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Am J Physiol Cell Physiol 290: C1399-C1410, 2006. First published December 21, 2005; doi:10.1152/ajpcell.00386.2005
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VASCULAR BIOLOGY

Glutamate induces oxidative stress and apoptosis in cerebral vascular endothelial cells: contributions of HO-1 and HO-2 to cytoprotection

Helena Parfenova,1 Shyamali Basuroy,1 Sujoy Bhattacharya,1 Dilyara Tcheranova,1 Yan Qu,2 Raymond F. Regan,2 and Charles W. Leffler1

1Laboratory for Research in Neonatal Physiology, Department of Physiology, Vascular Biology Center, University of Tennessee Health Science Center, Memphis, Tennessee; and 2Department of Emergency Medicine, Thomas Jefferson University, Philadelphia, Pennsylvania

Submitted 1 August 2005 ; accepted in final form 19 December 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In cerebral circulation, epileptic seizures associated with excessive release of the excitatory neurotransmitter glutamate cause endothelial injury. Heme oxygenase (HO), which metabolizes heme to a vasodilator, carbon monoxide (CO), and antioxidants, biliverdin/bilirubin, is highly expressed in cerebral microvessels as a constitutive isoform, HO-2, whereas the inducible form, HO-1, is not detectable. Using cerebral vascular endothelial cells from newborn pigs and HO-2-knockout mice, we addressed the hypotheses that 1) glutamate induces oxidative stress-related endothelial death by apoptosis, and 2) HO-1 and HO-2 are protective against glutamate cytotoxicity. In cerebral endothelial cells, glutamate (0.1–2.0 mM) increased formation of reactive oxygen species, including superoxide radicals, and induced major keystone events of apoptosis, such as NF-{kappa}B nuclear translocation, caspase-3 activation, DNA fragmentation, and cell detachment. Glutamate-induced apoptosis was greatly exacerbated in HO-2 gene-deleted murine cerebrovascular endothelial cells and in porcine cells with pharmacologically inhibited HO-2 activity. Glutamate toxicity was prevented by superoxide dismutase, suggesting apoptotic changes are oxidative stress related. When HO-1 was pharmacologically upregulated by cobalt protoporphyrin, apoptotic effects of glutamate in cerebral endothelial cells were completely prevented. Glutamate-induced reactive oxygen species production and apoptosis were blocked by a CO-releasing compound, CORM-A1 (50 µM), and by bilirubin (1 µM), consistent with the antioxidant and cytoprotective roles of the end products of HO activity. We conclude that both HO-1 and HO-2 have anti-apoptotic effects against oxidative stress-related glutamate toxicity in cerebral vascular endothelium. Although HO-1, when induced, provides powerful protection, HO-2 is an essential endogenous anti-apoptotic factor against glutamate toxicity in the cerebral vascular endothelium.

endothelium; carbon monoxide; bilirubin; injury; reactive oxygen species; heme oxygenase

GLUTAMATE IS THE MAJOR EXCITATORY neurotransmitter in the central nervous system. In cerebral circulation, glutamate is a vasodilator that contributes to increases in cerebral blood flow in response to seizures, matching metabolic demands of active neurons, and, therefore, preventing subsequent neuronal injury (28). The intracellular concentration of glutamate in the brain is in the 10–3 M range, whereas the extracellular concentration is maintained at 10–6 to 10–5 M (13, 28). Under certain pathological conditions such as ischemia, hypoxia, and brain injury, the overall glutamate concentration in the brain can be elevated up to 50-fold, and local concentrations can be much higher (13). At excitotoxic concentrations (10–3 M range), glutamate triggers neuronal damage and death (14, 15). Glutamate-induced brain cell injury is involved in the pathogenesis of neurodegenerative disorders, stroke, seizures, and epilepsy (10, 13, 20, 28, 33).

Very little is known about the damaging effects of glutamate in the cerebral vasculature. Glutamatergic seizures increase the formation of reactive oxygen species (ROS) by the cerebral cortex (3) and cause sustained cerebral vascular endothelial injury (37). Brain endothelial cells express ionotropic and metabotropic glutamate receptors and may respond directly to the neurotransmitter (12, 38, 48). Receptor-mediated effects of glutamate in the cerebral endothelium include increased oxidative stress and changes in blood-brain barrier (5, 12, 47, 48). Overall, these findings suggest that cerebral vascular endothelium is among the targets for glutamate cytotoxicity in the brain.

Oxidative stress injury is a leading mechanism of glutamate neurotoxicity. In neurons, activation of ionotropic glutamate receptors results in massive calcium entry, calcium overload in mitochondria, energy failure, excessive ROS formation, and, eventually, cell death (16, 20). Heme oxygenase (HO) is a component of the endogenous cell defense against oxidative stress (24, 36). HO degrades heme to biliverdin, which is then reduced to bilirubin by biliverdin reductase. Overall, HO-catalyzed reactions reduce the amount of pro-oxidant heme and increase bilirubin, which scavenges ROS by the redox cycling with biliverdin (15, 30). Carbon monoxide (CO), an end product of heme catabolism via the HO pathway, is a vasodilator messenger in cerebral circulation (24). Although CO irreversibly binds to heme and can be lethal to animals by blocking oxygen transport via hemoglobin, low concentrations of CO may have anti-apoptotic effects in vascular endothelial and smooth muscle cells (6, 7, 26).

Two major HO isoforms, inducible (HO-1) and constitutive (HO-2), the products of separate genes, catalyze heme degradation by similar mechanisms (24, 36). However, significant differences in gene regulation, molecular structure, and tissue distribution suggest that HO isoforms may have distinct cellular functions. HO-1 is induced by a variety of cell- and species-dependent stress factors, including oxidative stress (17, 36). HO-1 expression is regulated at the level of gene transcription via multiple stress- and antioxidant response regulatory elements in the promoter region of the HO-1 gene (1, 23, 34). Cytoprotective effects of HO-1 have been demonstrated in neurons (11, 46) and endothelial cells (5, 6, 36, 42, 54). HO-1 is not detectable in cerebral vasculature under physiological conditions and is not induced by epileptic seizures (24, 37). However, pharmacological induction of HO-1 in cerebral circulation provided complete protection against seizure-related cerebral vascular dysfunction (38).

Little is known about the role of HO-2 in oxidative stress-related vascular injury. HO-2 is essential in neuronal functions (14, 15), in regulation of cerebral blood flow (9, 24), and in preventing seizure-related cerebral vascular injury (9, 37). Overall, these data indicate that HO-2 may be an important player in the glutamate-related stress defense mechanism in the cerebral circulation.

The present study addresses the hypotheses that 1) glutamate induces oxidative stress-related endothelial cell death, and 2) HO-1 and HO-2 are protective against glutamate-induced endothelial cytotoxicity. We investigated the effects of glutamate on cell survival in primary cultures of cerebral vascular endothelial cells from newborn pigs that constitutively express high levels of HO-2, and from adult HO-2-knockout (KO) mice (HO2KO). To compare the roles of HO-1 and HO-2 isoforms in cell survival, we pharmacologically induced HO-1 overexpression in porcine cerebral vascular endothelial cells. In addition, we tested anti-apoptotic and anti-oxidant capacities of the end products of HO-catalyzed heme degradation, bilirubin, and CO against glutamate-induced endothelial cell death.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Protocols involving piglets were approved by the Animal Care and Use Committee at the University of Tennessee Health Science Center, and those protocols involving mice were approved by the Institutional Animal Care and Use Committee at Thomas Jefferson University. All procedures were done using aseptic techniques.

