellet was resuspended in lysis buffer [40 mM N-(2-hydroxyethyl) piperazine-N-(3-propane sulfonic acid) (EPPS), pH 7.6, containing 1 mg/ml lysozyme, 150 mM NaCl, 0.5 mM L-arginine, 4 µM BH4, 2 µM flavin adenine dinucleotide (FAD), and 10% glycerol] and protease inhibitor cocktail (Sigma) were added according to manufacturer's recommendation. The bacterial suspension was incubated with mild shaking at 4°C for 30 min to ensure complete cell lysis. Cells were broken by sonication using three 25-s pulses followed by three cycles of freezing and thawing. Cell debris was removed by centrifugation at 30,000 g for 30 min at 4°C. The supernatant was then applied to a Ni-NTA His-Bind Superflow (Novagen) column pre-equilibrated with buffer A (40 mM EPPS, pH 7.6, containing 150 mM NaCl, 10% glycerol, and 0.5 mM L-arginine). The column was washed with 5 bed volumes of buffer A followed by buffer B (buffer A with 25 mM imidazole). The bound protein was then eluted with buffer C (buffer A + 200 mM imidazole). The heme-containing fractions were pooled and concentrated using centriprep-100 YM-10 (Millipore). The concentrated protein were dialyzed against three changes of buffer A containing 4 µM BH4 and 1 mM DTT. The protein were further purified by using a 2'5'-ADP-sepharose column equilibrated with 40 mM Tris buffer, pH 7.6, containing 1 mM L-arginine, 3 mM DTT, 4 µM BH4, 4 µM FAD, 10% glycerol, and 150 mM NaCl (buffer D) and washed with buffer D containing 400 mM NaCl to prevent nonspecific binding. eNOS was then eluted with buffer E (buffer D with 5 mM 2'-AMP). The heme-containing fractions were pooled, concentrated, and dialyzed at 4°C against buffer D containing 1 mM DTT, 4 µM BH4, 4 µM FAD, and 10% glycerol and stored at –80°C until used. The DTT, BH4, and FAD were removed by repeated buffer exchange using centricon before use.

Cell culture. Primary cultures of ovine fetal pulmonary artery endothelial cells (PAEC) were isolated as described previously (18). Cells were maintained in DMEM containing phenol red supplemented with 10% fetal calf serum (Hyclone, Logan, UT), antibiotics, and antimycotics (MediaTech, Herndon, VA) at 37°C in a humidified atmosphere with 5% CO2-95% air. Cells were utilized between passages 3 and 10, seeded at ~50% confluence, and utilized when fully confluent.

The Y+ transporter carries both ADMA and L-arginine into the cell. The L-arginine levels in our culture medium are 84 mg/l (~500 µM). Thus the dose of ADMA used must be able to effectively compete for uptake with L-arginine. For this reason we chose to use 100 µM of ADMA. Our initial data indicated that 100 µM ADMA rapidly raises intracellular concentrations from ~5 µM to ~20 µM within 1 h (4-fold, see Fig. 2A).


Figure 2
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  		Fig. 2. ADMA induces eNOS uncoupling in pulmonary arterial endothelial cells (PAEC). PAEC were exposed or not exposed to ADMA (100 µM, 1 h), and then cellular ADMA levels were determined (A). In addition, the effect of increased ADMA on cellular superoxide levels were quantified using dihydroethidium (DHE) and fluorescent microscopy (B) or by ESR (C). The effect of ADMA on NO generation was also determined (D). The increased superoxide generation induced by ADMA is blocked by the addition of the NOS inhibitor ethylisothiourea (ETU, 100 µM), excess L-arginine (1 mM), or PEG-SOD (100 units). NO generation is also decreased, indicating ADMA is uncoupling eNOS. n = 4–6. Values are means ± SE. *P < 0.05 vs. Control; {dagger}P < 0.05 vs. ADMA alone.

 
Measurement of ADMA. ADMA was analyzed by high-performance liquid chromatography (HPLC) using modifications of a previously described method. Briefly, the crude fraction of cell lysates was isolated using a solid-phase extraction column, and subsequently, ADMA was separated using precolumn derivatization with {sigma}-phthaldialdehyde (OPA) reagent (4.5 mg/ml in borate buffer, pH 8.5, containing 3.3 µl/ml β-mercaptoethanol) before injection. HPLC was performed using an Amersham Biosciences AKTA purifier system (GE Healthcare, Piscataway, NJ) with a Nucleosil Phenyl reverse-phase column (4.6 x 250 mm; Supelco, Bellefonte, PA) equipped with a Jasco FP-2020 fluorescence detector (Jasco, Tokyo, Japan). ADMA levels were quantified by fluorescence detection at 450 nm (emission) and 340 nm (excitation). Mobile phase A was composed of 95% potassium phosphate (50 mM, pH 6.6) and 5% methanol, and mobile phase B was composed of 100% methanol. ADMA was separated using a pregradient wash of 25% mobile phase B (flow rate 0.8 ml/min), followed by a linear increase in mobile phase B concentration from 20% to 25% over 7 min, followed by a constant flow at 25% for 10 min, and another linear increase from 25% to 27% mobile phase B over 5 min followed by constant flow at 27% mobile phase B for another 7 min. Retention time for ADMA was ~28 min. ADMA concentrations were calculated using standards and an internal homoarginine standard. The detection limit of the assay was 0.1 µmol/l.

