Peroxisome proliferators compete and ameliorate Hcy-mediated endocardial endothelial cell activation

Matthew J. Hunt and Suresh C. Tyagi

Department of Physiology and Biophysics, University of Mississippi Medical Center, Jackson, Mississippi 39216


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

To determine whether homocysteine (Hcy)-mediated activation of endocardial endothelial (EE) cells is ameliorated by peroxisome proliferator-activated receptor (PPAR), we isolated EE cells from mouse endocardium. Matrix metalloproteinase (MMP) activity and intercellular adhesion molecule (ICAM)-1 in EE cells were measured in the presence and absence of Hcy, and ciprofibrate (CF; PPAR-alpha agonist) or 15-deoxy-Delta 12,14-prostaglandin J2 (PGJ2; PPAR-gamma agonist) by zymography and Western blot analyses, respectively. Results suggest that Hcy-mediated MMP activation and ICAM-1 expression are ameliorated by CF and PGJ2. To test the hypothesis that Hcy competes with other ligands for binding to PPARalpha and -gamma , we prepared cardiac nuclear extracts. Extracts were loaded onto an Hcy-cellulose affinity column. Bound proteins were eluted with CF and PGJ2. To determine conformational changes in PPAR upon binding to Hcy, we measured PPAR fluorescence at 334 nm. Dose-dependent increase in PPAR fluorescence demonstrated a primary binding affinity of 0.32 ± 0.06 µM. There was dose-dependent quenching of PPAR fluorescence by fluorescamine-homocysteine (F-Hcy). PPAR-alpha fluorescence quenching was abrogated by the addition of CF but not by PGJ2. PPAR-gamma fluorescence quenching was abrogated by the addition of PGJ2 but not by CF. These results suggest that Hcy competes with CF and PGJ2 for binding to PPAR-alpha and -gamma , respectively, indicating a role of PPAR in amelioration of Hcy-mediated EE dysfunction.

metalloproteinase; prostaglandin; fibrate; leukotriene; receptor; binding; microvessel; hydroxyeicosatetraeonic acid; epoxyeicosatrienoic acid; fluorescence resonance energy transfer


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
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DISCUSSION
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HOMOCYSTEINE (Hcy) induces nuclear factor-kappa B (NF-kappa B) (6, 66). On the other hand, a negative correlation between high levels of Hcy and peroxisome proliferator-activated receptor (PPAR) expression has been demonstrated (7, 24). PPAR induces a proliferation of the organelle within liver cells, where fatty acid oxidation takes place (51). These peroxisomes increased in both size and number in response to proliferator treatment, with a corresponding drop in circulating plasma lipids (20, 49). There is also an induction of expression of an entire suite of genes associated with fatty acid synthesis, transport, and catabolism. PPAR-alpha is expressed in adipose, liver, heart, muscle, and kidney (41). PPAR-beta is ubiquitous throughout the body. PPAR-gamma is most closely associated with fat (41). Although arachidonic acid (AA), metabolite hydroxyeicosatetraenoic acid (HETE), and leukotriene B4 (LB4) are the endogenous ligands of PPAR-alpha (11, 71), AA binds both PPAR-alpha and PPAR-gamma . On the other hand, LB4 binds preferentially PPAR-alpha (11). Among most of the fibrates, ciprofibrate has the most potent effect in ameliorating endothelial dysfunction (17). The 15-deoxy-Delta 12,14-prostaglandin J2 (PGJ2) is a potent agonist of PPAR-gamma (19). PPAR agonists promote anti-inflammatory effects (33). Although a direct connection of fatty acid metabolism and anti-inflammatory activity of PPAR is unclear, one might link the induction of antioxidant enzymes by PPAR with its anti-inflammatory activity. PPAR, upon induction, promotes the synthesis of SOD and catalase (22, 44). Meanwhile, PPAR decreases NADPH oxidase (22, 23). The inverse relationship between Hcy and PPAR is also unclear; however, elevation of Hcy is associated with a decrease in polyunsaturated fatty acids (PPAR ligand) (15). Hcy inhibits the formation of prostaglandins (67) and methylates the nucleic acid (70). It is also possible that in the condition of low PPAR activity, Hcy may increase oxidative stress by auto-oxidation, releasing superoxides by increasing NADPH oxidase (2, 72) and promoting the oxidative inflammatory condition. Hcy induces endocardial endothelial (EE) dysfunction (39) and instigates elastinolysis (28, 48), including arteriosclerosis (56), endothelial cell desquamation (53), thromboresistance (34), smooth muscle cell proliferation (57, 58), collagen synthesis (35, 58), oxidation of low-density lipoprotein (21), increased monocyte adhesion to the vessel wall (29), platelet aggregation (10), coagulation (46), blood rheology (14), and activation of plasminogen and metalloproteinase (20a, 64). Agonists of PPAR decrease the mRNA of plasminogen activator, increase the mRNA of plasminogen activator inhibitor (68), and decrease the metalloproteinase activity (37).