HO2KO mice. HO2KO mice used in this study were descendants of those produced by Poss et al. (41) using the HO-2-targeting construct that replaced a 3.4-kb region corresponding to rat HO-2 exons 4 and 5 with a pgk-neo cassette and were cross-bred with wild-type (WT) C57BL/6 x 129/Sv mice. The genotype was identified by PCR using the following primers: HO-2A: GAG TTG CTG GCT TGG CTT ATA G; HO-2B: TTC CGG TGT AGC TCC GTG GGG; Neo 1 insert: GCT TGG GTG GAG AGG CTA TTC; Neo 2 insert: CAA GGT GAG ATG ACA GGA GAT C (45). All breeding mice were obtained by pairing heterozygotes, and the offspring were genotyped to obtain HO-2+/+ and HO-2–/– breeders. Both WT and KO mice were obtained from this colony; therefore, the genetic backgrounds of HO2KO and WT mice were identical, other than a lack of the HO-2 gene. Mice were bred and then euthanized at Thomas Jefferson University, and the brains were sent to the University of Tennessee Health Science Center, where endothelial cells were isolated and grown.

Cerebrovascular endothelial cell primary cultures. Brain cortex was obtained from ketamine/acepromazine-anesthetized newborn pigs (1–5 days of age) or from isoflurane-anesthetized HO-2-KO or -WT mice (3–4 mo of age). The tissue was gently homogenized in medium 199 (1:10) with the use of a glass homogenizer and a loose-fitting pestle. Cerebral microvessels were collected by filtration of the homogenate through 300- and 60-µm nylon mesh screens consecutively. Microvessels were treated with collagenase dispase (2 mg/ml for 2 h at 37°C), and dissociated cerebral microvascular endothelial cells (CMVEC) were separated on a Percoll density gradient (38). CMVEC were plated on Matrigel-coated plates (3 x 104 cells/well) and cultured at 37°C (5% CO2-95% air) in DMEM with 20% fetal bovine serum (FBS), 30 µg/ml endothelial cell growth supplement, 1 U/ml heparin, and an antibiotic/antimycotic mixture for 5–6 days until confluent. Endothelial cells identified by immunostaining for von Willebrand factor accounted for at least 95% of the total cell population. All experiments were performed with confluent quiescent cells in primary cultures (piglet CMVEC) or in first-passage cultures (murine CMVEC). To achieve quiescence, CMVEC were exposed to a serum-depleted medium (0.1% FBS) for 15–20 h before the experiment.

Detection of oxidant generation. To assess intracellular ROS, two independent oxidant-sensitive probes were used (Molecular Probes, Eugene, OR). Dihydrorhodamine 123 (DHR123) is a cell-permeable, oxidant-sensitive, nonfluorescent probe that is oxidized to fluorescent rhodamine 123 (Rh123) by a variety of intracellular ROS species, especially H2O2 and peroxynitrite (21, 22). Rh123 is preferentially located to the mitochondria. Dihydroethidium (DHE) is generally used specifically to assess superoxide (O2–·) production (21, 22, 40). Cytosolic DHE exhibits blue fluorescence; once it is oxidized by O2–· to ethidium and oxyethidium, it intercalates with DNA in the nucleus and emits red fluorescence.

To visualize ROS formation in live cells using fluorescence microscopy, CMVEC were loaded with 50 µM DHR123 for 30 min at 37°C, and further incubated with either peroxynitrite (0.1–1.0 mM), H2O2 (0.1–2.0 mM), arachidonic acid (10–200 µM), or glutamate (0.1–2 mM) for an additional 30 min. Cells were rapidly rinsed three times with Dulbecco’s phosphate-buffered saline (DPBS) to remove the free probe. The live cells were immediately observed under a Nikon Diaphot microscope with a fluorescein filter coupled to a MacQuadra 950 computer system with a Power Mac 601 processor.

For measurement of ROS generation in response to glutamate and oxidants, CMVEC were loaded with 50 µM DHR123 for 20 min at 37°C, and the incubation was continued in the presence of glutamate (0.1–2 mM), peroxynitrite (0.1–1.0 mM), H2O2 (0.1–2.0 mM), or arachidonic acid (10–200 µM) for an additional 20 min. CMVEC were washed twice with cold DPBS to remove the free probe, scraped in 1 ml of ice-cold DPBS, and placed into microtubes. The cells were pelleted at 3,000 g for 10 min at 4°C and resuspended in 0.5 ml of ice-cold DPBS. The cell suspensions were sonicated on ice for 15 s, clarified by centrifugation, and 100-µl aliquots were dispensed into Falcon 96-well black microplates (Becton Dickinson, Franklin Lakes, NJ) in quadruplicate. Probe-free cell sonicates were used for blanks. Fluorescence was measured with a Fusion {alpha}HT Universal Microplate Analyzer (Packard Instruments, Meriden, CT) at excitation and emission wavelengths of 470 and 525 nm, respectively (21, 22, 40).

To detect the effects of heme metabolites on ROS formation under glutamate-induced apoptosis conditions, quiescent CMVEC were incubated with 1 mM glutamate in serum- and glutamine-free DMEM for 3 h at 37°C. DHR123 (50 µM) or DHE (20 µM) were added to the incubation media for the last 20 min of incubation. To detect overall glutamate-induced oxidative stress, both attached and floating cells were collected. CMVEC were mechanically dislodged, diluted with ice-cold DPBS, and pelleted at 3,000 g for 10 min at 4°C. The process was repeated twice to remove free probes. The cell pellet was disrupted by sonication in 100 µl of ice-cold DPBS, and cell debris were removed by centrifugation. Supernatant (total cell lysate) was transferred to Falcon 96-well black microplates in duplicate, and fluorescence was measured with a Fusion {alpha}HT Universal Microplate Analyzer (Packard Instruments) using excitation and emission wavelengths of 470/525 nm for Rh123 and 490/605 nm for ethidium (21, 22, 40).

Induction of HO-1 expression in porcine CMVEC. Cobalt protoporphyrin (CoPP) is a potent HO-1-inducing agent in cerebral microcirculation (37). CMVEC were treated with 20 µM CoPP in DMEM-0.1% FBS for 3 h. After CoPP was removed, the cells were incubated overnight in fresh DMEM-0.1% FBS. Under these conditions, HO-1 expression was upregulated ~10-fold.

Glutamate-induced apoptosis in CMVEC. CMVEC were incubated with 1–2 mM glutamate in glutamine-free DMEM-0% FBS for 3 h at 37°C in the absence or presence of tin protoporphyrin (SnPP), antioxidants (SOD and bilirubin), or CORM-A1. During the experiment, the cells were protected from light to avoid heme pigment photodegradation. All solutions were freshly prepared. The bilirubin/albumin mixture (molar ratio, 1:5) was prepared as described by Dore et al. (15). A boron-based carbonylating agent, sodium boranocarbonate (CORM-A1), spontaneously and slowly releases CO in neutral and acidified aqueous solutions (pH ≤7.4) and elicits pharmacological effects of CO (2, 31). CORM-A1 was diluted in Krebs buffer (pH 7.4) immediately before the experiment.

Caspase-3 activity detection by Western blot analysis. Caspase-3 activation was detected by formation of its proteolytic fragments (17 and 19 kDa) using highly selective polyclonal antibodies against cleaved caspase-3-Asp175 (Cell Signaling, Beverly, MA). Total proteins from control or treated CMVEC (10–20 µg protein/lane) were separated by 12% SDS-PAGE and transferred onto Hybond-P membranes (Amersham Biosciences, Piscataway, NJ). The membranes were blocked for 1 h at room temperature with 5% nonfat dry milk in 0.1% Tween 20-Tris-buffered saline and probed with cleaved caspase-3-Asp175 antibodies (1:1,000 dilution), followed by peroxidase-conjugated donkey anti-rabbit IgG (dilution 1:20,000; Sigma, St. Louis, MO). The antigen expression was normalized to actin detected by reprobing the membranes with anti-actin monoclonal antibodies (dilution 1:10,000, Roche Molecular Biochemicals, Indianapolis, IN). Proteins were visualized with Western Lightning Chemiluminescence Reagent Plus (Perkin Elmer Life Sciences, Boston, MA) and quantified by digital densitometry using NIH ImageJ version 1.33 software.

