Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2012 Apr 5.
Published in final edited form as: Free Radic Biol Med. 2006 Apr 26;41(4):568–578. doi: 10.1016/j.freeradbiomed.2006.04.010

17β-Estradiol Reverses Shear Stress-Mediated LDL Modifications

Juliana Hwang +,&, Mahsa Rouhanizadeh *,&, Ryan T Hamilton +, Tiantian C Lin *, Jason P Eiserich **, Howard N Hodis +,#, Tzung K Hsiai *,+
PMCID: PMC3320656  NIHMSID: NIHMS365913  PMID: 16863990

Abstract

Within arterial bifurcations or branching points, oscillatory shear stress (OSS) induces oxidative stress mainly via the NADPH oxidase system. It is unknown whether 17β-estradiol (E2) can regulate OSS-mediated low density lipoprotein (LDL) modifications. Bovine aortic endothelial cells (BAECs) were pre-treated with E2 at 5 nmol/L, followed by exposure to OSS (0 ± 3.0 dynes/cm2sec and 60 cycles/min) in a flow system. E2 decreased OSS-mediated NADPH oxidase mRNA expression, and E2-mediated ·NO production was mitigated by the ·NO synthase inhibitor L-NAME. The rates of O2−· production in response to OSS increased steadily as determined by superoxide dismutase-inhibited ferricytochrome c reduction; however, pre-treatment with E2 decreased OSS-mediated O2−· production (n=4, P<0.05). In the presence of native LDL (50 μg/mL), E2 also significantly reversed OSS-mediated LDL oxidation as determined by high performance liquid chromatography. In the presence of O2−· donor, xanthine oxidase (XO), E2 further reversed XO-induced LDL lipid peroxidation (n=3, P<0.001). Mass spectra were acquired in the m/z 400–1800 range, revealing XO-mediated LDL protein nitration involving tyrosine 2535 in the α-2 domains, whereas pre-treatment with E2 reversed this observation, consistent with the changes in nitrotyrosine intensities by the dot blots. E2 plays an indirect antioxidative role. In addition to up-regulation of eNOS and down-regulation of Nox4 expression, E2 influences LDL modifications via lipid peroxidation and protein nitration.

Keywords: 17β-Estradiol, Shear stress, LDL lipid oxidation and nitration, NADPH oxidase, Superoxide anion, Nitric oxide

Introduction

The cardiovascular protective effects of estrogen in the regulation of endothelial function has become important in pathological states in which hemodynamics, specifically shear stress, influences the bioavailability of ·NO [1]. Shear stress is a tangential force that exerts on the vascular endothelial cells by virtue of blood viscosity and velocity gradients [2]. Estrogen relaxes arteries, increases blood flow, and minimizes thickening of the carotid artery where oscillatory shear stress (OSS) develops in both humans and animals [3]. OSS is characterized as bidirectional net zero forward flow that commonly occurs within vascular bifurcations and branching points. OSS is a potent stimulus for proatherogenic expression of adhesion molecules and chemokines [4, 5] and enhancing vascular NADPH oxidase activity [6, 7]. Our previous studies demonstrated that OSS increased production of superoxide anion (O2−·) [7, 8] and oxidative modification of low density lipoprotein (LDL) [5].

It has long been suspected that the level of circulating 17β-estradiol (E2) contributes to gender-specific cardio-protection [9–11]. E2 up-regulates endothelial ·NO synthase (eNOS) gene expression leading to an increase in ·NO production [12] and restores the regulation of wall shear stress in arterioles of spontaneously hypertensive male rats [13]. Further, estrogen has antioxidant effects that oppose oxidation of LDL [14, 15]. Vascular endothelial cells (EC) become hyperpermeable in response to oxidized LDL [16] and hyperlipoproteinemia [17] that favor intimal uptake of LDL and local oxidative degradation of trapped LDL [18–20]. Thus, oxidized LDL plays an important role in inflammatory responses and oxidative stress at the atherosclerosis prone regions [21, 22]. However, the mechanism(s) by which 17β-estradiol (E2) mediates the effects of pro-atherogenic OSS within arterial bifurcations or curvatures remain unclear.

In the present study, we used a dynamic device generating precise flow patterns to investigate the antioxidant role of E2 that occurs in human carotid arteries [2, 23, 24]. OSS-induced oxidative modification of LDL was accompanied by increased O2−· and decreased nitric oxide (·NO) production. In contrast, E2 reversed OSS-mediated LDL oxidation that was accompanied by decreased O2−· and increased NO formation. Pre-treatment of E2 further attenuated xanthine oxidase-mediated LDL lipid peroxidation and protein nitration. Mass spectra analyses in the m/z 400–2000 range revealed LDL apo B 100 lipoprotein modifications involving tyrosine 2535 in the α-2 domains, consistent with XO-mediated nitrotyrosine by dot blots. These findings suggest that E2 at a physiological concentration (5 nmol/L) suppresses OSS-mediated O2−· production and reverses LDL post-translational protein modifications.

Methods and Materials

Isolation of LDL from Human Plasma

Venous blood was obtained from fasting adult human volunteers under institutional review board approval at the University of Southern California Atherosclerosis Research Unit. Plasma was pooled and immediately separated by centrifugation at 1500 g for 10 minutes at 4°C. LDL isolation was performed by density gradient centrifugation using a SW-41 rotor (Beckman, Fullerton, CA) utilizing a Beckman L-80 XP type centrifuge as described by Hodis et al [25]. LDL (δ=1.019–1.063) was then collected and washed with phosphate buffered saline (PBS) several times using a Millipore (Bedford, MA) centrifugal filtering device with a 30kDa cutoff. Concentrated LDL was then sterilized, stored at 4°C and used within 2–4 weeks. Protein concentrations were measured using Bio-Rad DC assay (Bio-Rad, Richmond, CA) with BSA used as a standard.

Experimental Protocol

Confluent bovine aortic endothelial cell (BAEC) monolayers were exposed to shear stress in a dynamic flow system (Fig. 1). The production of O2−· was measured by cytochrome c reduction rates and the expression of Nox4 and eNOS by quantitative real-time RT-PCR (described below). Native LDL at 50 μg/mL was added to the culture media maintained under the four conditions described below: (Control, E2, OSS and E2 + OSS) and aliquots of LDL isolated from the culture medium at 4 hours (and transferred to PBS) were subjected to anion exchange high performance liquid chromatography (HPLC) to analyze oxidized LDL, measured as LDL− [5, 26]. In some experiments, 100μM N(G)-nitro-L-arginine methyl ester (L-NAME) (Sigma, St. Louis, MO), an inhibitor of NOS, was added at 20 min prior to loading with native LDL.

Fig. 1.

Fig. 1

Open in a new tab

(a) A dynamic flow system: Confluent BAEC monolayers were seeded on the bottom of the dynamic flow channel which will be mounted on the inverted microscope while an array of MEMS sensors will be embedded on the upper wall. (b) Monocyte and or BAEC were captured and digitized. (c) A magnification of shear stress sensor array. (d) Real-time recording of pulsatile (PSS) vs. oscillatory shear stress (OSS). This newly designed physiologic flow model enabled us to implement the specific shear stress values with temporal variations. The effluents was used for biochemical analyses, specifically, analysis of NO2−/NO3−, cytochrome c reduction assay for superoxide (O2−·) and high performance liquid chromatography (HPLC) for LDL oxidation. The recovered BAEC were used for gene and protein expression.

