Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M400101200 on January 29, 2004

J. Biol. Chem., Vol. 279, Issue 18, 19099-19112, April 30, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/18/19099    most recent
M400101200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Shang, T.
Right arrow Articles by Kalyanaraman, B.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shang, T.
Right arrow Articles by Kalyanaraman, B.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

1-Methyl-4-phenylpyridinium-induced Apoptosis in Cerebellar Granule Neurons Is Mediated by Transferrin Receptor Iron-dependent Depletion of Tetrahydrobiopterin and Neuronal Nitric-oxide Synthase-derived Superoxide*

Tiesong Shang{ddagger}, Srigiridhar Kotamraju{ddagger}, Shasi V. Kalivendi{ddagger}, Cecilia J. Hillard§, and B. Kalyanaraman{ddagger}

From the {ddagger}Department of Biophysics and Free Radical Research Center and the §Department of Pharmacology and Toxicology, Medical College of Wisconsin, Milwaukee, Wisconsin 53226

Received for publication, January 6, 2004 , and in revised form, January 28, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In this study, we investigated the molecular mechanisms of toxicity of 1-methyl-4-phenylpyridinium (MPP+), an ultimate toxic metabolite of a mitochondrial neurotoxin, 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine, that causes Parkinson-like symptoms in experimental animals and humans. We used rat cerebellar granule neurons as a model cell system for investigating MPP+ toxicity. Results show that MPP+ treatment resulted in the generation of reactive oxygen species from inhibition of complex I of the mitochondrial respiratory chain, and inactivation of aconitase. This, in turn, stimulated transferrin receptor (TfR)-dependent iron signaling via activation of the iron-regulatory protein/iron-responsive element interaction. MPP+ caused a time-dependent depletion of tetrahydrobiopterin (BH4) that was mediated by H2O2 and transferrin iron. Depletion of BH4 decreased the active, dimeric form of neuronal nitric-oxide synthase (nNOS). MPP+-mediated "uncoupling" of nNOS decreased .NO and increased superoxide formation. Pretreatment of cells with sepiapterin to promote BH4 biosynthesis or cell-permeable iron chelator and TfR antibody to prevent iron-catalyzed BH4 decomposition inhibited MPP+ cytotoxicity. Preincubation of cerebellar granule neurons with nNOS inhibitor exacerbated MPP+-induced iron uptake, BH4 depletion, proteasomal inactivation, and apoptosis. We conclude that MPP+-dependent aconitase inactivation, Tf-iron uptake, and oxidant generation result in the depletion of intracellular BH4, leading to the uncoupling of nNOS activity. This further exacerbates reactive oxygen species-mediated oxidative damage and apoptosis. Implications of these results in unraveling the molecular mechanisms of neurodegenerative diseases (Parkinson's and Alzheimer's disease) are discussed.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Parkinson's disease (PD)1 is characterized by mitochondrial complex I defects, elevated iron levels in brain tissue, tetrahydrobiopterin (BH4) and dopamine deficiencies, and {alpha}-synuclein accumulation in Lewy body aggregates (1-4). The exact molecular mechanisms leading to the pathophysiology of PD are not well understood. 1-Methyl-4-phenylpyridinium ion (MPP+), a mitochondrial complex I inhibitor, produces Parkinson-like symptoms in humans and laboratory animals, and has been used to investigate the mechanism of pathogenesis of PD (5, 6). MPP+ is the ultimate metabolite of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP), a contaminant found in illicit narcotics (7, 8). MPP+ is taken up into dopaminergic neurons via the dopamine transporter and accumulates into mitochondria, where it inhibits complex I activity. Although the mechanism of MPP+-induced neurotoxicity is not fully understood, there is increasing evidence supporting the involvement of reactive oxygen species (ROS) and reactive nitrogen species (RNS) (1, 9).

Previous studies using the neuronal nitric-oxide synthase (nNOS) knockout mice implicate nitric oxide (.NO) and peroxynitrite (ONOO-) in MPP+-induced neurodegeneration (10). However, therapeutic intervention studies with nitric-oxide synthase (NOS) inhibitors in MPTP-treated mice demonstrated the opposite result, i.e. the attenuation of .NO was more deleterious than protective (11-14). Decreased formation of .NO-derived metabolites (nitrite and nitrate) in cerebrospinal fluids was observed in PD (15). Furthermore, low levels of tetrahydrobiopterin (BH4) were detected during the onset and progression of PD (2, 16, 17). BH4 is a critical cofactor for NOS activity (18, 19). Evidence indicates that BH4, by acting as a "redox switch," plays a critical role in increasing not only the rate of .NO generation by NOS, but also in controlling the formation of superoxide and hydrogen peroxide (20-22). Pharmacological manipulation of BH4 has been suggested as a therapeutic strategy to modulate .NO and superoxide generation in neuronal cells and endothelial cells (23-27). De novo biosynthesis of BH4 is catalyzed by guanosine 5'-triphosphate cyclohydrolase I (GTPCH I) (28, 29). Mammalian cells can also generate BH4 by another pathway wherein sepiapterin is converted to BH4 by sepiapterin reductase and dihydrofolate reductase (28, 29). Intracellular BH4 levels are inhibited by 2,4-diamino-6-hydroxypyrimidine (DAHP), an inhibitor of GTPCH I, and enhanced by sepiapterin treatment of cells. However, modulation of BH4 synthesis has yielded opposing results in cardiovascular and cerebrovascular systems (23-27). Blockade of BH4 biosynthesis in animals protects against postischemic neuronal injury (23), whereas increased intracellular BH4 from supplementation with sepiapterin enhanced endothelial function thereby inhibiting postischemic heart injury (24, 25). Thus, the exact relationship between .NO, BH4, and cytotoxicity/cytoprotection still remains ambiguous and further mechanistic studies clarifying the role of BH4 in oxidative injury in cellular systems deserve additional consideration.