HSP90 immunoprecipitation and Western blot analysis. PAEC were exposed or not exposed to ADMA (100 µM) and then solubilized with a lysis buffer containing 1% Triton X-100, 20 mM Tris, pH 7.4, 100 mM NaCl, 1 mM EDTA, 1% sodium deoxycholate, 0.1% SDS, and protease inhibitor cocktail (Pierce). Insoluble proteins were precipitated by centrifugation at 13,000 rpm for 10 min at 4°C, and the supernatants were then incubated overnight with the anti-HSP90 antibody (2 µg) at 4°C followed by incubation in Protein G plus protein A agarose (Calbiochem) for 2 h. The immune complexes were precipitated by centrifugation, washed three times with lysis buffer, boiled in SDS sample buffer, and subjected to SDS-PAGE on 4–12% polyacrylamide gels and transferred to a nitrocellulose membrane (Bio-Rad). The membranes were blocked with 5% nonfat dry milk in Tris-buffered saline containing 0.1% Tween (TBST). The primary antibody eNOS (1:500, BD Transduction Laboratories) was then added and incubated for 1 h at room temperature. The membrane was then washed three times with TBST (10 min) and then incubated with a secondary antibody coupled to horseradish peroxidase. The membrane was then washed three times with TBST as described above. Reactive bands were visualized using the SuperSignal West Femto Maximum Sensitivity Substrate Kit (Pierce, Rockford, IL) and Kodak 440CF image station (Kodak, New Haven, CT). The intensity of the reactive bands was quantified using the Kodak 1D software. The efficiency of each immunoprecipitation was normalized by reprobing the membrane with the immunoprecipitating antibody (HSP90).

DHE fluorescence analysis. PAEC were seeded onto 96-well plates (Costar) and allowed to adhere for at least 18 h. Cells were then washed in PBS and incubated in serum-free DMEM in the presence or absence of ADMA (100 µM, 1 h). Dihydroethidium (DHE; 5 µM; Molecular Probes) was added to the media 15 min before the end of the experiment. Oxidation of DHE was observed after excitation at 518 nm and emission at 605 nm as described below.

Plasma membrane isolation. Plasma membrane was isolated from ADMA-treated and untreated cells using Pierce Mem-PER Eukaryotic Membrane Protein Extraction Reagent Kit according to manufacturer's protocol. Briefly, 5 x 106 cells were pelleted by centrifugation and then lyzed. The supernatant obtained by centrifugation at 10,000 g for 3 min was incubated at room temperature for 20 min. The tubes were centrifuged at 10,000 g for 2 min, and the hydrophobic phase (bottom layer) was carefully separated from hydrophilic phase (top layer). The hydrophobic phase contains the plasma membrane fraction.

Mitochondria isolation. After ADMA treatment for 1 h, the mitochondria were isolated using Pierce Mitochondria isolation kit using manufacturer's protocol. Briefly, 2 x 107 were pelleted by centrifugation at 850 g for 2 min, and Mitochondria Isolation Reagent A was added to the pellet. Cells were vortexed for 5 s and then incubated on ice for 2 min. Mitochondria Isolation Reagent B was added and then followed by vortexing for 5 s. Tubes were incubated on ice for 5 min and vortexing being done every 1 min. Mitochondria Isolation Reagent C was added, and the tubes were inverted several times to mix. The supernatant obtained by centrifugation at 700 g for 10 min at 4°C was transferred to a new tube and centrifuged at 3,000 g for 15 min. Mitochondria Isolation Reagent C was added to the pellet and centrifuged at 12,000 g for 5 min. The pellet contains the mitochondrial fraction. To determine the effect of eNOS inhibition on ADMA-induced eNOS translocation, cells were treated for 30 min with ethylisothiourea (ETU, 100 µM) before addition of ADMA. To detect how L-arginine effects ADMA-mediated eNOS translocation, L-arginine (500 µM) was added along with ADMA. To determine the effect of SOD on ADMA-stimulated eNOS translocation, cells were pretreated with 100 U/ml PEG-SOD (30 min) before ADMA addition.

Detection of mitochondrial superoxide levels. Mitochondrial superoxide production was measured using MitoSOX Red mitochondrial superoxide indicator (Molecular Probes), a fluorogenic dye for selective detection of superoxide in the mitochondria of live cells. MitoSOX Red reagent is live-cell permeant and is rapidly and selectively targeted to the mitochondria. Once in the mitochondria, MitoSOX Red reagent is oxidized by superoxide and exhibits bright red fluorescence upon binding to nucleic acids. Briefly, PAEC were treated with ADMA (100 µM, 30 min), and cells were washed with fresh media and then incubated in media containing MitoSOX Red (2 µM) and ADMA (100 µM) for a further 30 min at 37°C in dark conditions. Cells were washed with fresh serum-free media and imaged using fluorescence microscopy as described below using an excitation of 510 nm and an emission at 580 nm.

