Department of Physiology and Biophysics, University of Mississippi Medical Center, Jackson, Mississippi 39216
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ABSTRACT |
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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- agonist) or 15-deoxy-
12,14-prostaglandin
J2 (PGJ2; PPAR-
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 PPAR
and -
, 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-
fluorescence quenching was
abrogated by the addition of CF but not by PGJ2. PPAR-
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-
and -
,
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
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INTRODUCTION |
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HOMOCYSTEINE (Hcy)
induces nuclear factor-B (NF-
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-
is expressed
in adipose, liver, heart, muscle, and kidney (41).
PPAR-
is ubiquitous throughout the body. PPAR-
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-
(11, 71), AA binds both PPAR-
and PPAR-
. On the other hand, LB4 binds preferentially PPAR-
(11). Among most of the
fibrates, ciprofibrate has the most potent effect in ameliorating
endothelial dysfunction (17). The
15-deoxy-
12,14-prostaglandin J2
(PGJ2) is a potent agonist of PPAR-
(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- 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-
and that fibrate displaces Hcy from PPAR-
. 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?
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MATERIALS AND METHODS |
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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 M1 · 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- was specifically eluted by using 0.1 mM CF
instead of NaCl in washing buffer in the elution step. To displace
PPAR-
, 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-
and PPAR-
was estimated on the basis of an absorbance of 6.5 at 280 nm for 10 mg/ml (1%) protein.
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--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-, PPAR-
, 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-
(Calbiochem), PPAR-
(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- (25). The tyrosine
fluoresces at 340 nm when excited at 295 nm (16, 32).
Intrinsic fluorescence at 334 nm of PPAR-
and PPAR-
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-
or PPAR-
were related
by the following equation: [PPAR]bound =
= [(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
)]
(Kd/PPAR × n), where
n is the number of binding sites,
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.
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RESULTS |
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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- and PPAR-
in Hcy-mediated EE activation.
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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- and PPAR-
were present in these fractions, Western
blot analysis was performed on fractions eluted by CF and
PGJ2. The results suggest that CF eluted PPAR-
(Fig.
3B, left inset) and PGJ2 eluted PPAR-
(Fig. 3B, right inset). There was no
PPAR in the elution by PGI2 (Fig. 3A).
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Purification of PPAR- and PPAR-
.
PPAR-
and PPAR-
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-
was
slightly higher than that of PPAR-
(Fig.
4).
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Hcy induces conformational changes in PPAR.
The immunopurified fractions of PPAR- and PPAR-
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-
fluorescence elicits 2.45 ± 0.57 binding sites of Hcy per PPAR-
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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Displacement of Hcy by CF from PPAR- and not by
PGJ2.
To determine whether Hcy can be displaced by PPAR-
agonist, we
prepared F-Hcy. Because F-Hcy absorbs at 340 nm (Fig.
6), the increase in PPAR-
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-
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-
, but not
with PGJ2 (a PPAR-
agonist).
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Displacement of Hcy by PGJ2 from PPAR- and not by
CF.
To determine whether homocysteine can be displaced by PPAR-
agonist,
we carried out the titration of F-Hcy/PPAR-
. The increase in
PPAR-
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-
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-
, but not with CF (a PPAR-
agonist).
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DISCUSSION |
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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- 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-
and PPAR-
.
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-
and PPAR-
(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- and fibrate in the micromolar range
(42). Our results suggest an ~10-fold stronger affinity of Hcy to PPAR-
than to fibrate. We also observed more than one binding site of Hcy to PPAR-
. These results may suggest that Hcy has
two different sites on PPAR-
and two different functions. To confirm
the Hcy binding to PPAR-
by displacement titration, we used F-Hcy
(58). The interaction between PPAR-
and F-Hcy suggests
that CF competes specifically with F-Hcy by binding to PPAR-
(Fig.
7). The Western blot analysis of cardiac fractions eluted by
PGJ2 (Fig. 2C) and displacement of
F-Hcy/PPAR-
complex with PGJ2 reveal the significant
amount of PPAR-
expression in the heart (Fig. 8). These results
suggest that there are PPAR-
and -
in the heart and that Hcy
competes with PPAR agonists for binding to these receptors.
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ACKNOWLEDGEMENTS |
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This work was supported in part by National Institutes of Health Grants GM-48595 and HL-71010 and by the Kidney Care Foundation.
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FOOTNOTES |
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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.
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