Endothelial Cell Culture

Between passages 3 and 6, BAECs were seeded on Cell-Tak cell adhesive (Becton Dickson Labware, Bedford, MA) and Vitrogen (Cohesion, Palo Alto, RC 0701) coated glass slides (5 cm2) at 3 × 106 cells per slide. Cells were grown to confluent monolayers in high glucose (4.5 g/L) Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 15% heat inactivated fetal bovine serum (Hyclone), 100 U/mL penicillin-streptomycin (Irvine Scientific) and 0.05% amphotericin B (Gibco) for 48 hours in 5% CO2 at 37°C.

Dynamic Flow System

The physiological flow system was designed to generate well-defined temporal shear stress gradients and was mounted on an inverted microscope (Olympus FV-IX701) for real-time visualization of BAECs [27, 28] (Fig. 1). The system was capable of simulating both pulsatile and oscillatory flow conditions reported in the human arterial system [2, 23]. Micro Electro Mechanical Systems (MEMS) sensors were embedded in the channel to monitor real-time shear stress [29]. The circulating DMEM culture medium was maintained at 37 °C and pH 7.4 (Accumet AP) using 5% CO2.

Shear Stress Protocols

Confluent BAEC monolayers between passages 3 and 6 were plated on glass slides and incubated in phenol red free DMEM, 10% male human serum solution, 1% penicillin-streptomycin, and 0.05% fungizone. Cells were subjected to four conditions: (1) static condition, (2) pre-treatment with E2 at 5 nmol/L for 12 hours at static condition, (3) OSS at 60 cycles/min and 0 ± 3.0 dynes/cm2sec (time-averaged shear stress (τave) = 0 dynes/cm2) for 4 hours, and (4) pre-treatment with E2 for 12 hours, followed by OSS for 4 hours. After 4 hours, BAECs were collected for quantification of Nox4 and eNOS mRNA expression. Effluents were collected for cytochrome c reduction assay and quantitative measurements of NO2− and NO3−. Cells were also exposed to the above four conditions in the presence of 50 μg/ml of native LDL After 4 hours, the medium was aliquoted to assess oxidative modification of LDL as described below.

Separation of Native and Oxidatively Modified LDL Subspecies by HPLC

The technique used for separating LDL was similar to that described previously [8]. Analysis of oxidatively modified LDL or electronegative LDL (LDL−) was used as a measure of LDL modification according to methods previously described [25]. The total LDL and LDL subfractions were eluted through an anion exchange column (UNO-Q1, BioRad, Hercules, CA) at 1.0 mL/min and the effluent monitored at 280 nm. Native LDL (nLDL) and oxidatively modified LDL (or LDL− due to its electronegativity) [30] were isolated by a stepwise sodium chloride gradient. Area under the curve was measured and oxidatively modified LDL (LDL−) percentage was determined. Accordingly, the ratio of LDL− relative to total LDL was compared as a measure of LDL modification [5].

Quantitative Real-Time RT-PCR

After BAECs were exposed to the flow conditions, total RNA was isolated using the RNeasy kit (Qiagen). Real-time RT-PCR was performed according to the recommendations of the PE Applied Biosystems TaqMan PCR Core Reagent Kit [31]. RT and PCR were performed beginning with a manual ramp rate at 50°C for 2 minutes, 60°C for 30 minutes, and then 95°C for 5 minutes, followed by PCR for 50 cycles at 94°C for 20 seconds and annealing from 58 °C to 65 °C for 1 minute (MJ Research Opticon® System). CT is the threshold cycle number at which the initial amplification becomes detectable by fluorescence [32]. ΔRn normalizes fluorescence. TaqMan probes [33] were used for further specificity and sensitivity. The difference in CT values for various flow conditions vs. control was used to mathematically determine the relative difference in the level of Nox4 and eNOS mRNA expression [31]. For quantification of relative gene expression, the target sequence was normalized to the expressed housekeeping gene GAPDH.

Measurement of Nitrate/Nitrite (NO3−/NO2−)

Quantitative measurements of NO2− and NO3− were performed as an index of global nitric oxide (·NO) production essentially following methods described previously [34, 35]. The level of ·NO produced by BAEC subjected to static (no flow) or oscillatory shear stress was measured by the Griess reaction [36]. Briefly, aliquots of the cell culture medium were collected from different treatment conditions for NO2− and NO3− using the Griess reaction (Cayman Nitrite/Nitrate Colorimetric Assay Kit, MI, USA). Briefly, collected cell culture medium was incubated with nitrate reductase for 2 hours to convert NO3− to NO2− [37]. After incubation, Griess reagents were added to the medium and NO2− levels quantified spectrophotometrically at 540 nm. The amount of ·NO produced by E2 was verified by addition of N(G)-nitro-l-arginine methyl ester (L-NAME) [38] at 100μM, a dose at which optimal inhibition of ·NO production was demonstrated. The results were cross-checked with an analytical procedure based on acidic reduction of NO2− and NO3− to ·NO by vanadium (III) and purging of ·NO with helium into a stream of ozone and detected by an Antek 7020 chemiluminescence ·NO detector (Antek Instruments, Houston, TX). At room temperature, vanadium (III) only reduced NO2−, whereas NO3− and other redox forms of ·NO (such as S-nitrosothiols) were reduced only if the solution was heated to 90–100 °C, so that the total ·NO could be quantified. This allowed for determination of the relative levels of NO2− and NO3−, which might be indicative of differential oxidative metabolic routes of ·NO. For most analyses, 25–50 μl of extracellular fluids were utilized for this assay, and determinations made in duplicate. Quantification was performed by comparison with standard solutions of NO2− and NO3−.

Measurement of Superoxide (O2−·) Formation

The production of O2−· was measured by cytochrome c reduction rates as described previously [39, 40]. In each case, the medium contained 100μM acetylated-ferricytochrome c (Sigma Aldrich). Control samples were placed in a tissue culture dish in complete media with cytochrome c (100μM) and incubated at 37°C. Aliquots of culture medium (300μL) were collected at 0, 1, 2, 3, and 4 hours for absorbance measurements at 550nm (Beckman DU 640 Spectrophotometer). For the OSS conditions, BAEC monolayers were placed in the flow system. At 0, 1, 2, 3, and 4 hours, aliquots of medium bathing the BAECs in the flow apparatus were aspirated and mixed with acetylated-ferricytochrome c (1mM) and monitored for the absorbance at 550nm. The specificity for reduction by O2−· was established by parallel measurements in the presence of PEG-SOD (60 μg/mL). The corrected rates for SOD-inhibited cytochrome c reduction were plotted after computing O2−· formation using an extinction coefficient: E550=2.1 × 104 M−1cm−1 at an absorbance wavelength of 550 nm.

Xanthine Oxidase-Mediated LDL Modifications

BAECs at passage 5 were grown to confluent monolayers in high glucose (4.5 g/L) phenol red free Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% male human serum (Omega Scientific, CA), 1% L-glutamine-penicillin-streptomycin solution (Sigma Aldrich) in 5% CO2 at 37°C. BAECs were treated with E2 at 5nM/L for 24 hours, followed by exposure to LDL (50μg/mL) in the presence and absence of xanthine oxidase (20mU/mL) for 4 hours. The culture media were collected for the determination of modified LDL (LDL−) by HPLC and LDL protein nitration by dot blot analysis for nitrotyrosine, respectively. LDL lipid peroxides were measured using the leucomethylene blue assay using tert-butyl hydroperoxide as a standard. Measurements were made at 650 nm.

Dot Blot Analysis

Dot blots were performed by spotting 12 μg of LDL protein which were exposed to BAEC for 4 hours. Membranes were soaked in methanol and washed with TBS-T. Samples were then blocked in milk and incubated with a 1:3000 dilution of monoclonal nitro-tyrosine antibody (Upstate, Lake Placid, NY) overnight followed by incubation with HRP-conjugated goat anti-mouse secondary antibody at a dilution of 1:10000. Blots were detected using ECL chemiluminescence kit (Pierce, Rockford, IL).