Although the treatment of PPAR-alpha agonists has been reported to ameliorate the Hcy-mediated endothelial dysfunction, the levels of Hcy were not decreased (3, 4, 12). In fact, some studies have suggested an increase in the levels of Hcy following fibrate therapy (3, 4, 12). This may suggest that fibrate and Hcy compete for the same binding site on PPAR-alpha and that fibrate displaces Hcy from PPAR-alpha . This would lead to an increase or no change in the plasma levels of Hcy, suggesting that fibrate ameliorates the effect of Hcy and not the metabolism. This is the paradox: drugs that repair endothelial dysfunction may increase Hcy accumulation. Therefore, the aim of this study was, is endothelial dysfunction by Hcy due to a decrease in PPAR function?


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Materials. D,L-Homocysteine, cysteine, fluorescamine, and ciprofibrate (CF) were obtained from Sigma Chemical. PGJ2 was obtained from Calbiochem. Fluorescamine-homocysteine (F-Hcy) and -cysteine (F-Cys) were prepared by using a 1:5 ratio of fluorescamine and subsequent gel-filtration chromatography to separate the complex from free fluorescamine and Hcy or Cys as described previously (58). F-Hcy has an absorbance at 334 nm and fluoresces at 480 nm. Free fluorescamine and Hcy do not absorb at 334 nm (58). To determine competitive binding among Hcy, fibrate, and PGJ2, we obtained an Hcy-cellulose affinity column from Sigma Chemical. The concentration of Hcy was estimated on the basis of colorimetric reaction with 5,5'-dithiobis(2-nitrobenzoic acid) and an increase in the absorbance at 340 nm, using a coefficient of 13,600 M-1 · cm-1 (64). The concentrations of CF and PGJ2 are based on the weight measurements. All reagents were dissolved in 50 mM Tris · HCl (pH 7.4). Fetal calf serum (FCS), minimum essential medium with Earle's salts (MEM), collagen, laminin-coated culture plates, and Hanks' balanced salt solution were obtained from Collaborative Research. Trypsin was obtained from GIBCO. Collagenase was from Worthington.

Microvessel EE cells. Mouse hearts were perfused with saline. The endocardium was carefully separated with a razor blade. EE cells were harvested by trypsin (0.1%) and collagenase digestion (200 units). This treatment detaches endothelial cells from basement membrane without disturbing the interstitium (60). These cells were further recovered to homogeneity into one major band (mostly endothelial) by centrifugation on Ficoll-Paque (Pharmacia, Biotech). EE cells were sorted from nonendothelial cells on the basis of their "cobblestone" characteristics and by the presence of factor VIII antigen. EE were maintained at 37°C in a humidified 5% CO2 atmosphere in MEM supplemented with 10% FCS, containing 2 mM L-glutamine and glucose. Isolated EE cells were cultured on collagen-coated plates in medium supplemented with 10% FCS and 2 mM glutamine. Cultures were routinely checked for the presence of mycoplasma (8), which has been shown to stimulate metalloproteinase (27).

Treatment of EE cells. The confluent cells were seeded at a density of 106 in a 35-mm disk in serum-free MEM for 24 h. The EE cells were treated with 0, 6, and 12 µM Hcy. To determine the role of PPAR agonists, we cotreated the EE cells with Hcy (12 µM) and CF (12 µM) or PGJ2 (12 µM) in serum-free medium. In controls, EE cells were treated with CF or PGJ2 (12 µM) alone. The medium was analyzed for metalloproteinase activity by gelatin zymography. The EE cell homogenates were analyzed for ICAM-1 and actin by Western blot analysis.