Fluorescence microscopy. A PC-based imaging system consisting of the following components: an Olympus IX51 microscope equipped with a CCD camera (Hamamatsu Photonics, Hamamatsu City, Japan) was used for acquisition of fluorescent images. Fluorescent-stained cells were observed with the appropriate excitation and emission, and the average fluorescent intensities (to correct for differences in cell number) were quantified using ImagePro Plus v.5.0 imaging software (Media Cybernetics, Silver Spring, MD).

EPR spectroscopy and spin trapping. To detect superoxide generation in intact cells, EPR measurements were performed as described previously (65). After an overnight serum starvation of the cells, 20 µl of spin-trap stock solution consisting of 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine·HCl (CMH, Alexis Biochemicals, San Diego, CA), 20 µM in DPBS + 25 µM desferrioxamine (Calbiochem, La Jolla, CA), and 5 µM diethyldithiocarbamate (Alexis Biochemicals, Lausen, Switzerland) + 2 µl DMSO were added to each well before shear stimulation. Adherent cells were trypsinized and pelleted at 500 g after a 45-min incubation at 37°C postshear to allow entrapment of superoxide by the spin trap. Cell pellet was washed and suspended in a final volume of 35 µl DPBS (containing desferrioxamine and diethyldithiocarbamate), loaded into a 50-µl capillary tube, and analyzed with a MiniScope MS200 EPR (Magnettech, Berlin, Germany) at a microwave power of 40 mW, modulation amplitude of 3,000 mG, and modulation frequency of 100 kHz. EPR spectra were analyzed measured for amplitude using ANALYSIS software (version 2.02; Magnettech), and experimental groups were compared using statistical analysis described below. In the experiments examining the role of ADMA in modulating superoxide production, the cells were preincubated with the ADMA (100 µM) for 30 min before the addition of the CMH. To determine the effect of eNOS inhibition on ADMA-induced superoxide production, cells were treated for 30 min with ETU (100 µM) before addition of ADMA. To detect how L-arginine effects ADMA-mediated superoxide generation, L-arginine (500 µM) was added along with ADMA. To determine the effect of SOD on ADMA-stimulated superoxide generation, cells were pretreated with 100 U/ml PEG-SOD (30 min) before ADMA addition. EPR measurements were performed as described above.

Effect of ADMA on NOx and superoxide generation by eNOS in vitro. The in vitro reaction was conducted in 50 µl of buffer containing 50 mM HEPES (pH 7.4), 1 mM NADPH, 100 µM L-arginine, 1 mM Ca2+, 10 µg/ml calmodulin, 4 µM tetrahydrobiopterin, and 1 µg eNOS. eNOS-dependent superoxide generation in vitro was performed in 50 µl of reaction mix containing 50 mM HEPES (pH 7.4), 1 mM NADPH, 1 mM Ca2+, 10 µg/ml calmodulin, 4 µM tetrahydrobiopterin, and 1 µg eNOS and CMH hydrochloride. In the experiments determining the effect of ADMA, eNOS and HSP90 were incubated for 15 min before measurements were made. The effect of ADMA on superoxide generation as then determined as described above. The effect of ADMA on NO generation was determined using an NO-sensitive electrode with a 2-mm diameter tip (ISO-NOP sensor, WPI) connected to an NO meter (ISO-NO Mark II, WPI) as described previously (65).

Effect of ADMA on SODactivity. Endothelial cells from fetal lamb were treated with ADMA for 1 h, and SOD activity in treated and untreated cells was measured using Cayman Chemical Superoxide Dismutase Assay Kit according to manufacturer's instructions. The method utilizes a tetrazolium salt to quantify superoxide radicals generated by xanthine oxidase and hypoxanthine. Briefly, cells were lysed in 20 mM HEPES buffer (pH 7.2) containing 1 mM EGTA, 210 mM mannitol, and 70 mM sucrose and then centrifuged at 1500 g for 5 min at 4°C, and the supernatant was collected for SOD assay. The samples and standards were added in duplicate to a sample plate provided with the kit. Reactions were initiated by adding xanthine oxidase to all wells, and the samples were incubated on a shaker at room temperature for 20 min. The absorbance of each standard and sample was read at 450 nm by the use of an absorbance microplate reader. SOD activity was calculated from the linear regression of the standard curve by substituting the linearized rate for each sample. One unit was defined as the amount of enzyme needed to exhibit 50% dismutation of the O2bullet– radical.

Dot-blot analysis for 3-nitrotyrosine. Mitochondrial fractions (50 µg) prepared from PAEC exposed to ADMA (100 µM, 0–8 h) were blotted onto nitrocellulose membranes using a Bio Dot apparatus (Bio-Rad, Hercules, CA). The membrane was rinsed in 20 ml TBST and blocked with 20 ml 5% nonfat milk in TBST for 1 h, followed by an incubation with a 3-nitrotyrosine (3-NT) antibody (1:1,000, Calbiochem, San Diego, CA) at 4°C overnight. After three washes with TBST, the membrane was incubated with goat antimouse IgG horseradish peroxidase-conjugated secondary antibody (1:2,000, Pierce, Rockford, IL) for 1 h at room temperature. After washing was completed, the dots were visualized with chemiluminescence using a Kodak Digital Science Image Station (NEN) and analyzed using the KED-1 software. All captured and analyzed images were determined to be in the dynamic range of the system. The same membrane was then probed with β-actin to normalize for loading.