Liquid Chromatography and Tandem Mass Spectrometry (LC/MS/MS) Analyses

  1. In-solution tryptic digest was carried in 50 mM ammonium carbonate buffer with 5mM CaCl2 using reductively methylated trypsin (Promega) to reduce self-hydrolysis. A 20:1 protein to enzyme ratio was used for the digestion reaction which was carried out at 37°C for at least three hours. Digestion products were prepared for LC/MS/MS analysis by drying down the reaction sample using an APD SpeedVac (ThermoSavant), resuspending evaporate in 5% acetonitrile/0.1 formic acid, and subjecting this sample to ZipTip purification using an elution buffer of 70% acetonitrile/0.1% formic acid. The ZipTip eluent was evaporated using the APD SpeedVac.

  2. Analysis of tryptic peptide sequence tags was performed by tandem MS. The digest samples were suspended in 10 μl 60% formic acid. Chromatographic separation was achieved using a ThermoFinnigan Surveyor MS-Pump with a BioBasic-18 100 mm C 0.18 mm reverse phase capillary column. Mass analysis was done with a ThermoFinnigan LQ Deca XP Plus ion trap mass spectrometer equipped with a nanospray ion source employing a 4.5 cm long metal needle, in the data-dependent acquisition mode. Electrical contact and voltage application to the probe tip took place via the nanoprobe assembly. Spray voltage was set to 2.9 kV and heated capillary temperature at 190°C. The column was equilibrated for 5 min with 95% solution A, 5% solution B (A, 0.1% formic acid in H2O; B, 0.1 % formic acid in acetonitrile) prior to sample injection. A linear gradient was initiated 5 min after sample injection ramping to 35% A, 65% B after 50 min and 20% A, 80% B after 60 min. Mass spectra were acquired in the m/z 400–800 range.

  3. Protein identifications were carried out with the MS/MS search software Mascot 1.9 (Matrix Science) with confirmatory or complementary analyses with TurboSequest as implemented in the Bioworks Browsers 3.2, build 41 (ThermoFinnigan). NCBI Sus scrofa protein sequences were used as the primary search database; searches were complemented with the NCBI non-redundant protein database. The presence of nitrotyrosine rendered an additional mass of 45 daltons to the peptides.

Statistical Analysis

Data are expressed as mean ± SD and compared among separate experiments. For comparisons between two groups, statistical analysis was performed using the two sample independent-groups t-test. Comparisons of multiple mean values were made by one-way analysis of variance (ANOVA), and statistical significance among multiple groups determined using the Tukey procedure (for pairwise comparisons of means between static-like and pulsatile flow conditions). P-values of < 0.05 are considered statistically significant.

Results

17-β Estradiol Down-Regulated Oscillatory Shear Stress-Mediated NADPH Oxidase Subunit (Nox4) Expression and O2−· Production

Under basal static cell culture conditions, Nox4 mRNA expression was markedly down-regulated in BAEC by E2 compared with that of control cells (Fig. 2). Nox4 mRNA expression was further up-regulated in response to OSS; however, the presence of E2 (5nmol/L) inhibited OSS-mediated Nox4 mRNA expression by 35.0 ±4.2 % (P<0.05, n=4). Similarly, the rates of O2−· production in response to OSS increased steadily; however, the presence of E2 decreased OSS-mediated O2−· production (n=4, P<0.05) (Fig. 3), consistent with decreased Nox4 expression. The rates of O2−· production remained relatively steady under static conditions whereas the rates in the presence of E2 were lower than that of controls (n=4, P<0.05). The apparent negative rates for O2−· production were due to the lower measured values for optical densities (OD at 550 nm) after incubation as compared to initial values at time 0. Rates of O2−· production were correlated with Nox4 mRNA expression in the presence and absence of E2.

Fig. 2.

Fig. 2

Open in a new tab

Relative NADPH oxidase subunit (Nox4) mRNA expression normalized to GAPDH at 4 hours. In the presence of E2, Nox4 mRNA decreased significantly compared with control (*, P<0.05,n=4). Pre-treatment with E2 under oscillatory shear stress (OSS) significantly down-regulated Nox4 expression by 35.0±4.2% compared with OSS (P<0.05, n=4).

Fig. 3.

Fig. 3

Open in a new tab

17β-Estradiol (E2) inhibits OSS-induced superoxide (O2−·) production. The rate of O2−· production in BAEC in response to OSS increased steadily whereas treatment with E2 decreased the rate of O 2−· formation (n=4, P<0.05). The rate of O2−· production remained relatively steady under static conditions whereas the rate of O2−· production in the presence of E2 was relatively lower than that of controls (n=4, P<0.05). The negative rates for O2−· production were due to the lower measured values for optical densities (OD at 550 nm) after incubation with E2 as compared to initial values at time 0.

17-β Estradiol Up-regulated eNOS Expression and ·NO Production in the Presence of Oscillatory Shear Stress

The presence of E2 noticeably increased eNOS mRNA expression by 40-fold ± 5 under static conditions (Fig. 4). While eNOS mRNA was down-regulated in response to OSS, the presence of E2 suppressed the inhibitory effect of OSS by up-regulating eNOS mRNA expression by 20-fold ± 1 (n=4, P < 0.001). The changes in eNOS mRNA were also correlated with an increase in ·NO production in the presence of E2 as compared with the control conditions (3.98±0.14 μM versus 7.37±0.89 μM, respectively P<0.05, n=4) (Fig. 6). While OSS decreased ·NO formation, the presence of E2 restored ·NO production to levels intermediate between controls and E2 pre-treatment (OSS=3.66±0.64 μM versus E2+OSS = 5.65±0.26 μM, P<0.05, n=4) (Fig. 6a). The relative difference in NO2− production was statistically insignificant between OSS and E2+ OSS conditions (Control = 0.82±0.02 μM, E2 = 1.21 ± 0.06 μM, OSS = 0.55±0.03 μM, E2+OSS = 0.93±0.032 μM) (Fig. 6b). The reversing effect of E2 on total ·NO formation was reflected largely by the increased formation of NO3− (Control = 3.16±0.13μM, E2 = 6.16±0.68μM, OSS = 3.11±0.49 μM, E2+OSS = 4.73±0.23 μM) (Fig. 6c). Overall, the levels of ·NO production correlated with the general trend of eNOS mRNA expression. The amount of ·NO produced as a result of E2 treatment was verified by addition of L-NAME (100 μM) which mitigated E2-mediated ·NO production (Fig. 7).

Fig. 4.

Fig. 4

Open in a new tab

Relative eNOS mRNA expression normalized by GAPDH. E2 pre-treated BAEC was associated with a significant increase in eNOS mRNA expression (*, P< 0.001, n=4). OSS decreased eNOS mRNA expression while pre-treatment with E2 restored eNOS expression (**, P<0.001, n=4).

Fig. 6.

Fig. 6

Open in a new tab

Total NO, NO2− and NO3− production. (a) Total NO: Pretreatment of BAEC with E2 significantly increased total NO production compared to control (static) conditions. OSS decreased NO production compared to E2. Pretreatment with E2 increased NO production compared to OSS (P<0.01, n=4). (b) NO2− production: the relative difference in NO2− production in response to OSS vs. E2+OSS was statistically insignificant (P<0.05, n=4). (c) NO3− formation (total NO-NO2−): the reversing effect of E2 of total NO was reflected largely by the increased formation of NO3− (P<0.01, n=4).