Cardiac nuclear extracts. To prepare cardiac nuclear protein extract, we harvested hearts from 10 wild-type mice (C57BL/6J). We selected C57BL/6J mice because this strain serves as the accepted background for genetically engineered mice. Also, the results from other laboratories have suggested extensive genetic homology between mice and humans (45). Nuclear proteins were isolated by a modification of the protocol of Dignam et al. (13). Briefly, after being washed with PBS, hearts were homogenized and centrifuged in buffer A (10 mM HEPES, pH 7.9, 1.5 mM MgCl2, 10 mM KCl, and 1 mM DTT). After centrifugation, the pellet was resuspended in buffer A containing 0.1% Triton X-100 by gentle pipetting. After incubation for 10 min at 4°C, the homogenate was centrifuged and the nuclear pellet was washed once with buffer A and resuspended in buffer C [10 mM HEPES, pH 7.9, 25% (vol/vol) glycerol, 0.4 M NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, and 1 mM DTT]. The suspension was incubated for 30 min at 4°C and then followed by centrifugation at 20,000 g for 10 min. The resulting supernatant was dialyzed in 50 mM Tris · HCl (pH 7.4) buffer. The extract was loaded onto the Hcy-cellulose affinity column, with prior activation with 5 ml of 0.5 M 2-mercaptoethanol and 0.1 M NaCl. The column was washed and preequilibrated with loading buffer (50 mM Tris · HCl, pH 7.4, 20 mM EDTA, and 0.1 M NaCl) as described previously (18). The bound protein was washed several times with loading buffer containing 0.5 M NaCl (18). After washing, the PPAR-alpha was specifically eluted by using 0.1 mM CF instead of NaCl in washing buffer in the elution step. To displace PPAR-gamma , we used the buffer containing 1 mM PGJ2. PGI2 was used as a control. The fractions eluted were measured for their absorption at 280 nm for protein by using a UV-based microplate reader (Molecular Devices). Loading buffer containing CF or PGJ2 was used as a reference. The concentration of PPAR-alpha and PPAR-gamma was estimated on the basis of an absorbance of 6.5 at 280 nm for 10 mg/ml (1%) protein.

PPAR-alpha and PPAR-gamma were purified by antibody affinity chromatography and immunoprecipitation with anti-PPAR-alpha or anti-PPAR-gamma conjugated with IgG-Sepharose. The antibody antigen was dissociated by using 10% SDS and gel filtration in 50 mM Tris · HCl (pH 7.4) buffer. The purity of PPAR was determined by a single band on SDS-PAGE silver stain and characterized by Western blot analysis.

SDS-PAGE and zymography. SDS-PAGE was performed on vertical slab gels according to a modification of the procedure of Laemmli (31) by using a 10% acrylamide/0.3% bis at pH 8.8 in the presence of reducing agents. All solutions contained 0.1% SDS. The gels were stained with Bio-Rad silver reagent. The actin band was identified by Western blot analysis with anti-beta -actin antibody (Sigma Chemical). The gelatin-gel zymography was performed on 1% gelatin SDS-PAGE under nonreducing conditions as described previously (59). The lytic band intensity was scanned and normalized with actin. The mean ± SE from at least 4 independent experiments is reported.

PPAR-alpha , PPAR-gamma , and ICAM-1 Western blots. The fractions eluted with CF and PGJ2 were concentrated. The concentrates were loaded onto a 10% SDS-PAGE under reducing condition as described previously (59). The protein was transferred to nitrocellulose membrane. The membrane was blocked with 5% fat-free milk. The blots were developed by using PPAR-alpha (Calbiochem), PPAR-gamma (Calbiochem), or ICAM-1 (Chemicon) antibodies, respectively. A secondary antibody alkaline phosphatase-conjugated detection system was used to identify the bands.

Fluorescence measurements. Excitation and emission spectra were recorded on a computer-controlled Spex Datamate spectrofluorometer as described previously (61, 62). The excitation and emission slits were adjusted for 1.25- and 2.50-nm band-pass width, respectively. Spectra were recorded at 1-nm intervals and corrected for baseline and instrument response. Samples were prepared and incubated for appropriate times before measurements were taken at 25°C in 0.3 × 0.3-cm microcells.