Detection of NOx in pulmonary arterial endothelial cells. NO generated by PAECs was measured using an NO-sensitive electrode with a 2-mm diameter tip (ISO-NOP sensor, WPI) connected to an NO meter (ISO-NO Mark II, WPI) as described previously (33).

Cellular ATP levels. Assay for ATP was done using the firefly luciferin-luciferase method using a commercially available kit (Invitrogen). ATP is consumed and light is emitted when firefly luciferase catalyzes the oxidation of luciferin. The amount of light emitted during the reaction is proportional to the availability of ATP. Luminescence was measured using a Fluoroscan Ascent FL luminometer (Thermo Electron).

Statistical analysis. Statistical calculations were performed using the GraphPad Prism V. 4.01 software. The means ± SE were calculated for all samples, and significance was determined by either by the unpaired t-test or ANOVA. A value of P < 0.05 was considered significant.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effect of ADMA on superoxide and NO production by eNOS in vitro. To investigate the effect of ADMA on eNOS uncoupling, we initially utilized recombinant human eNOS. Our data indicate that superoxide production from eNOS was significantly enhanced (Fig. 1A, P < 0.05 vs. no ADMA) in the presence of ADMA, whereas the NO produced was significantly diminished (Fig. 1B, P < 0.05 vs. no ADMA).


Figure 1
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  		Fig. 1. Asymmetric dimethylarginine (ADMA) uncouples purified recombinant human endothelial nitric oxide (NO) synthase (eNOS). Purified human eNOS (7 pmol) was incubated with 5 µM BH4 and 0.1 mM L-arginine in the presence and absence of 0.1 mM ADMA. Electron spin resonance (ESR) was used to determine eNOS superoxide generation over a 20-min incubation period (A). Simultaneously, NO generation was determined using an NO sensor (B). ADMA significantly increases superoxide generation and decreases NO generation from eNOS. n = 6. Values are means ± SE. *P < 0.05 vs. no ADMA.

 
Effect of ADMA on superoxide generation in pulmonary arterial endothelial cells. To begin to determine the significance of increased ADMA levels on superoxide generation in PAEC, we initially determined the effect of adding exogenous ADMA to intracellular ADMA concentrations. Our data indicate that the addition of ADMA (100 µM, 1 h) to PAEC resulted in a significant (4.3-fold) increase in intracellular ADMA levels (Fig. 2A, P < 0.05 vs. Control). We next examined the effect of ADMA on NOS-dependent superoxide production utilizing the oxidation to dihydroethidium to ethidium. Superoxide generation was found to be significantly increased in ADMA-exposed PAEC (Fig. 2B, P < 0.05 vs. control). This ADMA-mediated increase in superoxide formation was blocked in the presence of the NOS inhibitor ETU, indicating that ADMA also induces eNOS uncoupling in PAEC (Fig. 2B, P < 0.05 vs. ADMA alone). We subsequently confirmed this ADMA-mediated increase in superoxide generation using EPR analysis (Fig. 2C). Furthermore, this increase in NOS-derived superoxide was associated with a decrease in NO generation (Fig. 2D).

To confirm that the ADMA-mediated increase in superoxide was due to uncoupling of eNOS, cells were pretreated with the nonspecific NOS inhibitor ETU. ETU caused a significant decrease in ADMA-treated cells (Fig. 2C). Furthermore, we found that the addition of excess L-arginine (which competes with ADMA for the Y+-transporter) or the superoxide scavenger PEG-SOD both significantly reduced the ADMA-mediated increase in superoxide (Fig. 2C). In addition, we found that ADMA did not alter SOD activity in the cell, indicating that the increase in superoxide was not due to a loss of SOD activity (Fig. 3).


Figure 3
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  		Fig. 3. ADMA does not alter superoxide dismutase (SOD) activity in PAEC. PAEC were exposed or not exposed to ADMA (100 µM, 1 h), and then cellular SOD activity was determined. ADMA exposure does not alter SOD activity. Values are expressed as mean units of activity per microgram of protein ± SE. N = 3.

 
ADMA increases 3-NT levels in pulmonary arterial endothelial cells. Because ADMA appeared to be causing uncoupling of eNOS, we then determined whether this induced 3-NT levels in PAEC (as a surrogate marker for peroxynitrite generation). Utilizing dot-blot analysis, we found that 3-NT levels were significantly increased in whole cell extracts of PAEC exposed to ADMA (Fig. 4A, P < 0.05 vs. Control). Similar increases in 3-NT levels were also observed in isolated mitochondria from ADMA-exposed PAEC, although the duration of the increase in 3-NT levels was longer (Fig. 4B, P < 0.05 vs. Control).


Figure 4
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  		Fig. 4. ADMA exposure increases peroxynitrite levels in PAEC. Cells were exposed or not exposed to ADMA (100 µM) for 0–24 h, and whole cell extracts (A) or mitochondrial fractions (B) were prepared and peroxynitrite exposure was estimated using dot-blot analysis to measure 3-nitrotyrosine (3-NT) protein levels. 3-NT levels are increased as early as 0.5 h of exposure in the whole cell extracts and are maintained for at least 6 h in the mitochondrial fraction. N = 6. Values are means ± SE. *P < 0.05 vs. Control.