Fig. 7.

Fig. 7

Open in a new tab

Addition of N(G)-nitro-l-arginine methyl ester (L-NAME) at 100 μM mitigated E2-mediated NO production (control: 4.04±0.37μM L-NAME: 2.03±0.68 μM E2 : 7.7±0.58 μM L-NAME+E2 : 4.09±0.47μM ).

17-β Estradiol Reverses OSS-Mediated LDL Oxidation

The presence of E2 decreased the extent of LDL modifications (LDL−) under both static conditions and OSS conditions as demonstrated by the % LDL− formation (Fig. 8). The chromatogram revealed two distinct sets of peaks: native LDL and LDL− under static and OSS conditions in the presence and absence of E2 (Fig. 8a). The relative levels of oxidized LDL as expressed in % LDL− formation were compared in response to OSS in the presence and absence of E2 pre-treatment (Fig. 8b). OSS induced the highest level of LDL modifications as measured by optical density (OD) of LDL−, followed by control, E2+OSS, and E2. While OSS significantly increased LDL modifications, the presence of E2 significantly attenuated OSS-induced LDL− formation (LDL− to LDL ratios: control=1.05±0.34, E2=0.58±0.18, OSS=2.28±0.41, OSS+E2=0.80±0.25, P<0.01, n=4). The extent of % LDL− formation correlated with the regulation of Nox4 mRNA expression and the relative rates of O2−· and ·NO production in the presence of E2 (Figs. 2,3 and 6).

Fig. 8.

Fig. 8

Open in a new tab

(a) Chromatogram. The first set of peaks reflects nLDL and the second LDL−. OSS induced the highest optical density (OD) of LDL−, followed by control, E2+OSS, and E2. (b) LDL− to total LDL ratios as a measure of LDL oxidative modification. E2 decreased LDL oxidation compared with control (*, P<0.05, n=5). While OSS increased oxidative modification of LDL, pre-treatment with E2 significantly decreased OSS-mediated LDL oxidation (**, P<0.01, n=5).

17-β Estradiol Reverses Xanthine Oxidase-Mediated LDL Modification via Reduction in Lipid Peroxides and Protein Nitration

To elucidate a potential mechanism by which E2 influenced LDL post-translational modifications in connection with the regulation of O2·− and ·NO production by endothelial cells, we studied xanthine oxidase-mediated LDL lipid oxidation and protein nitration. Addition of XO significantly increased LDL− formation (Fig. 9) and LDL lipid peroxidation (Fig. 10) compared with control, whereas pre-treatment with E2 reversed XO-induced LDL modifications (*, P<0.01, n=3). LC/MS/MS analyses of XO-mediated LDL apo B 100 nitration revealed a tyrosine residue 2535 modification in a tryptic peptide (Y*LSLGQVYISDWWTLAAK) from α-2 helix domains (Fig. 11), whereas pre-treatment with E2 reversed this observation.

Fig. 9.

Fig. 9

Open in a new tab

E2 reverses xanthine oxidase (XO)-mediated LDL oxidation. While addition of XO significantly increased LDL oxidation compared with control, pre-treatment with E2 reversed XO-induced LDL oxidation (*, P<0.05, n=3).

Fig. 10.

Fig. 10

Open in a new tab

(a) E2 reverses xanthine oxidase (XO)-mediated lipid peroxidation (LP). XO increased LP to total protein ratios; pre-treatment with E2 significantly decreased XO-mediated lipid peroxidation (**, P<0.01, n=3).

Fig. 11.

Fig. 11

Open in a new tab

MS/MS spectrum for tryptic peptide carrying a nitrative modification at tyrosine 2535. Above the spectrum is the peptide sequence with observed b and y ions indicated by their fragmentation number above (y ions) or below (b ions) the sequences. ++ indicates doubly charged ions. Single-charged ions are not designated with a +. The ion annotation is based on results presented in Sequest’s “display ions view” window that were corroborated by the data file. Y* indicates a nitrotyrosine.

Sequence coverage for LDL protein modifications in control and XO- and/or E2-treated samples was approximately 40% in samples subjected to LC/MS/MS. The tyrosine modification was observed over the course of analysis of three separate samples preparations. The tryptic peptide containing the identified tyrosine modification was observed by Mascot software analysis and LC/MS/MS analyses (Table 1). Nitration of tyrosine 2535 was observed in the same doubly charged 18 amino acid tryptic peptide in three separate LC/MS/MS analyses. Representative MS/MS spectrum for the tryptic peptide in the m/z 400–2000 range carrying a nitrative modification is presented (Fig. 11). The precursor ion was doubly charged with a mass of 1353.18 Da corresponding to an 18-residue tryptic peptide, comprising amino acids 2535 – 2559, with one nitration modification. Fragment b and y ions were observed for nearly all-observable single-charged fragments. LDL protein nitration was further corroborated by the dot blot analyses for nitrotyrosine (Fig. 12). While XO significantly increased nitrotyrosine intensity, E2 significantly decreased its intensity (**, P<0.01, n=3). Collectively, these findings suggest that E2 reverses LDL− formation via lipid peroxidation and post-translation apo B100 protein nitration.

Table I.

LDL apo B 100 modifications in the m/z 400 – 2000 range. Nitrated tyrosines in the amino acid sequence of the tryptic fragments are denoted as asterisks. “M” is the Mascot score. Nitrotyrosine is absent in the samples treated with LDL and E2 (n=3).

Treatments Structure Numerical sequence Protein Sequence Nitrotyrosine Mascot score
LDL control α–2 2535–2559 y*lslvgqvystlvtyisdwwtlaak 1 35
LDL + XO α–2 2535–2559 y*lslvgqvystlvtyisdwwtlaak 1 25
LDL + E2 α–2 2535–2559 ylslvgqvystlvtyisdwwtlaak 0 NA
LDL + E2 + XO α–2 2535–2559 y*lslvgqvystlvtyisdwwtlaak 1 10
Open in a new tab

Fig. 12.

Fig. 12

Open in a new tab

(a) Dot plot for nitrotyrosine as a finger print for LDL protein nitration. (b) Densitometry of nitrotyrosine. E2 attenuates XO-induced LDL protein nitration as measured by the presence of nitrotyrosine. While XO increased LDL protein nitration, pre-treatment with E2 significantly decreased XO-mediated LDL nitration.

Discussion

The main finding of this study was that at a physiological concentration (5 nmol/L), E2 reversed OSS-mediated LDL oxidation. In the presence of E2, expression of the NADPH oxidase (Nox4) was down-regulated, whereas expression of eNOS was up-regulated. These changes in Nox4 and eNOS expression coincided with a decrease in O2−· formation and an increase in ·NO production, respectively, suggesting that the effect of E2 was mediated, in part, via concomitant down-regulation of NADPH oxidase (Nox4) and up-regulation of eNOS. This relative expression of Nox4 and eNOS indicates that E2 may act at the transcriptional level to suppress oxidant production.

Vascular cells use reactive oxygen (ROS) and reactive nitrogen species (RNS) to modify LDL. ECs produce ROS and RNS from the enzymes and ·NO synthase and by specific homologues of NADPH oxidase (gp91phox, Nox1, and Nox4) [41]. We previously demonstrated that NADPH oxidases were capable of producing high levels of ROS in blood vessels in response to specific stimuli arising from flow conditions. Consequently, increasing quantities of modified LDL species were formed under oscillatory flow conditions that favor ROS production, whereas lower levels of LDL modification occur in response to pulsatile flow (unidirectional forward flow) conditions that produce lower levels of ROS [5]. Here, we further showed that estrogen attenuated the levels of LDL modification, suggesting linked effects between pulsatile shear stress and estrogen. The increase in NAD(P)H oxidase-derived O2−· level can account for the cross-talks with endothelium-derived ·NO ; thus, the balance between O2−· and ·NO production may influence the ·NO signaling function, antioxidant action and generation of ONOO− [42, 43]. The exposure of LDL to ONOO− results in the nitration of apo B-100 tyrosine residues [44], the initiation of lipid peroxidation [45, 46], accompanied by the depletion of lipid-soluble antioxidants [47, 48].