Treatment of fluorescence data. Although there is no tryptophan in a PPAR molecule, there are 12-15 tyrosine residues, and 6 or 7 of them are in the ligand-binding pocket of PPAR-alpha (25). The tyrosine fluoresces at 340 nm when excited at 295 nm (16, 32). Intrinsic fluorescence at 334 nm of PPAR-alpha and PPAR-gamma was recorded with excitation at 295 nm. The inner filter effects due to protein and ligands were corrected by using the equation F = Fobs antilog [(Aex + Aem)/2], where F is the corrected fluorescence intensity, Fobs is the observed intensity, Aex is the absorbance of the solution at the excitation wavelength, and Aem is the absorbance of the solution at the emission wavelength (32). The concentrations of bound and unbound PPAR-alpha or PPAR-gamma were related by the following equation: [PPAR]bound alpha  = [(FPPAR - Fsample)/(FPPAR - Fbuffer)], where Fsample is the fluorescence of reaction mixture and Fbuffer is the fluorescence background of the buffer alone. The binding constant and number of binding sites were estimated by using a nonlinear least-squares analysis: [(FPPAR - Fsample)/(FPPAR - Fbuffer)] = [Hcy/PPAR] × 1/[n(1 - alpha )] - (Kd/PPAR × n), where n is the number of binding sites, alpha  is the fraction of bound PPAR, and Kd is the dissociation constant of the primary binding site.

Statistical analysis. Values are given as means ± SE from n = 4 in each group. Differences between groups were evaluated by using ANOVA followed by the Bonferroni post hoc test (55), focusing on the effects of Hcy (EE cells to Hcy-treated EE cells) and treatment (EE + Hcy-treated cells compared with EE + Hcy cells cotreated with CF or PGJ2). P < 0.05 was considered significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Is EE activation by Hcy due to attenuation of PPAR? In response to Hcy loading, EE cells synthesize enhanced levels of matrix metalloproteinase (MMP)-2 at 72 kDa. The cotreatment of EE cells with Hcy plus CF or PGJ2 ameliorates the MMP induction (Fig. 1). There is an Hcy dose-dependent increase in ICAM-1 expression in EE cells. This increase is abrogated by cotreatment with CF or PGJ2 (Fig. 2). These results suggest a role of PPAR-alpha and PPAR-gamma in Hcy-mediated EE activation.


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Fig. 1.   A: confluent endocardial endothelial (EE) cells (3-4 passages) were cultured in serum-free medium for 24 h. The medium was analyzed for matrix metalloproteinase (MMP) activity by using gelatin-gel zymography. The levels of EE cell actin were measured by Western analysis. Lane 1, EE cells alone; lane 2, EE cells treated with ciprofibrate (CF; 12 µM); lanes 3 and 4, EE cells treated with homocysteine (Hcy; 12 µM); lane 5, EE cells treated with Hcy + CF; lane 6, EE cells treated with Hcy + 15-deoxy-Delta 12,14-prostaglandin J2 (PGJ2; 12 µM). B: histographic presentation of metalloproteinase lytic activity [in arbitrary units (AU)] normalized with actin. Values are means ± SE from 4 independent EE cell culture experiments. *P < 0.05 compared with EE alone. **P < 0.05 compared with Hcy.



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Fig. 2.   A: confluent EE cells (3-4 passages) were cultured in serum-free medium for 24 h. The medium was analyzed for ICAM-1 and actin by Western blotting. Lane 1, EE cells alone; lane 2, EE cells treated with Hcy (6 µM); lane 3, EE cells treated with Hcy (12 µM). EE cells treated with 12 µM Hcy were cotreated with 12 µM CF (lane 4) or PGJ2 (lane 5). B: histographic presentation of ICAM-1 levels (in AU) normalized with actin. Values are means ± SE from 4 independent EE cell culture experiments. *P < 0.05 compared with EE alone. **P < 0.05 compared with Hcy.