 
ADMA stimulates eNOS redistribution to the mitochondrion. To further investigate the mechanism for the potentiated effect of ADMA on protein nitration within the mitochondria, we isolated plasma membrane and mitochondria from control and ADMA-treated PAEC and subjected them to Western blot analysis. Our data indicate that ADMA stimulated a redistribution of eNOS from the plasma membrane (Fig. 5, A and B, P < 0.05 vs. Control) to the mitochondria (Fig. 5, C and D, P < 0.05 vs. Control).


Figure 5
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  		Fig. 5. ADMA induces the redistribution of eNOS in PAEC. PAEC were treated with ADMA (100 µM, 1 h). Plasma membrane (A and B) and mitochondrial fractions (C and D) were then prepared and protein extracts subjected to Western blot analysis. A representative image is shown for each cell fraction. N = 3. Densitomeric values are means ± SE. *P < 0.05 vs. Control.

 
ADMA-mediated mitochondrial redistribution is peroxynitrite dependent. To further investigate the mechanism for the redistribution of eNOS from the plasma membrane to the mitochondria, we evaluated the role of the peroxynitrite produced by the ADMA-mediated uncoupling of eNOS. Our data indicated that the NOS inhibitor ETU caused a significant decrease in the ADMA-mediated eNOS mitochondrial translocation cells (Fig. 6, A and B). Furthermore, we found that the addition of excess L-arginine (which competes with ADMA for the Y+-transporter) or the superoxide scavenger PEG-SOD also both significantly reduced the ADMA-mediated eNOS mitochondrial translocation (Fig. 6, A and B). Furthermore, the peroxynitrite generator 3-morpholino-sydnonimine alone is able to stimulate eNOS translocation to the mitochondria (Fig. 6, C and D), whereas the peroxynitrite scavenger MnTymPyp reduces ADMA-mediated eNOS mitochondrial translocation (Fig. 6, C and D). Together, these data suggest a role for peroxynitrite in regulating the mitochondrial translocation of eNOS.


Figure 6
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  		Fig. 6. ADMA-mediated redistribution of eNOS in PAEC is peroxynitrite dependent. PAEC were treated with ADMA (100 µM, 1 h) in the presence and absence of the NOS inhibitor ETU (100 µM), L-arginine (1 mM), or PEG-SOD (100 units). Mitochondrial fractions were then prepared and protein extracts subjected to Western blot analysis. A representative image is shown (A). Densitomeric values are means ± SE (B). In addition, cells were treated for 1 h with the peroxynitrite generator 3-morpholino-sydnonimine (10 µM) or ADMA (100 µM) with or without the peroxynitrite scavenger MnTymPyp (25 µM). A representative image is shown (C). Densitomeric values are means ± SE (D). N = 3. *P < 0.05 vs. Control; {dagger}P < 0.05 vs. ADMA alone.

 
ADMA induces mitochondrial oxidative stress and increases UCP-2 expression in pulmonary arterial endothelial cells. To determine whether the increase in nitrated proteins in the mitochondria of ADMA-exposed cells was altering mitochondrial function, we utilized MitoSOX that we have previously shown to be a good marker of oxidative stress in the mitochondria (64, 65). We found that oxidative stress within the mitochondria was significantly increased in ADMA-exposed cells (Fig. 7, A and B, P < 0.05 vs. Control). Furthermore, this effect appears to be a specific effect of ADMA as the metabolites of ADMA, L-citrulline and dimethylamine did not increase oxidative stress in the mitochondria of PAEC (Fig. 7C). In addition, our data indicate that ADMA-exposed cells have a significantly increased expression of the mitochondrial uncoupling protein UCP-2 (Fig. 7, D and E, P < 0.05 vs. Control) a further indication of mitochondrial stress induced by ADMA. To further evaluate the role of ADMA in inducing mitochondrial dysfunction, we evaluated the dose- and time-dependent effects of ADMA on mitochondrial oxidative stress. We evaluated the effect of ADMA in both L-arginine containing and L-arginine-free medium. Our data indicate that in L-arginine-free medium exposure to a dose as low as 2 µM, ADMA significantly increases mitoSox fluorescence (Fig. 8A). In addition, we found that over a 24-h exposure time, there was a time-dependent increase in MitoSox Fluorescence induced by 100 µM ADMA in both L-arginine-containing and -free medium but only with L-arginine free when 2 µM was used (Fig. 8B). In all cases there was significantly more oxidative stress in the cells exposed to ADMA in L-arginine-free conditions (Fig. 8B). Finally, we found that even in L-arginine-free conditions, L-citrulline and dimethylamine did not increase oxidative stress in the mitochondria of PAEC (Fig. 8C).