Tyrosine nitration, a post-translational modification of proteins through the addition of a nitro (NO2) group in the ortho position of tyrosine residues [49], has been detected under physiological settings and in a number of pathological states including inflammatory and septic conditions [46, 50, 51]. Peroxynitrite reacts rather specifically with tyrosine residues in proteins leading to the formation of nitrotyrosine, which in itself could be toxic, e.g., by undergoing redox cycling, by interfering with signal transduction, or by becoming incorporated into the microtubules protein tubulin and distorting the cytoskeleton [52].

Misfolding of apoB may account for one of the mechanisms for LDL− formation. The α-2 and α-3 domains of apoB-100 contain the highest number of tyrosine residues (4.8% and 6.8% tyrosine respectively), and nitrotyrosine may play an important role in α helical integrity of apoB-100 [42]. Nitration of tyrosine residues could lend to a more hydrophilic residue carrying a dipole moment, altering the intact structure of α helices and other domains of the protein on the surface of the LDL particles. Mass spectra analysis (Fig. 11) demonstrated that pre-treatment with E2 suppressed xanthine oxidase-mediated LDL protein nitration at tyrosine 2535 in the α-2 domain. These findings are consistent with dot blots for nitrotyrosine as a finger print for peroxynitrite-mediated nitration that reacts rather specifically with tyrosine residues (Fig. 12). Thus, E2 plays an indirect antioxidant role in LDL− formation by reversing xanthine oxidase-mediated LDL lipid peroxidation and LDL apo B 100 nitration.

A subclass of LDL described on the basis of its greater electronegativity and oxidative status has been previously characterized [25, 26, 53]. These particles, which were referred to as LDL−, are enriched with lipid hydroperoxides and other peroxidation products as compared to the bulk of the unmodified, normal LDL (nLDL) recovered from human plasma. We have demonstrated that LDL− is a major carrier of lipid hydroperoxides and cholesterol oxides associated with plasma LDL and that LDL− may arise from oxidative events in the vasculature [54]. Decreased rates of O2−· production in glucose free medium containing 2-Deoxyglucose further implicate NADPH as an important cofactor for NADPH oxidase activity and the necessity of this activity to produce O2−· and modify LDL [5]. The variation in LDL− levels formed by BAECs under OSS conditions points to a role for NADPH oxidase activation in cell-mediated LDL modification.

The atheroprotective effect of estrogen favors women until menopause [55]. Several reports suggest that the atheroprotective effect of E2 may be mediated by increased local concentrations of ·NO [9, 56] through up-regulation of eNOS expression [57] and/or an estrogen receptor-mediated activation of eNOS via the phosphatidylinositol-3-(PI3) kinase/Akt kinase pathway [58, 59]. E2 transcriptionally up-regulates cytochrome P450 (CYP) activity leading to the enhanced arteriolar vasodilation response to shear stress in ·NO-deficient female rats and mice [1]. Interaction of cell membrane-bound estrogen receptors with PI3 kinase phosphorylates Akt induces phosphorylation of eNOS and release of ·NO [48, 60]. By inhibiting NADPH oxidase activity, E2 can exert an indirect ‘antioxidant’ effect through the reduced formation and interaction between O2−· and ·NO. Thus, the presence of estrogen exerts salutary effects on vascular endothelium largely via an increase in ·NO formation and a decrease in oxidant production.

In figure 3, the effect of E2 on the oxidative arm appears to be modest. There was a 35% change in Nox4 mRNA (Fig. 2); however, the effect of E2 on O2−· production appears not to be a reduction in NADPH oxidase but activities. One possibility was an increased loss of already formed O2−·. Another possibility was the effects of E2 on other sources of O2−· production, including xanthine oxidase and mitochondria [61, 62].

Furthermore, ·NO was metabolized or decomposed via various reactions to the major products NO2− and NO3− in vivo. They served as useful measurement of overall ·NO production and metabolism, both under basal physiological conditions, as well as during exposure to xenobiotics and/or inflammatory response. Although NO2− could also react with heme proteins and participate in further reactions (including nitration), NO3− was an ultimate end product [63]. However, in Figure 6a, ·NO production was comparable between control and OSS groups. OSS-mediated Nox4 expression and consequently, O2−· production (Figs. 2 and 3), was likely to react with ·NO to form peroxynitrite (ONOO−). ONOO− would further decompose upon protonation through rapid isomerization to NO3− at physiologic pH [64]. Thus, the formation of ONOO− could account for the paradoxical increase in NO3− between the control and OSS groups (Fig. 6c). In this context, measuring total NO3− and NO2− would not precisely reflect an impaired ·NO bioavailability in the presence of increased O2−· formation.

The increase in eNOS mRNA by E2 was impressive (Fig. 4); however, there was only a 2-fold change in ·NO under control conditions and a smaller change under OSS. Two reasons may have accounted for the disparity between eNOS mRNA expression and ·NO production. First of all, statistic analysis of real-time RT-PCR could have accentuated the differences between relative eNOS mRNA expressions. We used the delta delta Ct method as follows: CT is the threshold cycle number at which the initial amplification becomes detectable by fluorescence [65]. The difference in CT values for various flow conditions vs. control was used to mathematically determine the relative difference in the level of eNOS mRNA expression [65]. For quantification of relative eNOS mRNA expression, the difference in CT values was normalized to the expressed housekeeping gene GAPDH. Secondly, a loss of already formed ·NO could have attributed to the relatively small change in ·NO. Quantitative measurements of NO2− and NO3− were performed as an index of global nitric oxide (·NO) production [33,34]. Collected cell culture medium was incubated with nitrate reductase for 2 hours to convert NO3− to NO2− [3], followed by addition of Griess reagents. Quantification of NO2− levels were performed spectrophotometrically.

We demonstrated that E2 attenuated the extent of LDL oxidation in response to OSS. The hemodynamic forces, to which endothelial cells are exposed in vivo, especially at sites of arterial bifurcation, can predispose these regions to atherogenic processes that include up-regulation of O2−· formation [8] and subsequent oxidative reactions. Clinically known risk factors for coronary heart disease have been associated with endothelial dysfunction [66]. Enhanced oxidative stress may be another factor contributing to this dysfunction, although controversy remains [66]. A recent study in normotensive women showed that endothelial dysfunction secondary to acute endogenous estrogen deprivation is strongly linked to a reduction in the bioavailability of ·NO [67]. Moreover, E2 can attenuate O2−· formation in phagocytes [68] and endothelial cells [6]. In summary, the reduction in O2−· formation combined with the salutary effects of ·NO may account for decreased LDL oxidative modification by endothelial cells. Furthermore, E2 suppresses xanthine oxidase-induced LDL post-translational modifications via reductions in both lipid peroxidation and protein nitration. This indirect antioxidant activity may provide a new insight into a potential mechanism whereby E2 plays an atheroprotective role.

Fig. 5.

Fig. 5

Open in a new tab

Nox4 protein expression normalized with β-actin at 4 hours. E2 decreased Nox4 protein by 61±3% compared with control (*, P<0.05, n=4). E2 attenuated OSS-induced Nox4 protein expression by 23.9±5.4% compared with OSS alone (**, P<0.05, n=4).