DNA fragmentation. DNA fragmentation was detected by accumulation of mono- and oligonucleosomes in the CMVEC nuclei-free cytoplasmic fraction by sandwich enzyme immunoassay using antibodies against DNA and histones (Cell Death Detection ELISA Plus Kit; Roche Diagnostics, Mannheim, Germany) as described elsewhere (4). Briefly, control or treated cells on 12-well culture plates were washed twice with DPBS, lysed, and centrifuged to remove the nuclei. An aliquot of the nuclei-free supernatant was transferred to streptavidin-coated wells and incubated with anti-histone-biotin antibody and anti-DNA peroxidase-conjugated antibody for 2 h at room temperature. After incubation, the wells were washed three times, developed with 2,2'-azino-di(3-ethylbenzthiazolin-sulfonate), and the absorbance was read at 405 nm.

Cell viability. Cell viability was estimated from the number of attached (live) and floating (dead) cells. Attached cells were quantified by the amount of protein detected using a protein assay kit (Pierce Biotechnology, Rockford, IL). Floating cells were pelleted and counted using a microscope counting chamber.

Cytotoxicity assay. Cytotoxicity was estimated as an increase in membrane permeability (cytolysis) detected by release of a cytosolic component, adenylate kinase (AK), into the medium. AK activity in the medium was detected using the ToxiLight Cytotoxicity Assay Kit (Cambrex Bio Science, Rockland, ME) by AK-catalyzed ADP-ATP conversion; ATP was detected using the luciferase-based bioluminescence method according to the manufacturer’s instructions. Luminescence was measured with a Fusion {alpha}HT Universal Microplate Analyzer (Packard Instruments).

Western blot analysis for HO-1 and HO-2. Cells were extracted with 1% Triton X-100 in 50 mM Tris·HCl buffer, pH 7.4, containing protease inhibitors. Proteins (20–50 µg/lane) were separated by 12% SDS-PAGE and immunoblotted as described above. Hybond-P transfer membranes were probed with a polyclonal antibody to a human HO-2 synthetic peptide (1:5,000 dilution; SPA 897 from StressGen Biotechnologies, Victoria, BC, Canada), or a polyclonal antibody to a human HO-1 peptide (1:5,000 dilution; StressGen).

Immunofluorescence detection of HO-2 and NF-{kappa}B p65 in murine CMVEC. Confluent, quiescent murine CMVEC were fixed with 3.7% paraformaldehyde in DPBS (pH 7.4; 20 min, room temperature) and permeabilized by 0.1% Triton X-100-DPBS (20 min, room temperature). The nonspecific binding sites were blocked by 5% BSA-DPBS (1 h at room temperature). The cells were incubated with HO-2 isoform-specific polyclonal antibody (dilution 1:50; StressGen) or a monoclonal antibody to NF-{kappa}B p65 (dilution 1:30; BD Transduction, San Diego, CA) for 1 h at 37°C, followed by FITC-conjugated anti-mouse or anti-rabbit IgG (dilution 1:100; Vector Laboratories, Burlingame, CA) for 1 h at 37°C. For F-actin costaining, cells were also incubated with rhodamine-phalloidin (dilution 1:100; Molecular Probes, Eugene, OR). For negative controls, cells were incubated with the secondary antibody only. Coverslips were mounted on glass slides using anti-fade mounting medium (Vector Laboratories). Slides were viewed with the use of an Image Deconvolution System consisting of a Nikon Diaphot microscope in conjunction with a cooled charge-coupled device camera coupled to the MacQuadra 950 computer system. Images were deconvolved with the use of IPLab Spectrum software for image collection and Vaytech software for deconvolution. Digital processing of the images was done with the use of PhotoShop (Adobe Systems).

Statistical analysis. Data are presented as means ± SE of absolute values or percent of control. Data were analyzed using repeated-measures ANOVA, followed by Fisher’s test for protected least-significant differences to isolate differences between groups. A level of P < 0.05 was considered significant in all statistical tests.

Materials. Cell culture reagents were purchased from Life Technologies (Gaithersburg, MD), HyClone (S. Logan, UT), and Amersham Pharmacia Biotech. Matrigel (growth factor reduced) was from Becton Dickinson (Bedford, MA). SnPP, ZnPP, and CrMP were purchased from Frontier Scientific (Logan, UT). Arachidonic acid (peroxide free) was from Cayman Chemical (Ann Arbor, MI). Peroxynitrite was purchased from Upstate (Charlottesville, VA). CORM-A1 (sodium boranocarbonate) was a generous gift from Tyco Healthcare/Mallinckrodt (Petten, The Netherlands). All other reagents were obtained from Sigma.


   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Glutamate increases ROS formation in CMVEC. First, we visualized ROS accumulation in live CMVEC using a cell-permeable oxidant-sensitive probe, DHR123 (Fig. 1). The fluorescent DHR123 oxidation derivative, rhodamine 123 (Rh123), is preferentially localized to the mitochondria in a ROS-dependent reaction (22). In control quiescent cells, Rh123 fluorescence is minimal, indicating low basal ROS formation. H2O2 and peroxynitrite were used as the control treatments to induce severe oxidative stress. H2O2 (1 mM) and peroxynitrite (1 mM) greatly increased intracellular Rh123 fluorescence (Fig. 1), indicating that DHR123 is suitable for detecting ROS production in endothelial cells. Glutamate (1–2 mM) and arachidonate (0.1 mM) rapidly (within 20 min) increased intracellular ROS formation (Fig. 1). ROS formation in live CMVEC was also observed in response to lower doses of glutamate (0.1 mM) (data not shown). Quantification of Rh123 fluorescence after a 20-min treatment of CMVEC with glutamate and other oxidants was performed using fluorescence spectrophotometry (Fig. 2A). Within 20 min, glutamate (0.1–1 mM) significantly increased Rh123 fluorescence intensity. Glutamate-induced ROS formation was comparable to that induced by arachidonate (0.1 mM) and even by strong oxidants such as H2O2 (0.1–0.5 mM) and peroxynitrite (0.1 mM; Figs. 1 and 2A). When used at high concentrations (1 mM), H2O2 and peroxynitrite had much greater effect than glutamate on ROS formation in CMVEC.


Figure 1
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  		Fig. 1. Fluorescence microscopy of reactive oxygen species (ROS) accumulation in live cerebral microvascular endothelial cells (CMVEC). Confluent quiescent CMVEC from newborn pigs preloaded with 50 µM nonfluorescent oxidant-sensitive dihydrorhodamine-123 (DHR123) were incubated with either Krebs buffer (control), 1 and 2 mM glutamate, 100 µM arachidonic acid, 1 mM H2O2, or 1 mM peroxynitrite for 30 min. Mitochondria-localized oxidized fluorescence product rhodamine-123 (Rh123) was detectable in live cells using fluorescence microscopy.

 

Figure 2
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  		Fig. 2. Effects of glutamate and oxidative stress on ROS production (A) and endothelial cell viability (B). Confluent, quiescent CMVEC from newborn pigs preloaded with 50 µM DHR123 were incubated with either Krebs buffer (Ctrl), 0.1–1 mM glutamate, 0.01–0.1 mM arachidonic acid, 0.1–1 mM H2O2, or 0.1–1 mM peroxynitrite for 20 min. A: Rh123 fluorescence of the cell lysates measured in Fusion {alpha}HT microplate analyzer (excitation/emission wavelength, 470 and 525 nm) and expressed as an increase in optical density (OD) units over the baseline. Data represent averages of 8 independent experiments. B: cell viability detected by protein quantification in attached CMVEC is expressed as percentage of control basal values. Data represent averages of 6 independent experiments. Values are means ± SE. *P < 0.05 compared with the control value.