Figure 7
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  		Fig. 7. ADMA induces mitochondrial oxidative stress in PAEC. Cells were treated with ADMA (100 µM), L-citrulline (100 µM), or dimethyamine (100 µM) for 90 min. MitoSOX red mitochondrial superoxide indicator (Molecular Probes) was added to a final concentration of 5 µm, and the cells were incubated for another 30 min at 37°C for a total exposure time of 2 h. The ADMA-exposed cells were washed with PBS, and images were captured and the fluorescent intensity of the images (A) was quantified (B). Similarly, the fluorescent intensities of the L-citrulline- and dimethylamine-exposed cells were determined (C). L-Citrulline or and dimethylamine did not increase mitosox fluorescence. Images of 10 random fields were quantified for each sample. In addition, whole cell protein extracts (10 µg) were separated on 4–20% SDS-PAGE and electrophoretically transferred to polyvinylidene difluoride membranes. The membranes were then subjected to Western blot analysis using a rabbit polyclonal UCP2 antibody. Bands were visualized with chemiluminescence using a Kodak Digital Science Image Station. UCP2 protein levels were then normalized to β-actin protein. A representative blot is shown (D). Densitomeric values were then obtained (E). N = 4–8 for each group. Values are means ± SE. *P < 0.05 vs. Control.

 

Figure 8
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  		Fig. 8. Increased mitochondrial oxidative stress is modulated by L-arginine. Cells were incubated with increasing concentrations of ADMA (0–100 µM), L-citrulline (100 µM), or dimethyamine (100 µM) in the presence or absence of L-arginine for increasing times (0–24 h). MitoSOX red mitochondrial superoxide indicator (Molecular Probes) was added to a final concentration of 5 µm 30 min before analysis. The ADMA-exposed cells were then washed with PBS, and images of 10 random fields were captured and the fluorescent intensity of each was quantified. There is a dose-dependent increase in MitoSox fluorescence in the absence of L-arginine (A). Furthermore, the presence of L-arginine reduces the ADMA-mediated increase in MitoSox fluorescence (B). Treating cells with L-citrulline (100 µM) or dimethyamine (100 µM) for 2 h does not increase MitoSox fluorescence even in the absence of L-arginine (C). N = 3 for each group. Values are means ± SE. *P < 0.05 vs. Control.

 
ADMA decreases ATP levels in PAEC. To determine whether the ADMA-induced increase in mitochondrial oxidative stress was altering mitochondrial function, we next investigated the effect of ADMA on cellular ATP levels. Our data indicate that ADMA caused a significant reduction in cellular ATP levels that was maintained for at least 8 h (Fig. 9, P < 0.05 vs. control).


Figure 9
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  		Fig. 9. ADMA exposure decreases cellular ATP levels in PAEC. Cells were exposed or not exposed to either ADMA (100 µM) or the mitochondrial inhibitor 2,4-DNP (50 µM, as a positive control for mitochondrial dysfunction) for 0–8 h, and cellular ATP levels were determined using a luciferase-based ATP measurement kit. Both ADMA and 2,4-DNP significantly reduce cellular ADMA levels. N = 6. Values are means ± SE. *P < 0.05 vs. Control.

 
ADMA disrupts the interaction of HSP90 with eNOS. As mentioned previously the chaperone activity of HSP90 is ATP dependent. Thus we examined the effect of the ADMA-mediated decrease in cellular ATP levels on HSP90 activity by examining eNOS-HSP90 interactions. To accomplish this we utilized immunoprecipitation. Western blot analysis for eNOS on the immunoprecipitated HSP90 complex revealed that ADMA exposure led to significant decrease in the association of HSP90 with eNOS (Fig. 10, P < 0.05 vs. Control) in PAEC.


Figure 10
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  		Fig. 10. ADMA disrupts heat shock protein-90 (HSP90) activity in pulmonary arterial endothelial cells. PAEC were treated with ADMA (100 µM, 1 h). Cells were then washed with PBS, and lysates were prepared with modified RIPA buffer. Immunoprecipitation was performed by using an antibody to HSP90. Protein extracts were separated on 4–20% SDS-PAGE and electrophoretically transferred to polyvinylidene difluoride membranes. The membranes were then probed with the appropriate antibody to eNOS. A representative images is shown (A). Blots were also stripped and reprobed for HSP90 to normalize for the immunoprecipitation efficiency. Densitomeric values were obtained for total eNOS (B) protein. N = 5. Values are means ± SE. *P < 0.05 vs. untreated cells.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
The important findings of this study are as follows: 1) ADMA stimulates the redistribution of eNOS from the plasma membrane to the mitochondrion. 2) ADMA uncouples eNOS in both in vitro and PAEC. 3) The increased generation of peroxynitrite is required for eNOS redistribution to the mitochondrion. 4) The uncoupled eNOS localized to the mitochondrion results in an increase in nitrated proteins in the mitochondria. 5) These nitration events leads to mitochondrial dysfunction and reduced cellular ATP levels. 6) The reduction in ATP leads to a decrease in the chaperone activity of HSP90;, that 7) decreases the interaction of HSP90 with eNOS further reducing NO signaling.