Acknowledgments

This manuscript is dedicated to Alex Sevanian who passed away in 2005. This study was supported by American Heart Association BGI-0265166U (T. K. H.), National Institutes of Heath grants HL KO8-068689 (T.K.H), R01HL050350 (A. S., T. K. H., H. N. H.), NIA RO1AG-18798 (H. N. H.), RO1AG-17160 (H. N. H.), RO1AG-24154 (H. N. H.), NCCAM RO1AT001653 (H. N. H.), and National Heart Foundation/American Health Assistance Foundation H2003-028 (T. K. H.).

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  • 1.Huang ASD, Wu Z, Yan C, Carroll MA, Jiang H, Falck GK., JR Estrogen elicits cytochrome P450--mediated flow-induced dilation of arterioles in NO deficiency: role of PI3K-Akt phosphorylation in genomic regulation. Circ Res. 2004;94:245–52. doi: 10.1161/01.RES.0000111525.96232.46. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Fung Y. Biomechanics: Circulation. 2. Springer; 1997. [Google Scholar]
  • 3.Couzin J. Estrogen research. The great estrogen conundrum. Science. 2003;302:1136–8. doi: 10.1126/science.302.5648.1136. [DOI] [PubMed] [Google Scholar]
  • 4.Hsiai TK, CS, Wong PK, Ing M, Salazar A, Sevanian A, Navab M, Demer LL, Ho CM. Monocyte recruitment to endothelial cells in response to oscillatory shear stress. Faseb J. 2003;17:1648–57. doi: 10.1096/fj.02-1064com. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Hwang J, IM, Salazar A, Lassegue B, Griendling K, Navab M, Sevanian A, Hsiai TK. Pulsatile versus oscillatory shear stress regulates NADPH oxidase subunit expression: implication for native LDL oxidation. Circ Res. 2003;93:1225–32. doi: 10.1161/01.RES.0000104087.29395.66. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Wagner A, Schroeter MR, Hecker M. 17β-Estradiol inhibition of NADPH oxidase expression in human endothelial cells. The FASEB Journal. 2001;15:2121–2130. doi: 10.1096/fj.01-0123com. [DOI] [PubMed] [Google Scholar]
  • 7.De Keulenaer GW, CD, Ishizaka N, Nerem RM, Alexander RW, Griendling KK. Oscillatory and steady laminar shear stress differentially affect human endothelial redox state: role of a superoxide-producing NADH oxidase. Circ Res. 1998;82:1094–1101. doi: 10.1161/01.res.82.10.1094. [DOI] [PubMed] [Google Scholar]
  • 8.Hwang J, SA, Boo YC, Sorescu GP, McNally JS, Holland SM, Dikalov S, Giddens DP, Griendling KK, Harrison DG, Jo H. Oscillatory shear stress stimulates endothelial production of O2− from p47phox-dependent NAD(P)H oxidases, leading to monocyte adhesion. J Biol Chem. 2003;278:47291–8. doi: 10.1074/jbc.M305150200. [DOI] [PubMed] [Google Scholar]
  • 9.Hodis HN, MW, Azen SP, Lobo RA, Shoupe D, Mahrer PR, Faxon DP, Cashin-Hemphill L, Sanmarco ME, French WJ, Shook TL, Gaarder TD, Mehra AO, Rabbani R, Sevanian A, Shil AB, Torres M, Vogelbach KH, RHS Women’s Estrogen-Progestin Lipid-Lowering Hormone Atherosclerosis Regression Trial Research Group. N Engl J Med. 2003;349:535–45. doi: 10.1056/NEJMoa030830. [DOI] [PubMed] [Google Scholar]
  • 10.Hodis HN, MW, RAL Randomized controlled trial evidence that estrogen replacement therapy reduces the progression of subclinical atherosclerosis in healthy postmenopausal women without preexisting cardiovascular disease. Circulation. 2003;108:e5. doi: 10.1161/01.CIR.0000080080.76333.38. [DOI] [PubMed] [Google Scholar]
  • 11.Rubanyi GM, Freay AD, Kauser K, Sukovich D, Burton G, Lubahn DB, Couse JF, Curtis SW, Korach KS. Vascular estrogen receptors and endothelium-derived nitric oxide production in the mouse aorta. Gender difference and effect of estrogen receptor gene disruption. J Clin Invest. 1997;99:2429–2437. doi: 10.1172/JCI119426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Sumi D, LI Estrogen-related receptor alpha 1 up-regulates endothelial nitric oxide synthase expression. Proc Natl Acad Sci U S A. 2003;100:14451–6. doi: 10.1073/pnas.2235590100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Huang A, SD, Koller A, GK 17beta-estradiol restores endothelial nitric oxide release to shear stress in arterioles of male hypertensive rats. Circulation. 2000;101:94–100. doi: 10.1161/01.cir.101.1.94. [DOI] [PubMed] [Google Scholar]
  • 14.Wilcox JG, HJ, Hodis HN, Sevanian A, Stanczyk FZ, Lobo RA. Cardioprotective effects of individual conjugated equine estrogens through their possible modulation of insulin resistance and oxidation of low-density lipoprotein. Fertil Steril. 1997;67:57–62. doi: 10.1016/s0015-0282(97)81856-0. [DOI] [PubMed] [Google Scholar]
  • 15.Hwang J, PH, Hodis HN, Choi B, Sevanian A. Ascorbic acid enhances 17 beta-estradiol-mediated inhibition of oxidized low density lipoprotein formation. Atherosclerosis. 2000;150(2):275–84. doi: 10.1016/s0021-9150(99)00376-7. [DOI] [PubMed] [Google Scholar]
  • 16.Demaio L, Rouhanizadeh M, Reddy S, Sevanian A, Hwang J, Hsiai T. Oxidized phospholipids mediate occludin expression and phosphorylation in vascular endothelial cells. Am J Physiol Heart Circ Physiol. 2006;290(2):H674–83. doi: 10.1152/ajpheart.00554.2005. [DOI] [PubMed] [Google Scholar]
  • 17.Van Nieuw Amerongen GP, VM, Negre-Aminou P, Lankelma J, Emeis JJv, VWaH Simvastatin improves disturbed endothelial barrier function. Circulation. 2000;23:2803–9. doi: 10.1161/01.cir.102.23.2803. [DOI] [PubMed] [Google Scholar]
  • 18.Buchanan JRJ, Kleinstreuter C, Truskey GA, Lei M. Relation between nonuniform hemodynamics and sites of altered permeability and lesion growth at the rabbit aorto-celiac junction. Atherosclerosis. 1999;26:215–24. doi: 10.1016/s0021-9150(98)00264-0. [DOI] [PubMed] [Google Scholar]
  • 19.Deng X, MT, How T, Merhi M, King RM, Guidoin, Karino T. Luminal surface concentration of lipid (LDL) and its effect on the wall uptake of cholesterol by canine carotid arteries. J Vasc Surg. 1995;21:135–45. doi: 10.1016/s0741-5214(95)70252-0. [DOI] [PubMed] [Google Scholar]
  • 20.Henry PD, CC Inflammatory mechanisms of atheroma formation. Influence of fluid mechanics and lipid-derived inflammatory mediators. Am J Hypertension. 1993;6:328S–334S. doi: 10.1093/ajh/6.11.328s. [DOI] [PubMed] [Google Scholar]