 
Effects of glutamate, arachidonate, and severe oxidative stress on CMVEC viability. Endothelial viability after overnight exposure to glutamate and other oxidative stress-inducing agents was assessed by the number of substrate attached and floating cells (Fig. 2B) and by cytolysis detected as AK activity in the medium (Figs. 2B and 3). CMVEC showed remarkable resistance to severe oxidative stress caused by H2O2 and peroxynitrite. Endothelial cell death was observed only when high concentrations (0.5 and 1 mM) of these oxidants were used, whereas lower concentrations (0.1 M) did not alter cell viability (Fig. 2B). H2O2 (1 mM) caused endothelial cell death as indicated by cell detachment (36 ± 5% reduction in attached cells and 82 ± 2% increase in floating cells) and membrane damage (52 ± 13% increase in AK activity). Surprisingly, peroxynitrite (1 mM), which produced the greatest increase in ROS formation in CMVEC, had only mild effects on endothelial cell survival (26 ± 4% decrease in attached cells, 11 ± 5% increase in floating cells, and no changes in AK activity) (Figs. 2B and 3). Glutamate (0.5–1 mM, overnight) caused endothelial cell death as indicated by a 15 ± 6% decrease in attached cells (Fig. 2B), increased floating cells (24 ± 4%; Fig. 3A), and cytolysis (16 ± 9% increase in AK activity; Fig. 3B). Arachidonic acid (100 µM) also caused endothelial cell death comparable to the effects of glutamate (Figs. 2B and 3). Lower concentrations of glutamate (0.1 mM) and arachidonate (10–50 µM) did not decrease endothelial cell viability (Fig. 2B).


Figure 3
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  		Fig. 3. Effects of heme oxygenase (HO) inhibition on cell death caused by glutamate and other oxidative stress-inducing agents in piglet CMVEC. Confluent CMVEC from newborn pigs were incubated overnight with 1 mM glutamate, 0.1 mM arachidonic acid, 1 mM H2O2, or 1 mM peroxynitrite in the absence or presence of an HO inhibitor tin protoporphyrin (SnPP; 20 µM). Cell death detected by the number of floating cells (A) and release of cytosolic adenylate kinase (AK) activity into the media (B) expressed as percentage of control basal values. Data represent average of 8 independent experiments. Values are means ± SE. *P < 0.05 compared with the control value. {dagger}P < 0.05 compared with the corresponding indicated value.

 
Contribution of HO-2 activity to endothelial cell resistance to glutamate and oxidative stress. To investigate whether constitutive HO activity contributes to endothelial cell resistance to oxidative stress, CMVEC were exposed overnight to glutamate (1 mM), arachidonate (100 µM), and severe oxidative stress (H2O2 and peroxynitrite, 1 mM) in the presence of an HO inhibitor, SnPP (20 µM) in serum-depleted media (DMEM-0.1% FBS). SnPP at 10–50 µM effectively inhibits HO activity in isolated cerebral microvessels and in cultured CMVEC (24, 38). SnPP decreased cell resistance to serum withdrawal (Fig. 3, A and B). Furthermore, cell death induced by overnight exposure to 1 mM glutamate was exacerbated in CMVEC with inhibited HO activity, as indicated by increased numbers of detached floating cells and by an elevation in AK activity (Fig. 3, A and B). Other HO inhibitors, zinc mesoporphyrin (ZnMP) and chromium mesoporphyrin (CrMP), had similar effects (data not shown). In contrast, SnPP did not intensify endothelial cell death caused by arachidonic acid, H2O2, and peroxynitrite (Fig. 3, A and B). These data show that HO inhibition increased endothelial cell death caused by glutamate, but not by the other oxidative stress-inducing agents examined.

Effects of glutamate and oxidative stress on HO-1 and HO-2 expression in CMVEC. HO-1 expression in serum-deprived cells is minimal (Fig. 4, A and B). Prolonged exposure (20 h) to glutamate (1 mM), arachidonate (100 µM), and H2O2 (1 mM) did not upregulate HO-1 in CMVEC (Fig. 4B). HO-1 induction was observed only in cells treated with peroxynitrite (0.5 and 1.0 mM), but not with its inactive derivative (Fig. 4, A and B). The potency of peroxynitrite to induce HO-1 expression (threefold over the basal level) was only a fraction of the CoPP potency (Fig. 4, A and B). None of the agents tested, including glutamate, altered HO-2 expression in CMVEC (Fig. 4C).


Figure 4
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  		Fig. 4. Western immunoblot analysis of inducible HO (HO-1) and constitutive HO (HO-2) in piglet CMVEC. A: confluent CMVEC were incubated overnight with 0.2–1.0 mM peroxynitrite or 20 µM cobalt protoporphyrin (CoPP; representative blot). B and C: confluent CMVEC were incubated overnight with 1 mM glutamate, 0.1 mM arachidonic acid, 1 mM H2O2, and 0.2–1.0 mM active or degraded peroxynitrite. Cell lysates resolved by 12% SDS-PAGE were probed for HO-1, HO-2, and actin. HO-1 expression was normalized to actin (B), and HO-2 expression also was normalized to actin (C). Data represent averages of 3 independent experiments. Values are means ± SE. *P < 0.05 compared with the control value.

 
Apoptotic effects of glutamate in piglet CMVEC. Glutamate (1–2 mM, 3 h) initiated substantial early apoptotic changes in piglet CMVEC that included caspase-3 activation (4-fold increase; see Fig. 5), DNA fragmentation (38 ± 10% above the basal level; see Fig. 6A), and cell detachment (35 ± 8% above the baseline; see Fig. 6B). No increases in AK activity were observed (data not shown), suggesting that no cytolysis had occurred within a 3-h period of glutamate treatment. We compared the apoptotic effects of glutamate on endothelial cells with intact (control) or inhibited HO-2 activity. Because basal HO-1 expression was minimal (Fig. 5A), we assume that constitutive HO-2 activity was the major target for SnPP inhibition in quiescent CMVEC. SnPP alone (20 µM, 3-h treatment) increased caspase-3 activity (Fig. 5, A and B), DNA fragmentation, and cell detachment (Fig. 6, A and B). In CMVEC with inhibited HO-2 activity, pro-apoptotic effects of glutamate were greatly potentiated as indicated by dramatic increases in caspase-3 activation, DNA fragmentation, and numbers of floating cells (Figs. 5, A and B, and 6, A and B). These data are consistent with the essential role of HO-2 activity in the resistance of cerebral vascular endothelial cells to apoptosis induced by serum withdrawal and glutamate toxicity.


Figure 5
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  		Fig. 5. Effects of glutamate on caspase-3 activity in control and HO-1-overexpressing CMVEC from newborn pigs. HO-1 was pharmacologically induced by CoPP. A: representative immunoblots. B: caspase-3 17-kDa fragment expression normalized to actin. Control or HO-1-overexpressing CMVEC were treated for 3 h with 1 mM glutamate in the absence or presence of HO inhibitor (SnPP, 20 µM). Cell lysates resolved by 12% SDS-PAGE were probed for HO-1, caspase-3 proteolytic active fragment (17 kDa), and actin. Data represent averages of 2 independent experiments. Values are means ± SE. *P < 0.05 compared with control; {dagger}P < 0.05 compared with the corresponding indicated value.

 

Figure 6
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  		Fig. 6. Apoptotic effects of glutamate in control and HO-1-overexpressing CMVEC from newborn pigs. HO-1 was pharmacologically induced by CoPP (CoPP-CMVEC). Confluent quiescent CMVEC were incubated for 3 h with 1 mM glutamate in the absence or presence of HO inhibitor (SnPP, 20 µM). DNA fragmentation (A) and the number of floating cells (B) are expressed as a percentage of control basal values. Data represent averages of 8 independent experiments. Values are means ± SE. *P < 0.05 compared with control; {dagger}P < 0.05 compared with the corresponding indicated value.