Elution of protein by CF and PGJ2 from the Hcy-cellulose column. After extensive washing with NaCl, protein bound to the column was eluted by the addition of CF or PGJ2 to the elution buffer. The changes in the optical density at 280 nm of eluted fractions suggest that a protein bound to Hcy in the column is eluted by CF. Another protein was eluted by PGJ2 (Fig. 3B). The SDS-PAGE silver stain analysis of the fractions, isolated from the Hcy-cellulose column, reveal that a number of protein bands appear in the eluate by CF and PGJ2. This finding suggests that CF and PGJ2 elute several proteins from the Hcy-column. However, to determine whether PPAR-alpha and PPAR-gamma were present in these fractions, Western blot analysis was performed on fractions eluted by CF and PGJ2. The results suggest that CF eluted PPAR-alpha (Fig. 3B, left inset) and PGJ2 eluted PPAR-gamma (Fig. 3B, right inset). There was no PPAR in the elution by PGI2 (Fig. 3A).


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Fig. 3.   A: separation of protein bound to Hcy-cellulose column. Cardiac nuclear extracts were prepared from 10 mice. The extract was equilibrated with buffer (50 mM Tris · HCl, pH 7.2, 20 mM EDTA, and 0.1 M NaCl) and loaded onto the column, preequilibrated with the same buffer. The bound proteins were washed with 0.5 M NaCl. A: after extensive washing, PGI2 (0.1 mM) was added to elution buffer. B: CF (0.1 mM, from 1 M stock solution in ethanol) was added to elution buffer. The fractions were collected. After absorption reached background, 0.1 mM PGJ2 was added. The fractions were collected. The optical density at 280 nm (OD280 nm) was measured for each fraction by using loading buffer as a reference. Left inset: Western blot analysis of fractions eluted by CF using peroxisome proliferator-activated receptor (PPAR)-alpha antibody. Arrow indicates PPAR-alpha reactive band. Right inset: Western blot analysis of fractions eluted by PGJ2 from the Hcy-column using anti-PPAR-gamma antibody. Arrow indicates PPAR-gamma reactive band. The 10% SDS-PAGE was performed under reducing conditions and transferred to nitrocellulose membrane. The protein was blotted. An alkaline phosphatase-conjugated system was used to detect the labeling.

Purification of PPAR-alpha and PPAR-gamma . PPAR-alpha and PPAR-gamma were purified by antibody affinity chromatography. The results suggest a single subunit of PPAR of ~66 kDa molecular mass in range. The molecular mass of PPAR-alpha was slightly higher than that of PPAR-gamma (Fig. 4).


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Fig. 4.   Isolation and characterization of PPAR-alpha and PPAR-gamma from cardiac nuclear extracts by antibody affinity-column chromatography. Serial blots were analyzed for PPAR-alpha and PPAR-gamma , lanes 1 and 2, respectively. Left, SDS-PAGE silver stain of purified fractions; middle, Western blot analysis using anti-PPAR-gamma antibody; right, Western blot analysis using anti-body to PPAR-alpha .

Hcy induces conformational changes in PPAR. The immunopurified fractions of PPAR-alpha and PPAR-gamma demonstrate fluorescence with a maximum at ~334 nm. The peak maximum at ~334 nm was shifted to ~340 nm after the addition of Hcy (Fig. 5). These results suggest that Hcy binds and induces conformational changes in PPAR and that these changes are associated with the hydrophilic environment around the binding site in PPAR. With the use of nonlinear least-squares analysis, the dose-dependent increase in PPAR-alpha fluorescence elicits 2.45 ± 0.57 binding sites of Hcy per PPAR-alpha molecule. The Kd of the primary binding site is 0.32 ± 0.06 µM. There is no significant binding to Cys. These results suggest that Hcy binds to PPAR at the primary site with affinity in the submicromolar range.


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Fig. 5.   A: shift in PPAR-alpha fluorescence at 340 nm when excited at 295 nm upon binding to Hcy. The fluorescence of isolated PPAR-alpha as detected by antibody affinity chromatography was recorded by spectrofluorometer. Hcy was added to the aliquot containing PPAR-alpha . The baseline fluorescence of Hcy + buffer was recorded. B: fraction of change in PPAR-alpha (1 µM) fluorescence upon binding to Hcy or cysteine (Cys). Each point is an average of 4 independent data points. The reference containing Hcy + buffer was subtracted from each experimental point. The curves are the best fits to nonlinear least-squares analysis.