A number of studies suggest that impaired vasodilation in a variety of cardiovascular diseases is linked to the inhibition of NO generation by the amino acid ADMA (4, 6–8, 59). ADMA is an endogenous competitive inhibitor of NOS (4, 6–8, 59). ADMA is continuously produced in the course of normal protein turnover in tissues, including vascular endothelial cells, and is derived from hydrolysis of methylated proteins (4, 6–8, 59). Elevated plasma levels of ADMA have been reported in heart failure (19), atherosclerosis (38), diabetes mellitus (34), and hypertension(23). Furthermore, Gorenflo et al. (23) found ADMA plasma levels in patients with severe pulmonary hypertension were significantly increased (median, 0.55 µM/l; range, 0.25–0.79 µM/l) compared with those levels in control patients (median, 0.21 µM/l; range, 0.08–0.27 µM/l). This increase may account for the endothelial vasodilator dysfunction observed in this condition. Also, Boger and colleagues (9) previously demonstrated that cultured human endothelial cells produce superoxide (as determined by increased oxidation of DHE) in the presence of ADMA. This led to the hypothesis that ADMA may interrupt the NO-producing activity of NOS and "uncouple" the enzyme, which results in a "switch" of the enzymatic activity from NO to superoxide (9). The data we present here demonstrate that ADMA can uncouple eNOS both in vitro, using recombinant purified human eNOS, and in cultured PAEC. The direct uncoupling effect of ADMA on eNOS is interesting because a recent study found that ADMA does not uncouple purified neuronal NOS (13). These differing effects of ADMA on NOS isoforms with regard to NOS uncoupling suggest that ADMA may exert effects that are isoform dependent, and future studies to examine the effect of ADMA on inducible NOS coupling will also be required. Furthermore, our data indicate that the increased eNOS uncoupling in ADMA-exposed cells results in an increase in nitrated protein both in whole cell extracts and in isolated mitochondria. However, the effect on the mitochondria appears to be more sustained (6 h vs. 1 h). This is likely due to the fact that ADMA stimulates the redistribution of eNOS from the plasma membrane to the mitochondrion. eNOS redistribution to the outer mitochondrial membrane has been shown previously in both human umbilical vein endothelial cells and in human embryonic kidney cells transiently transfected with an eNOS expression construct (20). The key sequence required for mitochondrial localization has been localized to residues 628–632 (20). This sequence is located within the 45-amino acid autoinhibitory domain of eNOS that is an insertion in the flavin mononucleotide-containing domain and is thought to have an important role in regulating eNOS activity (51). 5 Further studies will be required to determine whether ADMA acts on these same residues to stimulate the redistribution of eNOS. Alternatively, it is also possible that the mitochondrion lacks the ability to remove nitrated proteins effectively including nitrated eNOS. Furthermore, our data indicate that the elevations in cellular and mitochondrial 3-NT decrease with time and that the cells are responding to the ADMA challenge. The mechanism by which occurs is not clear from our studies. However, one possible explanation is that as ADMA is actively degraded by DDAH, over time intracellular ADMA concentrations will tend to diminish as we added only one bolus dose of ADMA. Thus eNOS uncoupling will also tend to diminish, which would explain the recovery of the cells over time. Similarly, this could explain the apparent recovery of ATP levels in the cell. In addition, the mechanism by which ATP levels diminish is unclear. However, our recent in vivo studies in a lamb model of pulmonary hypertension secondary to increased pulmonary blood flow have shown that, indeed, our peroxynitrite-mediated nitration of the mitochondrial protein carnitine acetyl trasnferase (CrAT) may be important for the loss of mitochondrial function. Thus it is possible that the nitration mediated by ADMA leads to an inhibition of CrAT and subsequent decreases in ATP levels. Further studies will be required to test these possibilities.

With increasing knowledge of the role of ADMA in the pathogenesis of cardiovascular disease, ADMA is likely to become a goal for pharmacotherapeutic interventions (5). Several studies support the view that the ratio between L-arginine and ADMA is a key component in the regulation of eNOS activity, and elevated ADMA levels have been shown to antagonize endothelium-dependent vasodilation in humans (4, 6–8, 59), and the administration of L-arginine has been shown to improve endothelium-dependent vascular functions in subjects with high ADMA levels (10). ADMA can be metabolized via hydrolytic degradation to L-citrulline and dimethylamine by the enzyme DDAH (6). There are two currently identified isoforms of DDAH: DDAH1 and DDAH2 (11, 38). The endothelial cell predominant isoform is believed to be DDAH2 (31, 59) and the inhibition of DDAH activity results in vasoconstriction of vascular segments that can be reversed by L-arginine (35). The regulation of DDAH is not fully understood. However, in endothelial cells, oxidative stress induced by oxidized LDL or tumor necrosis factor-{alpha} has been shown to decrease DDAH activity (30), whereas Lin and colleagues (34) have shown that elevated glucose raises endothelial ADMA levels by inhibiting DDAH activity. Conversely, in an interesting study from Tran and co-workers (59), all-trans-Retinoic acid increased nitrite production by sEnd.1 cells without increasing eNOS expression. Rather, trans-Retinoic acid was shown to increase DDAH II gene expression and promoter activity with a subsequent reduction in ADMA levels. However, further studies will be required to determine whether altering the ADMA-to-L-arginine ratio with arginine supplementation or stimulating DDAH activity will reduce the mitochondrial dysfunction that we have found with elevated levels of ADMA.