 
Apoptotic effects of glutamate in HO-1-overexpressing CMVEC. HO-1 expression in piglet CMVEC was pharmacologically upregulated ~10-fold by CoPP (20 µM). In HO-1-overexpressing cells, glutamate (1–2 mM, 3 h) did not increase caspase-3 activity (Fig. 5, A and B) and did not alter DNA fragmentation (7 ± 3% above the baseline; P > 0.05) or cell detachment (8 ± 4% above the baseline; P > 0.05) (Fig. 6, A and B), indicating complete blockade of glutamate-induced apoptosis. SnPP treatment partially restored apoptotic effects of glutamate in HO-1-overexpressing CMVEC, as indicated by increased caspase-3 activity (Fig. 5, A and B), DNA fragmentation (12 ± 3% above the baseline, P = 0.05), and cell detachment (16 ± 4% above the baseline, P < 0.05, Fig. 6, A and B).

Apoptosis in response to serum withdrawal and glutamate in CMVEC from HO2KO and WT mice. We established primary cultures of CMVEC from HO2KO and WT mice. HO-2 and HO-1 distribution in murine CMVEC was visualized by immunofluorescence. In WT CMVEC, HO-2 was localized in the nuclear envelope and the perinuclear area of the cytoplasm, with no detectable immunostaining at the cell periphery (Fig. 7A). As expected, no HO-2 immunofluorescence was detected in HO2KO CMVEC (Fig. 7B). Similar to our findings in quiescent CMVEC from newborn pigs (39), basal HO-1 expression was detectable by immunofluorescence in both WT (Fig. 7C) and HO2KO CMVEC (Fig. 7D). In addition to the nuclear envelope and the endoplasmic reticulum area of the cytoplasm, HO-1 localization sites also include the nucleus, with selective accumulation in the nucleoli (Fig. 7, C and D). Nuclear localization sites may indicate the importance of HO-1 in the regulation of nuclear functions. HO-1 expression is the same in endothelial cells from WT and HO2KO mice, as indicated by immunocytochemistry (Fig. 7, C and D) and Western immunoblot analysis (data not shown). Previously, we reported (45) that HO-1 expression is not upregulated in HO-2-KO astrocytes and neuron/astrocyte cultures.


Figure 7
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  		Fig. 7. Primary cultures of CMVEC from wild-type (WT) (A and C) and HO-2 gene-deleted (B and D) mice. Immunofluorescence detection of HO-2 (A and B, green), HO-1 (C and D, green), and rhodamine-phalloidin F-actin costaining (A–D, red). KO, knockout.

 
HO-2 gene deletion did not reduce CMVEC viability. Growth of HO2–/– CMVEC was actually greater than that of WT cells (31 ± 3 and 24 ± 4 µg protein/well for HO2–/– and HO2+/+ CMVEC at confluence, respectively, n = 10 independent cultures; P < 0.01). However, HO-2 gene deletion reduced cell resistance to serum withdrawal. After overnight exposure to 0.1% FBS-DMEM to achieve quiescence, HO2–/– CMVEC death was ~30% higher than that in WT cells (2,180 ± 300 and 1,680 ± 360 floating cells/well for HO2–/– and HO2+/+ CMVEC, respectively, n = 10 independent cultures; P < 0.01). Furthermore, quiescent HO2–/– CMVEC had greater basal apoptosis than WT cells as detected by caspase-3 activity and DNA fragmentation (Fig. 8, A and B, and Fig. 9).


Figure 8
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  		Fig. 8. Apoptotic effects of glutamate in CMVEC from HO2KO and WT mice. Confluent quiescent murine CMVEC (WT and HO2–/–) were incubated for 3 h with 1 mM glutamate. A: caspase-3 activity detected by accumulation of the active proteolytic fragment (17 kDa); costaining for actin (representative blot). B: DNA fragmentation, the number of floating cells, and AK activity are expressed as a percentage of control basal values. Data represent averages of 7 independent experiments. Values are means ± SE. *P < 0.05 compared with the control.

 

Figure 9
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  		Fig. 9. Glutamate induced nuclear translocation of NF-{kappa}B in CMVEC from WT (A, B) and HO2KO (C, D) mice. Confluent, quiescent murine CMVEC (WT and HO2–/–) were incubated for 3 h without (A, C) or with 1 mM glutamate (B, D). Control and glutamate-treated CMVEC were immunostained with NF-{kappa}B polyclonal antibodies and visualized with FITC-conjugated anti-rabbit IgG. Note the nuclear translocation of NF-{kappa}B in glutamate-treated WT and HO2–/– CMVEC.

 
We compared the apoptotic effects of glutamate (1 mM, 3 h) in WT and HO2–/– murine CMVEC. In WT murine CMVEC, glutamate increased caspase-3 activity (Fig. 8A) and cell detachment (~20% increase over the baseline level, Fig. 8B), whereas no significant changes in DNA fragmentation or AK activity were documented (Fig. 8B). Overall, it appears that murine WT CMVECs are more resistant than piglet CMVEC to glutamate toxicity. In HO2–/– CMVEC, glutamate (1 mM, 3 h) greatly increased caspase-3 activity (Fig. 8A), DNA fragmentation, cell detachment, and elevated cytolysis-related AK activity in the media (Fig. 8B), indicating that HO-2 gene deletion increases endothelial susceptibility to glutamate-induced apoptosis and cytotoxicity.

Effects of glutamate on NF-{kappa}B localization in CMVEC from HO2KO and WT mice. NF-{kappa}B translocation from the cytoplasmic sites to the nucleus is a morphological indicator of apoptotic cell death (29, 43). Glutamate (1 mM, 3 h) caused nuclear translocation of NF-{kappa}B in murine WT and HO2–/– CMVEC (Fig. 9, A–D). In WT murine CMVEC, NF-{kappa}B is exclusively in the cytoplasm (Fig. 9A); glutamate caused nuclear translocation of NF-{kappa}B in selected cells (<10% of total cell population) (Fig. 9B). In HO2–/– CMVEC, nuclear localization of NF-{kappa}B was detectable already under basal conditions (Fig. 9C). Following a 3-h exposure of the HO2–/– CMVEC to glutamate (1 mM), nuclear localization of NF-{kappa}B (Fig. 9D) was observed in 30 ± 10% cells (5 microscopic fields in 3 independent preparations), indicating that glutamate-induced apoptosis is greatly exacerbated in HO-2 gene-deleted cerebral vascular endothelial cells.

Anti-apoptotic and anti-oxidant effects of SOD, bilirubin, and CO in a glutamate-induced cell death model in piglet CMVEC. We investigated whether SOD, bilirubin, and/or CO have cytoprotective roles in the model of glutamate-induced endothelial apoptosis. For these experiments, CMVEC were treated for 3 h with 2 mM glutamate, which caused severe endothelial cell apoptosis as detected by caspase-3 activation (Fig. 10, A and B), DNA fragmentation (Fig. 11A), and cell detachment (Fig. 11B). Bilirubin (1 µM) and a CO donor, CORM-A1 (50–100 µM) added to cerebral vascular endothelial cells for the duration of glutamate treatment prevented glutamate-induced caspase-3 activation, DNA fragmentation, and cell detachment (Figs. 10 and 11). SOD (1,000 U/ml) blocked all apoptotic changes (Figs. 10 and 11), indicating that O2–·-mediated oxidative stress triggers glutamate-induced apoptosis.


Figure 10
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  		Fig. 10. Superoxide dismutase (SOD), bilirubin, and CORM-A1 block glutamate-induced caspase-3 activation in CMVEC from newborn pigs. Confluent, quiescent CMVEC were incubated for 3 h with glutamate (2 mM) in the absence (open bar) and presence of SOD (1,000 U/ml), 1 µM bilirubin/5 µM albumin (BR) or CO-releasing molecule, CORM-A1 (50 and 100 µM). A: caspase-3 activity detected by accumulation of the active proteolytic fragment (17 kDa); costaining for actin (representative blot). B: caspase-3 activity normalized to actin expression. Data represent averages of 3 independent experiments. Values are means ± SE. *P < 0.05 compared with the control. {dagger}P < 0.05 compared with glutamate alone.