  • 21.Steinberg D, PS, Carew TE, Khoo JC, Witztum JL. Beyond cholesterol. Modifications of low-density lipoprotein that increase its atherogenicity [see comments] N Engl J Med. 1989;320:915–24. doi: 10.1056/NEJM198904063201407. [DOI] [PubMed] [Google Scholar]
  • 22.Witztum J. The oxidation hypothesis of atherosclerosis. Lancet. 1994;344:793–5. doi: 10.1016/s0140-6736(94)92346-9. [DOI] [PubMed] [Google Scholar]
  • 23.Ku DN, GD, Zarins CK, Glagov S. Pulsatile flow and atherosclerosis in the human carotid bifurcation. Positive correlation between plaque location and low oscillating shear stress. Arteriosclerosis. 1985;5:293–302. doi: 10.1161/01.atv.5.3.293. [DOI] [PubMed] [Google Scholar]
  • 24.Hsiai TK, CS, Honda HM, Hama S, Navab M, Demer LL, Ho CM. Endothelial cell dynamics under pulsating flows: significance of high versus low shear stress slew rates (d(tau)/dt. Ann Biomed Eng. 2002;30(5):646–56. doi: 10.1114/1.1484222. [DOI] [PubMed] [Google Scholar]
  • 25.Hodis HN, KD, Avogaro P, Bittolo-Bon G, Cazzolato G, Hwang J, Peterson H, Sevanian A. Biochemical and cytotoxic characteristics of an in vivo circulating oxidized low density lipoprotein (LDL−) J Lipid Res. 1994;35:669–77. [PubMed] [Google Scholar]
  • 26.Sevanian A, SR, Traldi P, Rossato P, Ursini F, Hodis H. Analysis of plasma cholesterol oxidation products using gas- and high- performance liquid chromatography/mass spectrometry. Free Radic Biol Med. 1994;17:397–409. doi: 10.1016/0891-5849(94)90166-x. [DOI] [PubMed] [Google Scholar]
  • 27.Nerem RM, AR, Chappell DC, Medford RM, Varner SE, Taylor WR. The study of the influence of flow on vascular endothelial biology. Am J Med Sci. 1998;316:169–75. doi: 10.1097/00000441-199809000-00004. [DOI] [PubMed] [Google Scholar]
  • 28.Papadaki MML. Methods in Molecular Medicine: Tissue Engineering Methods and Protocols. In: Yarmush ML, editor. Quantitative Measurement of Shear-Stress Effects on Endothelial Cells. Totowa, NJ: Humana Press Inc; 1998. pp. 577–593. [DOI] [PubMed] [Google Scholar]
  • 29.Hsiai TWP, Ing M, Navab M, Reddy S, Ho CM. Bio-MEMS sensors for real-time shear stress on endothelial cell dynamics. :7.4.2–5. The Second Joint Meeting of the IEEE Engineering in Medicine and Biology and the Biomedical Engineering Society; 2002; Huston, USA. [Google Scholar]
  • 30.Asatryan L, HR, Isas JM, Hwang J, Kayed R, Sevanian A. LDL phospholipid hydrolysis produces modified electronegative particles with an unfolded apoB-100 protein. J Lipid Res. 2005;46(1):115–22. doi: 10.1194/jlr.M400306-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Chu Y, HD, Knudtson KL, Lamping KG, Faraci FM. Quantification of mRNA for endothelial NO synthase in mouse blood vessels by real-time polymerase chain reaction. Arterioscler Thromb Vasc Biol. 2002;22:611–6. doi: 10.1161/01.atv.0000012663.85364.fa. [DOI] [PubMed] [Google Scholar]
  • 32.Livak KJ, ST Analysis of relative gene expression data using real-time quantitative PCR and the 2(−Delta Delta C(T)) Method. Methods. 2001;25(4):402–8. doi: 10.1006/meth.2001.1262. [DOI] [PubMed] [Google Scholar]
  • 33.Walker N. A technique whose time has come. Science. 2002;296:557–559. doi: 10.1126/science.296.5567.557. [DOI] [PubMed] [Google Scholar]
  • 34.Braman RS, HS Nanogram nitrite and nitrate determination in environmental and biological materials by vanadium (III) reduction with chemiluminescence detection. Anal Biochem. 1989;61:2715–2718. doi: 10.1021/ac00199a007. [DOI] [PubMed] [Google Scholar]
  • 35.Van Der Vliet A, NMN, Shigenaga MK, Eiserich JP, Marelich GP, CEC Myeloperoxidase and protein oxidation in cystic fibrosis. Am J Physiol. 2000;279:L537–L546. doi: 10.1152/ajplung.2000.279.3.L537. [DOI] [PubMed] [Google Scholar]
  • 36.Clarkson RB, NS, Smirnov A, Boyer S, Vahidi N, Nims RW, Wink DA. Direct measurement of the accumulation and mitochondrial conversion of nitric oxide within Chinese hamster ovary cells using an intracellular electron paramagnetic resonance technique. Biochim Biophys Acta. 1995;1243:496–502. doi: 10.1016/0304-4165(94)00181-v. [DOI] [PubMed] [Google Scholar]
  • 37.Crow JP, JB Reactions between nitric oxide, superoxide, and peroxynitrite: footprints of peroxynitrite in vivo. Adv Pharmacol. 1995;34:17–43. doi: 10.1016/s1054-3589(08)61079-0. [DOI] [PubMed] [Google Scholar]
  • 38.Espey MG, MK, Thomas DD, Xavier S, Citrin D, Vitek MP, DAW A chemical perspective on the interplay between NO, reactive oxygen species, and reactive nitrogen oxide species. Ann N Y Acad Sci. 2002;962:195–206. doi: 10.1111/j.1749-6632.2002.tb04068.x. [DOI] [PubMed] [Google Scholar]
  • 39.Aviram M, Rosenblat M, Etzioni A, Levy R. Activation of NADPH oxidase required for macrophage-mediated oxidation of low-density lipoprotein. Metabolism. 1996;45:1069–79. doi: 10.1016/s0026-0495(96)90005-0. [DOI] [PubMed] [Google Scholar]
  • 40.Navab M, BJ, Watson AD, Hama SY, Territo MC, Lusis AJ, Shih DM, Van Lenten BJ, Frank JS, Demer LL, Edwards PA, Fogelman AM. The Yin and Yang of oxidation in the development of the fatty streak. A review based on the 1994 George Lyman Duff Memorial Lecture. 1996; Arterioscler Thromb Vasc Biol. 1996;16:831–42. doi: 10.1161/01.atv.16.7.831. [DOI] [PubMed] [Google Scholar]
  • 41.Szocs K, Lassegue B, Sorescu D, Hilenski LL, Valppu L, Couse TL, Wilcox JN, Quinn MT, Lambeth JD, Griendling KK. Upregulation of Nox-based NAD(P)H oxidases in restenosis after carotid injury. Arterioscler Thromb Vasc Biol. 2002;22:21–27. doi: 10.1161/hq0102.102189. [DOI] [PubMed] [Google Scholar]
  • 42.Giordano F. Oxygen, oxidative stress, hypoxia and heart failure. J Clin Invest. 2005;115:500–8. doi: 10.1172/JCI200524408. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43.Madamanchi NR, VA, Runge MS. Oxidative stress and vascular disease. Arterioscler Thromb Vasc Biol. 2005;25:29–38. doi: 10.1161/01.ATV.0000150649.39934.13. [DOI] [PubMed] [Google Scholar]
  • 44.Leeuwenburgh C, Hardy MM, Hazen SL, Wagner P, Ohishi S, Steinbrecher UP, Heinecke JW. Reactive nitrogen intermediates promote low density lipoprotein oxidation in human atherosclerotic intima. J Biol Chem. 1997;272:1433–1436. doi: 10.1074/jbc.272.3.1433. [DOI] [PubMed] [Google Scholar]