Disruption of mitochondrial function is acknowledged as a critical event in a number of pathological conditions, including hypoxia-ischemic injuries (3), stroke (56), and diabetes (17, 39, 41, 44). There is also evidence for decreased mitochondrial function in aging-related neurodegenerative disorders (18, 22). In addition, a study in poultry has shown that lung mitochondrial dysfunction is present in broilers with pulmonary hypertension syndrome, and this was associated with oxidative stress as high dietary vitamin E attenuated the effect (29). Mitochondria are the source of superoxide anion radicals and H2O2 under physiological and pathological conditions. The increased production of reactive oxygen species from the mitochondria can be deleterious to the cell due to their ability to induce lipid peroxidation, protein oxidation, and DNA damage (49, 60). Indeed we, and others, have shown that reactive nitrogen species such as nitric oxide and peroxynitrite can lead to increased mitochondrial reactive oxygen species production (27, 43, 65). The mechanisms for these effects appear to involve the inhibition of mitochondrial complexes-I and -III (27, 43) as well as the disruption of zinc homeostasis (65). The data we present here indicate that mitochondrial dysfunction can also decrease HSP90 chaperone activity secondary to a loss of normal ATP generation. This reduction in HSP90 activity could have significant effects on cell function. With respect to endothelial biology and NO signaling, HSP90 is known to interact with a number of proteins required for efficient NO biosynthesis, including eNOS (24), soluble guanylate cyclase (42, 62), and possibly GTP cyclohydrolase (58). A loss of normal activity for any of these proteins would have a significant deleterious effect on NO signaling. However, the effects of a decrease in HSP90 activity would extend far beyond NO signaling. HSP90 is an essential chaperone for the proper folding of a large number of proteins. The importance of HSP90 function is demonstrated by the fact that in eukaryotes, the activity of HSP90 is essential for cell viability. The function of HSP90 is highly complex with recent studies only now beginning to identify signaling networks that are dependent of a functional HSP90. The complexity of these interactions have been demonstrated using the lower eukaryote Saccharomyces cerevisiae where HSP90 has been shown to interact directly or indirectly with more than 10% of all proteins in the cell. Thus any decreases in the activity of the HSP90 system could have dramatic effects on cell survival.

An oversupply of electrons in the mitochondrial transfer chain can also result in mitochondrial membrane hyperpolarization and the formation of reactive oxygen species (16, 32, 50, 63). These free radicals damage proteins and lipids and lead to dysfunction of mitochondria, the central mediators of programmed cell death (16, 32, 50, 63). Damaged mitochondria release proapoptotic factors that activate the cysteine protease family of caspases, which in turn propagate a death cascade (16, 32, 50, 63). Cellular energy and mitochondrial membrane function are regulated in part by the uncoupling proteins (UCPs) (14, 48). UCPs are mitochondria carrier proteins that dissipate the proton gradient of the inner mitochondria membrane. This uncoupling reaction can bypass the production of ATP by oxidative phosphorylation (14, 48). Increased expression of UCP isoforms is believed to be a marker of mitochondrial dysfunction (1, 28, 36, 40, 57). The different members of the UCP family have distinct tissue distributions (15, 45, 52). Tissue localization as well as regulation confers different roles for the family members. For example, UCP1 is strictly restricted to brown adipose tissue, and its principal function is believed to be the generation of body heat (15, 45, 52). UCP3 tissue distribution is predominantly limited to skeletal muscle, whereas UCP2 expression is widespread (15, 45, 52). However, it is still unclear the exact role played by UCP proteins in mitochondrial dysfunction. In addition, how their expression and activation are regulated has not been adequately resolved. Our data here link increased nitrosative stress in the mitochondria with increased UCP-2 expression, suggesting that mitochondrial dysfunction may be a signal for the cell to increase UCP-2 expression. Recent investigations (57) have demonstrated that UCP can also prevent mitochondrial damage during stroke, suggesting that increased expression of UCP may be an adaptive response for the cell to try and protect the mitochondria after an insult. However, further studies will be required to determine whether increases in UCP-2 expression are protective for the mitochondria in ADMA-challenged PAEC.

In conclusion, our data suggest that increased levels of ADMA can induce mitochondrial dysfunction in PAEC with a resulting decrease in cellular ATP levels and HSP90 chaperone activity. Furthermore, our data emphasize that mitochondrial dysfunction may be an understudied aspect of cardiovascular disorders, and that therapies aimed at maintaining mitochondrial function may have beneficial effects on endothelial function in these pathological states.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This research was supported in part by National Heart, Lung, and Blood Institute Grants HL-60190 (to S. M. Black), HL-67841 (to S. M. Black), HL-072123 (to S. M. Black), and HL-070061 (to S. M. Black), Grant 0550133Z from the American Heart Association, Pacific Mountain Affiliates (to S. M. Black), and by a grant from the Fondation Leducq (to S. M. Black).


   ACKNOWLEDGMENTS
 
The authors thank Dr. Jeff Fineman (University of California at San Diego) for critical reading of this manuscript and with help with the statistical analyses.


   FOOTNOTES
 

Address for reprint requests and other correspondence: S. M. Black, Vascular Biology Center: CB-3210B, Medical College of Georgia, 1459 Laney Walker Blvd., Augusta, GA 30912 (e-mail: sblack{at}mcg.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* N. Sud and S. M. Wells contributed equally to this work. Back


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