 

Figure 11
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  		Fig. 11. Antiapoptotic effects of SOD, bilirubin, and CORM-A1 against glutamate toxicity in CMVEC from newborn pigs. Confluent, quiescent CMVEC were incubated for 3 h with glutamate (2 mM) in the absence (open bar) and presence of 1,000 U/ml SOD, 1 µM bilirubin/5 µM albumin (BR) or CO-releasing molecule, CORM-A1 (50 µM). DNA fragmentation and the number of floating cells are expressed as a percentage of control basal values. Data represent averages of 3 independent experiments. Values are means ± SE. *P < 0.05, compared with control; {dagger}P < 0.05, compared with glutamate alone.

 
To investigate whether anti-apoptotic effects of HO metabolites are due to their ROS-scavenging capabilities, we used two oxidant-sensitive probes, DHR123 (ROS-sensitive probe more selective to H2O2 and peroxynitrite) and an O2–·-selective probe, DHE (21, 22, 40). Glutamate at apoptotic doses (1 mM, 3 h) greatly increased overall ROS and specifically O2–· formation in CMVEC as detected by increases in Rh123 and ethidium fluorescence, respectively (Fig. 12, A and B). SOD (1,000 U/ml) inhibited glutamate-evoked ROS formation detected by either DHR123 or DHE (Fig. 12, A and B), suggesting that O2–· is a major glutamate-induced oxidant. Bilirubin (1 µM) and CORM-A1 (50 µM) reduced glutamate-evoked Rh123 fluorescence (Fig. 12A) and completely blocked O2–· production (Fig. 12B). In a cell-free system, glutamate, SOD, bilirubin, and CORM-A1 did not change the fluorescence intensity of DHR123 or DHE (data not shown). These data indicate that bilirubin and CO are extremely effective anti-oxidants and anti-apoptotic factors in cerebral vascular endothelium.


Figure 12
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  		Fig. 12. Antioxidant effects of SOD, bilirubin, and CORM-A1 against glutamate-induced oxidative stress and apoptosis. Confluent, quiescent CMVEC from newborn pigs were incubated for 3 h with 1 mM glutamate alone (open bar) or in combination with SOD (1,000 U/ml), 1 µM bilirubin/5 µM albumin (BR), or CORM-A1 (50 µM). DHR123 (50 µM) or dihydroethidium (DHE; 100 µM) was included for the last 20 min of incubation. A: Rh123 fluorescence of the total cell lysates (excitation/emission wavelengths, 470/525 nm) expressed as percentage of control basal values. B: ethidium fluorescence of the total cell lysates (excitation/emission wavelengths, 490/605 nm) expressed as percentages of control basal values. Data represent averages of 3 independent experiments conducted in duplicate. Values are means ± SE. *P < 0.05 compared with the control values. {dagger}P < 0.05 compared with glutamate alone.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The major novel findings of our study are 1) glutamate at excitotoxic concentrations (0.5–2 mM) causes death of cerebral vascular endothelial cells via oxidative stress-related apoptosis, and 2) HO-1 and HO-2 are anti-apoptotic factors in glutamate-induced toxicity. Glutamate-induced endothelial apoptotic changes include nuclear translocation of NF-{kappa}B, caspase-3 activation, DNA fragmentation, cell detachment, and, at the later stages, membrane damage and cytolysis. Our results strongly suggest that HO-2 is highly expressed constitutively in the cerebral vasculature and is essential for endogenous protection against glutamate-induced oxidative stress and apoptosis. HO-1 is not expressed in cerebral vascular endothelium and is not induced by glutamate, suggesting that it is not an endogenous mediator of endothelial cell survival against glutamate-induced death. However, when pharmacologically upregulated, HO-1 provides complete protection from glutamate-induced apoptosis in cerebrovascular endothelium.

Apoptosis is characterized by a series of highly regulated events that result in DNA laddering, chromatin condensation, cell shrinkage, membrane blebbing, and, finally, cell death by fragmentation into apoptotic bodies, as well as phagocytosis of the cell remnants (19, 27). Cell detachment is often observed as a result of alteration in integrin signaling and loss of cell-cell contacts during apoptosis. DNA fragmentation by laddering is a hallmark of apoptosis. Major events leading to DNA fragmentation include sequential activation of caspases, a multimember family of cysteine proteases. Activation of the executioner caspases-3, -6, and-7 is thought to be a "point of no return" in the apoptotic signal transduction mechanism (19, 27, 35, 44).

Our data demonstrate that glutamate at high concentrations (mM range) causes endothelial cell death via apoptosis. In cultured cerebral vascular endothelial cells, glutamate initiates keystone events of apoptosis that include activation of the executioner caspase-3, DNA fragmentation, cell detachment, and, at the later stages, membrane damage and cytolysis. By initiating apoptotic changes within the first hours of intervention, glutamate may cause loss of endothelium-dependent vascular functions and possibly disruption of the blood-brain barrier. Indeed, our previous in vivo observations show that glutamatergic seizures extended for >1 h caused subsequent loss of cerebral vascular function (37). Glutamate caused a rapid decline of barrier functional integrity in human cerebral endothelial cells (47).

Oxidative stress appears to be a major mechanism leading to endothelial cell apoptosis caused by glutamate. In cerebral vascular cells from newborn pigs, glutamate (0.1–1 mM) increases ROS formation to a level comparable to that of strong prooxidants, including arachidonic acid, H2O2, or peroxynitrite. It appears that O2–· plays a major proapoptotic role in cerebral endothelial cells. SOD treatment blocked glutamate-induced ROS formation and provided protection from glutamate-induced DNA fragmentation and caspase-3 activation, indicating the importance of O2–· in triggering endothelial cell apoptosis. Glutamate-evoked oxidative stress also has been demonstrated in immortalized human cerebral endothelial cells (48). In neurons, oxidative stress is also a leading factor in glutamate excitotoxicity and apoptosis. Antioxidants (SOD, {alpha}-tocopherol, catalase, and ubiquinone) also protect neurons from glutamate toxicity (33).

The NF-{kappa}B pathway is involved in glutamate-induced apoptosis in cerebral endothelial cells. The transcription factor NF-{kappa}B family is involved in modulation of apoptosis following diverse injuries and oxidative stress (25, 29). Activation of the NF-{kappa}B pathway involves phosphorylation-driven translocation of the predominantly induced form, a p50 and p65/RelA heterodimer, from the cytoplasm to the nucleus, where it regulates gene transcription (25, 29, 43). Although NF-{kappa}B is often considered an anti-apoptotic protein, NF-{kappa}B activation can also enhance apoptosis in a stimuli-dependent manner (18, 25, 43, 52). NF-{kappa}B targets include antioxidant genes that regulate the synthesis of reduced glutathione and SOD (43). NF-{kappa}B targets may also include proapoptotic genes, such as TNF-{alpha} and other proinflammatory cytokines (25, 43, 52). In cerebral vascular endothelial cells, glutamate caused nuclear translocation of NF-{kappa}B consistent with subsequent transcriptional gene activation. In endothelial cells from HO2KO mice, glutamate-induced nuclear translocation of NF-{kappa}B was greatly enhanced coincident with the overall loss of resistance to glutamate cytotoxicity. This finding suggests that NF-{kappa}B activation may be proapoptotic in cerebral endothelial injury caused by glutamate. Proapoptotic effects of NF-{kappa}B in response to glutamate were also demonstrated in neurons. The NF-{kappa}B pathway mediated glutamate-induced apoptosis in neurons and glia, and blockage of NF-{kappa}B activation with aspirin was neuroprotective (8, 18).