Displacement of Hcy by CF from PPAR-alpha and not by PGJ2. To determine whether Hcy can be displaced by PPAR-alpha agonist, we prepared F-Hcy. Because F-Hcy absorbs at 340 nm (Fig. 6), the increase in PPAR-alpha fluorescence near 334 nm is quenched by the binding of F-Hcy in the proximity of the binding site in PPAR. The dose-dependent quenching of the fluorescence of PPAR-alpha is shown in Fig. 7. There was no significant quenching by F-Cys. The quenching was abrogated by CF (Fig. 7A) but not by PGJ2 (Fig. 7B). These results suggest that Hcy competes with CF on PPAR-alpha , but not with PGJ2 (a PPAR-gamma agonist).


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Fig. 6.   Spectral overlap of PPAR-alpha fluorescence and fluorescamine (F)-Hcy absorption. Samples were incubated in 50 mM Tris · HCl (pH 7.4) at 37°C before their spectra were recorded. PPAR-alpha (1 µM) fluorescence was recorded with excitation at 295 nm, and band-pass slits were set at 1.25 and 2.5 nm for excitation and emission, respectively. The absorption of F-Hcy (1 µM) was recorded. All spectra were corrected for buffer baseline and instrument response. The y-axis is offset to overlap the 2 spectra. The relative fluorescence intensity for PPAR-alpha and absorption for F-Hcy are reported.



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Fig. 7.   A: fractional changes in PPAR-alpha (1 µM) fluorescence at 340 nm when excited at 295 nm by binding to and quenching by F-Hcy or F-Cys. The reference containing F-Hcy or F-Cys + buffer was subtracted from each experimental point. The curves are the best fits to nonlinear least-squares analysis. Arrows represent the addition of difference doses of CF to the cuvette containing F-Hcy/PPAR-alpha . The reference fluorescence due to CF and F-Hcy was subtracted from the fluorescence of F-Hcy/PPAR-alpha . Each point is an average of 4 independent data points. B: fractional changes in PPAR-alpha (1 µM) fluorescence upon binding to and quenching by F-Hcy. The reference containing F-Hcy + buffer was subtracted from each experimental point. The curves are the best fits to nonlinear least-squares analysis. Arrows represent the addition of different doses of PGJ2 to the cuvette containing F-Hcy/PPAR-alpha . The reference fluorescence due to PGJ2 and F-Hcy was subtracted from the fluorescence of F-Hcy/PPAR-alpha . Each point is an average of 4 independent data points.

Displacement of Hcy by PGJ2 from PPAR-gamma and not by CF. To determine whether homocysteine can be displaced by PPAR-gamma agonist, we carried out the titration of F-Hcy/PPAR-gamma . The increase in PPAR-gamma fluorescence at 334 nm is quenched by the binding of F-Hcy in the proximity of the binding site in PPAR. The dose-dependent quenching of the fluorescence of PPAR-gamma is shown in Fig. 8. The quenching was abrogated by PGJ2 (Fig. 8A) but not by CF (Fig. 8B). These results suggest that homocysteine competes with PGJ2 on PPAR-gamma , but not with CF (a PPAR-alpha agonist).