  • 45.Radi R, Beckman JS, Bush KM, Freeman BA. Peroxynitrite-induced membrane lipid peroxidation: the cytotoxic potential of superoxide and nitric oxide. Arch Biochem Biophys. 1991;288:481–487. doi: 10.1016/0003-9861(91)90224-7. [DOI] [PubMed] [Google Scholar]
  • 46.Shishehbor M, Aviles RJ, Brennan ML, Fu X, Goormastic M, Pearce GL, Gokce N, Keaney JF, Jr, Penn MS, Sprecher DL, Vita JA, Hazen SL. Association of nitrotyrosine levels with cardiovascular disease and modulation by statin therapy. JAMA. 2003;289:1675–80. doi: 10.1001/jama.289.13.1675. [DOI] [PubMed] [Google Scholar]
  • 47.Hogg N, D-UV, Wilson MT, Moncada S. The oxidation of alpha-tocopherol in human low-density lipoprotein by the simultaneous generation of superoxide and nitric oxide. FEBS Lett. 1993;326. FEBS Lett. 1993;326(1–3):199–203. doi: 10.1016/0014-5793(93)81790-7. [DOI] [PubMed] [Google Scholar]
  • 48.Goss S, Hogg N, Kalyanaraman B. The effect of alpha-tocopherol on the nitration of gamma-tocopherol by peroxynitrite. Arch Biochem Biophys. 1999;363(2):333–40. doi: 10.1006/abbi.1998.1094. [DOI] [PubMed] [Google Scholar]
  • 49.Ischiropoulos H, et al. Peroxynitrite-Mediated Tyrosine Nitration Catalyzed by Superoxide-Dismutase. Archives of Biochemistry and Biophysics. 1992;298(2):431–437. doi: 10.1016/0003-9861(92)90431-u. [DOI] [PubMed] [Google Scholar]
  • 50.Podrez EA, et al. Myeloperoxidase-generated reactive nitrogen species convert LDL into an atherogenic form in vitro. Journal of Clinical Investigation. 1999;103(11):1547–1560. doi: 10.1172/JCI5549. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Foster MW, McMahon TJ, Stamler JS. S-nitrosylation in health and disease. Trends in Molecular Medicine. 2003;9(4):160–168. doi: 10.1016/s1471-4914(03)00028-5. [DOI] [PubMed] [Google Scholar]
  • 52.Zhang YJ, et al. Nitration and oligomerization of tau induced by peroxynitrite inhibit its microtubule-binding activity. Febs Letters. 2005;579(11):2421–2427. doi: 10.1016/j.febslet.2005.03.041. [DOI] [PubMed] [Google Scholar]
  • 53.Sevanian A, HJ, Hodis H, Cazzolato G, Avogaro P, Bittolo-Bon G. Contribution of an in vivo oxidized LDL to LDL oxidation and its association with dense LDL subpopulations. Arterioscler Thromb Vasc Biol. 1996;16:784–93. doi: 10.1161/01.atv.16.6.784. [DOI] [PubMed] [Google Scholar]
  • 54.Sevanian A, B-BG, Cazzolato G, Hodis H, Hwang J, Zamburlini A, Maiorino M, Ursini F. LDL− is a lipid hydroperoxide-enriched circulating lipoprotein. J Lipid Res. 1997;38:419–28. [PubMed] [Google Scholar]
  • 55.Turgeon JL, MD, Martin KA, PMW Hormone therapy: physiological complexity belies therapeutic simplicity. Science. 2004;304:1269–73. doi: 10.1126/science.1096725. [DOI] [PubMed] [Google Scholar]
  • 56.Guetta V, QA, Prasad A, Panza JA, Waclawiw M, Cannon RO. The role of nitric oxide in coronary vascular effects of estrogen in postmenopausal women. Circulation. 1997;95:2795–2801. doi: 10.1161/01.cir.96.9.2795. [DOI] [PubMed] [Google Scholar]
  • 57.Yang S, BL, Zhang L. Estrogen increases eNOS and NOx release in human coronary artery endothelium. J Cardiovasc Pharmacol. 2000;36:242–247. doi: 10.1097/00005344-200008000-00015. [DOI] [PubMed] [Google Scholar]
  • 58.Busse R, FI Endothelial dysfunction in atherosclerosis. J Vasc Res. 1996;33:181–194. doi: 10.1159/000159147. [DOI] [PubMed] [Google Scholar]
  • 59.Haynes MP, SD, Russell KS, Collinge M, Fulton D, Morales-Ruiz M, Sessa WC, Bender JR. Membrane estrogen receptor engagement activates endothelial nitric oxide synthase via the PI3-kinase-Akt pathway in human endothelial cells. Circ Res. 2000;87:677–682. doi: 10.1161/01.res.87.8.677. [DOI] [PubMed] [Google Scholar]
  • 60.Simoncini T, H-MA, Brazil DP, Ley K, Chin WW, JKL Interaction of oestrogen receptor with the regulatory subunit of phosphatidylinositol-3-OH kinase. Nature. 2000;407:538–541. doi: 10.1038/35035131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 61.Budhiraja R, et al. Estrogen modulates xanthine dehydrogenase/xanthine oxidase activity by a receptor-independent mechanism. Antioxidants & Redox Signaling. 2003;5(6):705–711. doi: 10.1089/152308603770380007. [DOI] [PubMed] [Google Scholar]
  • 62.Vina J, Borras C, Gomez-Cabrera MC, Orr WC. Part of the series: from dietary antioxidants to regulators in cellular signaling and gene expression. Role of reactive oxygen species and (phyto)oestrogens in the modulation of adaptive response to stress. Free Radic Res. 2006;40(2):111–9. doi: 10.1080/10715760500405778. [DOI] [PubMed] [Google Scholar]
  • 63.Pietraforte D, SA, Scorza GMM. Scavenging of reactive nitrogen species by oxygenated hemoglobin: globin radicals and nitrotyrosines distinguish nitrite from nitric oxide reaction. Free Radic Biol Med. 2004;37(8):1244–55. doi: 10.1016/j.freeradbiomed.2004.06.027. [DOI] [PubMed] [Google Scholar]
  • 64.Edwards JO, Plumb RC. The chemistry of peroxynitrites. Prog Inorg Chem. 1994;41:599–635. [Google Scholar]
  • 65.Ischiropoulos H, et al. Peroxynitrite-mediated tyrosine nitration catalyzed by superoxide dismutase. Arch Biochem Biophys. 1992;298:431–437. doi: 10.1016/0003-9861(92)90431-u. [DOI] [PubMed] [Google Scholar]
  • 66.Vogel R. Coronary risk factors, endothelial function, and atherosclerosis: a review. Clin Cardiol. 1997;20:426–432. doi: 10.1002/clc.4960200505. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 67.Virdis A, GL, Pinto S, Lombardo M, Petraglia F, Gennazzani A, Buralli S, Taddei S, Salvetti A. Mechanisms responsible for endothelial dysfunction associated with acute estrogen deprivation in normotensive women. Circulation. 2000;101:2258–2263. doi: 10.1161/01.cir.101.19.2258. [DOI] [PubMed] [Google Scholar]
  • 68.Bekesi GKR, Varbiro S, Racz K, Sprintz D, Feher J, Szekacs B. In vitro effects of different steroid hormones on superoxide anion production of human neutrophil granulocytes. Steroids. 2000;65:889–894. doi: 10.1016/s0039-128x(00)00183-5. [DOI] [PubMed] [Google Scholar]

RESOURCES