HO-1 is recognized as an endogenous antioxidant factor that is upregulated in response to oxidative stress in various cell types, including neurons and vascular cells (6, 11, 24, 36, 41, 46, 54). HO-1 expression is transcriptionally regulated via cell-specific regulatory elements in the promoter region of the HO-1 gene, which include stress response elements or antioxidant response elements in conjunction with the redox-sensitive transcription factor Nrf2 (1, 34) and NF-{kappa}B (23). Therefore, we asked whether HO-1 is protective against glutamate-induced oxidative stress-related endothelial apoptosis. HO-1 is not detectable in the brain and cerebral microvessels under physiological conditions or following epileptic seizures (9, 37, 38) but can be upregulated by CoPP, a potent inducer of HO-1 in vivo and in vitro (24, 37). HO-1-overexpressing cerebral vascular endothelial cells demonstrated remarkable resistance against glutamate-induced apoptosis.

We examined whether glutamate and other oxidative stress-inducing agents upregulate HO-1 expression in cerebral vascular endothelial cells. Glutamate at cytotoxic concentrations (up to 2 mM), as well as arachidonic acid and H2O2, failed to induce HO-1 in cerebral vascular endothelial cells. Moderate HO-1 induction (3-fold) was observed only in response to peroxynitrite, an agent that caused the most severe oxidative stress. HO-1 induction by peroxynitrite may be cytoprotective because long-term survival was much higher in endothelial cells exposed to peroxynitrite than in those exposed to H2O2. However, it should be noted that SnPP did not reverse anti-apoptotic effects of HO-1 induction by peroxynitrite, suggesting that the mechanisms of HO-1 protection are not limited to HO activity. The anti-apoptotic role of HO-1 induced by peroxynitrite has been reported for bovine aortic endothelial cells (17).

Constitutive HO-2 provides essential but not sufficient protection against glutamate-induced apoptosis in cerebral vascular endothelial cells. Inhibition of HO-2 activity in cerebral vascular endothelial cells from newborn pigs greatly potentiated apoptosis caused by serum withdrawal and glutamate. Similarly, HO-2 gene deletion also increased the sensitivity of cerebral vascular endothelial cells to glutamate-induced apoptosis. These data demonstrate that HO-2 is protective against glutamate toxicity in the cerebral circulation. HO-2 protection appears to have a stimuli-specific character because endothelial cell death caused by other oxidative stress-inducing agents, namely, arachidonate, H2O2, and peroxynitrite, was not altered by HO activity inhibition. The mechanisms of ROS-specific and cell-specific relationships between HO-2 and oxidative stress are not completely understood. One of the possible explanations is that glutamate and ionotropic glutamate receptor agonists rapidly increase HO-2 activity, indicating a functional coupling between glutamate and HO in cerebral vascular endothelial cells (38, 39). Because glutamate induces cell death even in HO-2-expressing cerebral vascular endothelial cells, we conclude that HO-2 is essential but not sufficient for complete protection against glutamate toxicity.

We present herein data demonstrating that bilirubin and CO, the end products of HO-catalyzed heme degradation, are potent antioxidants and are cytoprotective against the apoptotic effects of glutamate in the cerebral vascular endothelium. Bilirubin also has protective effects against glutamate neurotoxicity (15). The antioxidant properties of bilirubin are credited to its ability to undergo the redox cycling in an ensemble with its precursor, biliverdin, and biliverdin reductase. Our data show that in cultured cerebral vascular endothelial cells, bilirubin blocked O2–· and other ROS responses to excitotoxic glutamate. Both pro- and anti-apoptotic effects of CO have been reported in different cell types. The anti-apoptotic effects of CO against TNF-{alpha}-mediated apoptosis have been observed in endothelial cells and fibroblasts (6, 49), but others found that CO had proapoptotic effects (51). We have provided direct experimental evidence that CO is a potent antioxidant in cerebral vascular endothelial cells. The CO donor CORM-A1 abrogated glutamate-evoked production of O2–· and other cellular oxidants in endothelial cells and completely prevented glutamate-induced apoptosis. Antioxidant properties of CO may involve inhibition of free radical generation via a direct interaction with the heme proteins of the mitochondrial electron transport chain (49). We suggest that CO is protective against glutamate-induced cerebrovascular endothelial injury mainly because of its antioxidant properties. In addition, CO may also interact with other mechanisms involved in apoptosis, such as the p38 MAPK or cGMP pathways (49), the caspase activation cascade (50, 55), and NF-{kappa}B-mediated signaling (6, 7).

Overall, glutamate causes endothelial cell death by oxidative stress-related apoptosis in cerebral vascular endothelial cells. Bilirubin and CO, the products of heme degradation by HO-2 or HO-1 activities, are potent antioxidants and anti-apoptotic factors against endothelial injury caused by excitotoxic glutamate. Although HO-1 can provide complete protection against glutamate-induced apoptosis, it is not induced in the cerebral vasculature by glutamate in vitro or in glutamatergic seizures in vivo and therefore does not appear to provide endogenous protection against glutamate toxicity in the cerebral endothelium. In contrast, HO-2 is an essential physiologically relevant component of endogenous defense against glutamate-induced apoptosis in cerebral vascular endothelium.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This research was supported by the National Institute of Neurological Disorders and Stroke and by the National Heart, Lung, and Blood Institute.


   ACKNOWLEDGMENTS
 
The authors thank Danny Morse and Greg Short for helping with preparation of the figures. We are deeply grateful to Hector Knight from Tyco Healthcare-Mallinckrodt Medical (Petten, The Netherlands) for the generous gift of CORM-A1.


   FOOTNOTES
 

Address for reprint requests and other correspondence: H. Parfenova, Dept. of Physiology, Univ. of Tennessee Health Science Center, 894 Union Ave., Memphis, TN 38163 (e-mail: hparf{at}physio1.utmem.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.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Alam J and Cook JL. Transcriptional regulation of the heme oxygenase-1 gene via the stress response element pathway. Curr Pharm Des 9: 2499–2511, 2003.[CrossRef][ISI][Medline]

2. Alberto R, Ortner K, Wheatley N, Schibli R, and Schubiger AP. Synthesis and properties of boranocarbonate: a convenient in situ CO source for the aqueous preparation of [99mTc(OH2)3(CO)3]+. J Am Chem Soc 123: 3135–3136, 2001.[CrossRef][ISI][Medline]

3. Armstead WM, Mirro R, Leffler CW, and Busija DW. Cerebral superoxide anion generation during seizures in newborn pigs. J Cereb Blood Flow Metab 9: 175–179, 1989.[ISI][Medline]

4. Bhattacharya S, Ray RM, Viar MJ, and Johnson LR. Polyamines are required for activation of c-Jun NH2-terminal kinase and apoptosis in response to TNF-{alpha} in IEC-6 cells. Am J Physiol Gastrointest Liver Physiol 285: G980–G991, 2003.[Abstract/Free Full Text]

5. Bresgen N, Karlhuber G, Krizbai I, Bauer H, Bauer HC, and Eckl PM. Oxidative stress in cultured cerebral endothelial cells induced chromosomal aberrations, micronuclei, and apoptosis. J Neurosci Res 72: 327–333, 2003.[CrossRef][ISI][Medline]

6. Brouard S, Otterbein LE, Anrather J, Tobiasch E, Bach FH, Choi AM, and Soares MP. Carbon monoxide generated by heme oxygenase 1 suppresses endothelial cell apoptosis. J Exp Med 192: 1015–1026, 2000.[Abstract/Free Full Text]

7. Brouard S, Berberat OP, Tobiash E, Seldon PM, Bach FH, and Soares MP. Heme oxygenase-1 derived carbon monoxide requires the activation of the transcription factor NF{kappa}B to protect endothelial cells form TNF-{alpha} mediated apoptosis. J Biol Chem 277: 17950–17961, 2002.[Abstract/Free Full Text]

8. Caccamo D, Campisi A, Curro M, Bramanti V, Tringali M, Volti GL, Vanella A, and Ientile R. Antioxidant treatment inhibited glutamate-evoked NF-{kappa}B activation in primary astroglial cell cultures. Neurotoxicology 26: 915–921, 2005.[CrossRef][ISI][Medline]