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Fig. 8.   A: fractional changes in PPAR-gamma (1 µM) fluorescence at 340 nm when excited at 295 nm by binding to and quenching by F-Hcy. The reference containing F-Hcy + buffer was subtracted from each experimental point. The curves are the best fits to a nonlinear least-squares analysis. Arrows represent the addition of difference doses of PGJ2 to the cuvette containing F-Hcy/PPAR-gamma . The reference fluorescence due to PGJ2 and F-Hcy was subtracted from the fluorescence of F-Hcy/PPAR-gamma . Each point is an average of 4 independent data points. B: fractional changes in PPAR-gamma (1 µM) fluorescence upon binding to and quenching by F-Hcy. The reference containing F-Hcy + buffer was subtracted from each experimental point. The curves are the best fits to nonlinear least-squares analysis. Arrows represent the addition of different doses of CF to the cuvette containing F-Hcy/PPAR-gamma . The reference fluorescence due to CF and F-Hcy was subtracted from the fluorescence of F-Hcy/PPAR-gamma . Each point is an average of 4 independent data points.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Lowering of the levels of Hcy demonstrates reduction in ischemic events (50). The plasma Hcy levels are positively related to blood pressure in the Hordaland Homocysteine Study of 16,000 people 40-67 yr old (43). Similarly, Malinow and coworkers (36) found that hypertensive men had higher Hcy levels than nonhypertensive men. Sutton-Tyrrell and colleagues (54) also found a significant association of Hcy levels with systolic hypertension. Elevation of Hcy is associated with an increase in blood pressure in rats (39) and minipigs (48). The agonist of PPAR decreases systolic blood pressure (17, 52). Hcy induces cardiac hypertrophy in rats (39). Ventricular pressure-overload studies in mice have demonstrated reduced expression of PPAR-alpha during cardiac hypertrophy (1). Peroxisome proliferators ameliorate cardiac hypertrophy (69). An inverse relationship between oxidative microvessel endothelial cell density and cardiac hypertrophy has been suggested (47). The role of Hcy in vascular endothelial dysfunction has been studied extensively; however, little attention has been given to the role of Hcy in capillary EE. In an acute study we have demonstrated that Hcy impairs EE by decreasing the bioavailability of nitric oxide (63). In a chronic study of 4-wk hyperhomocysteinemia, Ungvari et al. (65) demonstrated that reduced activity of nitric oxide in arterioles may contribute to the microvascular impairment by Hcy. We have demonstrated that 12-wk hyperhomocysteinemia induces apoptosis in EE (39) and instigates EE dysfunction (38). Hcy has an intracellular cytosolic redox receptor in vascular cells (58), and nicotinamide, an inhibitor of poly(ADP-ribose)synthetase, an enzyme that can be activated by peroxynitrite and oxidants (5, 9), reverses Hcy-mediated endothelial dysfunction (40). Here we suggest that Hcy activates EE by antagonizing PPAR-alpha and PPAR-gamma .

To determine the source of PPAR and Hcy interaction in the heart, we cultured EE cells in the presence and absence of Hcy. Hcy attenuates endothelial function (40), and PPAR agonists ameliorate the endothelial dysfunction (17). Also, PPAR agonists inhibits the metalloproteinase activation in macrophage (37). It was unclear whether PPAR agonists ameliorate Hcy-mediated endothelial activation. ICAM-1 has been used as a marker of endothelial cell activation (30). Our data demonstrate that Hcy activates EE cells and increases metalloproteinase activity and ICAM-1 expression in response to antagonize PPAR-alpha and PPAR-gamma (Figs. 1 and 2).

The shift in PPAR fluorescence by Hcy elicits physical binding between PPAR and Hcy. The shift suggests that tyrosines are near the Hcy binding site in PPAR (Fig. 5). Previous studies demonstrate a binding constant between PPAR-alpha and fibrate in the micromolar range (42). Our results suggest an ~10-fold stronger affinity of Hcy to PPAR-alpha than to fibrate. We also observed more than one binding site of Hcy to PPAR-alpha . These results may suggest that Hcy has two different sites on PPAR-alpha and two different functions. To confirm the Hcy binding to PPAR-alpha by displacement titration, we used F-Hcy (58). The interaction between PPAR-alpha and F-Hcy suggests that CF competes specifically with F-Hcy by binding to PPAR-alpha (Fig. 7). The Western blot analysis of cardiac fractions eluted by PGJ2 (Fig. 2C) and displacement of F-Hcy/PPAR-gamma complex with PGJ2 reveal the significant amount of PPAR-gamma expression in the heart (Fig. 8). These results suggest that there are PPAR-alpha and -gamma in the heart and that Hcy competes with PPAR agonists for binding to these receptors.


    ACKNOWLEDGEMENTS

This work was supported in part by National Institutes of Health Grants GM-48595 and HL-71010 and by the Kidney Care Foundation.


    FOOTNOTES

Address for reprint requests and other correspondence: S. C. Tyagi, The Univ. of Mississippi Medical Center, Dept. of Physiology & Biophysics, 2500 North State St., Jackson, MS 39216-4505 (E-mail: styagi{at}physiology.umsmed.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.

10.1152/ajpcell.00152.2002

Received 5 April 2002; accepted in final form 23 May 2002.


    REFERENCES
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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