9. Carratu P, Pourcyrous M, Fedinec A, Leffler CW, and Parfenova H. Endogenous heme oxygenase prevents impairment of cerebral vascular functions caused by seizures. Am J Physiol Heart Circ Physiol 285: H1148–H1157, 2003.[Abstract/Free Full Text]

10. Chapman AG. Glutamate receptors in epilepsy. Prog Brain Res 116: 371–383, 1998.[ISI][Medline]

11. Chen K, Gunter K, and Maines MD. Neurons overexpressing heme oxygenase-1 resist oxidative stress-mediated cell death. J Neurochem 75: 304–313, 2000.[CrossRef][ISI][Medline]

12. Collard CD, Park KA, Montalto MC, Alapati S, Buras JA, Stahl GL, and Colgan SP. Neutrophil-derived glutamate regulates vascular endothelial barrier function. J Biol Chem 277: 14801–14811, 2002.[Abstract/Free Full Text]

13. Coyle JT and Puttfarcken P. Oxidative stress, glutamate, and neurodegenerative disorders. Science 262: 689–695, 1993.[Abstract/Free Full Text]

14. Dore S, Goto S, Sampei K, Blackshaw S, Hester LD, Ingi T, Sawa A, Traystman RJ, Koehler RC, and Snyder SH. Heme oxygenase-2 acts to prevent neuronal death in brain cultures and following transient cerebral ischemia. Neuroscience 99: 587–592, 2000.[CrossRef][ISI][Medline]

15. Dore S, Takahashi M, Ferris CD, Zakhary R, Hester LD, Guastella D, and Snyder SH. Bilirubin, formed by activation of heme oxygenase-2, protects neurons against oxidative stress injury. Proc Natl Acad Sci USA 96: 2445–2450, 1999.[Abstract/Free Full Text]

16. Duchen MR. Mitochondria and calcium: from cell signaling to cell death. J Physiol 529: 57–68, 2000.[Abstract/Free Full Text]

17. Foresti R, Sarathchandra P, Clark JE, Green CJ, and Motterlini R. Peroxynitrite induces haem oxygenase-1 in vascular endothelial cells: a link to apoptosis. Biochem J 339: 729–736, 1999.

18. Grilli M, Pizzi M, Memo M, and Spano P. Neuroprotection by aspirin and sodium salicylate through blockade of NF-{kappa}B activation. Science 274: 1383–1385, 1996.[Abstract/Free Full Text]

19. Hengartner MO. The biochemistry of apoptosis. Nature 407: 770–776, 2000.[CrossRef][Medline]

20. Henshall DC and Simon RP. Epilepsy and apoptosis pathways. J Cereb Blood Flow Metab 25: 1557–1572, 2005.[CrossRef][ISI][Medline]

21. Itoh S, Lemay S, Osawa M, Che W, Duan Y, Tompkins A, Brookes PS, Sheu SS, and Abe J. Mitochondrial Dok-4 recruits Src kinase and regulates NF-{kappa}B activation in endothelial cells. J Biol Chem 280: 26383–26396, 2005.[Abstract/Free Full Text]

22. Kooy NW, Royall JA, Ischiropoulos H, and Beckman JS. Peroxynitrite-mediated oxidation of dihydrorhodamine 123. Free Radic Biol Med 16: 149–156, 1994.[CrossRef][ISI][Medline]

23. Lavrovsky Y, Schwartzman ML, and Abraham NG. Novel regulatory sites of the human heme oxygenase-1 promoter region. Biochem Biophys Res Commun 196: 336–341, 1993.[CrossRef][ISI][Medline]

24. Leffler CW, Parfenova H, Jaggar JH, and Wang R. Carbon monoxide and hydrogen sulfide: gaseous messengers in cerebrovascular circulation. J Appl Physiol 100: 1065–1076, 2006.[Abstract/Free Full Text]

25. Lipton SA. Janus faces of NF-{kappa}B: neurodestruction versus neuroprotection. Nat Med 3: 20–22, 1997.[CrossRef][ISI][Medline]

26. Liu XM, Chapman GB, Peyton KJ, Schafer AI, and Durante W. Carbon monoxide inhibits apoptosis in vascular smooth muscle cells. Cardiovasc Res 55: 396–405, 2002.[Abstract/Free Full Text]

27. Meier P, Finch A, and Evan G. Apoptosis in development. Nature 407: 796–801, 2000.[CrossRef][Medline]

28. Meldrum BS. Concept of activity-induced cell death in epilepsy: historical and contemporary perspectives. Prog Brain Res 135: 3–11, 2002.[Medline]

29. Mercurio F and Manning AM. NF-{kappa}B as a primary regulator of the stress response. Oncogene 18: 6163–6171, 1999.[CrossRef][ISI][Medline]

30. Minetti M, Mallozzi C, Di Stasi AM, and Pietraforte D. Bilirubin is an effective antioxidant of peroxynitrite-mediated protein oxidation in human blood plasma. Arch Biochem Biophys 352: 165–174, 1998.[CrossRef][ISI][Medline]

31. Motterlini R, Sawle P, Hammad J, Bains S, Alberto R, Foresti R, and Green CJ. CORM-A1: a new pharmacologically active carbon monoxide-releasing molecule. FASEB J 19: 284–286, 2005.[Abstract/Free Full Text]

32. Murphy TH, Miyamoto M, Sastre A, Schnaar RL, and Coyle JT. Glutamate toxicity in a neuronal cell line involves inhibition of cystine transport leading to oxidative stress. Neuron 2: 1547–1558, 1989.[CrossRef][ISI][Medline]

33. Nakao N and Brundin P. Neurodegeneration and glutamate induced oxidative stress. Prog Brain Res 116: 246–263, 1998.

34. Nguyen T, Sherratt PJ, and Pickett CB. Regulatory mechanisms controlling gene expression mediated by the antioxidant response element. Annu Rev Pharmacol Toxicol 43: 233–260, 2003.[CrossRef][ISI][Medline]

35. Nicholson DW. Caspase structure, proteolytic substrates and function during apoptotic cell death. Cell Death Differ 6: 1028–1042, 1999.[CrossRef][ISI][Medline]

36. Otterbein LE and Choi AM. Heme oxygenase: colors of defense against cellular stress. Am J Physiol Lung Cell Mol Physiol 279: L1029–L1037, 2000.[Abstract/Free Full Text]

37. Parfenova H, Carratu P, Tcheranova D, Fedinec A, Pourcyrous M, and Leffler CW. Epileptic seizures cause extended postictal cerebral vascular dysfunction that is prevented by HO-1 overexpression. Am J Physiol Heart Circ Physiol 288: H2843–H2850, 2005.[Abstract/Free Full Text]

38. Parfenova H, Fedinec A, and Leffler CW. Ionotropic glutamate receptors in cerebral microvascular endothelium are functionally linked to heme oxygenase. J Cereb Blood Flow Metab 23: 190–197, 2003.[CrossRef][ISI][Medline]

39. Parfenova H, Neff RA III, Alonso JS, Shlopov BV, Jamal CN, Sarkisova SA, and Leffler CW. Cerebral vascular endothelial heme oxygenase: expression, localization, and activation by glutamate. Am J Physiol Cell Physiol 281: C1954–C1963, 2001.[Abstract/Free Full Text]

40. Pias EK, Ekshyyan OY, Rhoads CA, Fuseler J, Harrison L, and Aw TY. Differential effects of superoxide dismutase isoform expression on hydroperoxide-induced apoptosis in PC-12 cells. J Biol Chem 278: 13294–13301, 2003.[Abstract/Free Full Text]

41. Poss KD, Thomas MJ, Ebralidze AK, O’Dell TJ, and Tonegawa S. Hippocampal long-term potentiation is normal in heme oxygenase-2 mutant mice. Neuron 15: 867–873, 1995.