Oxidant-induced iron signaling and the subsequent occurrence of free radical-mediated oxidative damage are becoming increasingly understood as playing highly significant roles in neurodegeneration (30-32). Mitochondrial toxins that stimulate and H2O2 cause excessive accumulation of cellular iron (33, 34). Iron has been shown to be abundantly present in the substantia nigra, whose levels have been reported to be consistently enhanced in the PD brain (4). A major pathway by which neuronal cells acquire iron is via the transferrin receptor (TfR), which facilitates the uptake of iron-loaded transferrin (35). In a previous study, we showed that MPP+ up-regulates TfR expression and iron uptake, leading to oxidative stress and apoptosis in neuronal cells (36).

Recently, we and others reported that .NO is critical to regulating oxidant-induced intracellular iron uptake (37-39). In this study, we seek to understand better the interaction between .NO, BH4, iron, and neuronal apoptosis in response to MPP+ treatment. For these studies, we used cerebellar granule neurons (CGNs) for the following reasons: (i) CGNs express an organic cation transporter 3 that makes them vulnerable to MPP+ toxicity, much like neurons expressing the dopamine transporter (40); (ii) CGNs express a high level of nNOS that makes it possible to test the "uncoupling" mechanism of nNOS influenced by BH4 modulation; and (iii) MPP+ induces an iron-dependent apoptosis in CGNs (36). Results indicate that MPP+-induced BH4 depletion is dependent on TfR-iron signaling and is caused by MPP+-dependent TfR up-regulation, which ultimately results in nNOS uncoupling (i.e. switching from .NO to superoxide generation). Increasing the intracellular BH4 levels by sepiapterin supplementation partially restored mitochondrial respiratory enzyme activity, restored the nNOS activity (.NO generation), inhibited superoxide formation from the uncoupling mechanism, and prevented TfR-dependent iron accumulation and apoptotic cell death. Implications for the role of nNOS in iron signaling in PD are discussed.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—1-Methyl-4-phenylpyridinium (MPP+), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT), deferoxamine, and NG-nitro-L-arginine-methyl ester HCl (L-NAME) were purchased from Sigma. L-Sepiapterin, DAHP, N-acetylserotonin (NAS), and NG-monomethyl-L-arginine monoacetate (L-NMMA) were bought from Alexis Biochemicals (San Diego, CA). 7-Nitroindazole was purchased from Calbiochem-Novabiochem Corp. (La Jolla, CA). The fluorogenic substrates for proteasome assay Boc-VLK-AMC (Boc-Val-Leu-Lys-7-amido-4-methylcoumarin) was bought from Sigma, and N-Suc-LLVY-AMC (N-succinyl-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin) was bought from Biomol%20Research%20Laboratories">Biomol Research Laboratories, Inc. (Plymouth Meeting, PA). HBED (N,N'-bis(2-hydroxybenzyl ethylenediamine-N,N'-diacetic acid) was a kind gift from Dr. Cherakuri Muralikrishna (National Cancer Institute, Bethesda, MD.). Carboxy-2',7'-dichlorodihydrofluorescein diacetate (carboxy-H2DCFDA) and dihydroethidium (hydroethidine or DHE) were purchased from Molecular Probes Inc. (Eugene, OR). QuickPrep micro mRNA purification kit, Ready-to-go You-Prime-First-Strand Beads, and First-strand cDNA synthesis kit were purchased from Amersham Biosciences. TRIzol reagent was obtained from Invitrogen. The caspase 3 assay kit (colorimetric) was purchased from Sigma. ApoAlert DNA fragmentation assay kit was purchased from Clontech (Palo Alto, CA). Mouse-anti-human TfR antibody was purchased from Zymed Laboratories Inc. (South San Francisco, CA). Mouse anti-rat nNOS protein antibody was purchased from BD Transduction Laboratories (San Diego, CA). Mouse-anti-actin antibody was purchased from Chemicon International Inc. (Temecula, CA). Monoclonal antibody 42/6 against human TfR was a gift from Dr. Ian Trowbridge (Salk Institute, San Diego, CA). 55Fe (ferric chloride) (1 mCi/100 µl) was obtained from PerkinElmer Life Sciences. Rat GTP cyclohydrolase I (GTPCH I) and rat transferrin receptor-specific cDNA primers (shown below) were synthesized by Operon Technologies (Alameda, CA). Other salts and buffers were obtained from the usual commercial sources.

CGN Primary Culture—CGNs were prepared from 6-8-day-old rat pups of either sex exactly as described previously (41). Cells were plated at density of 1.4 x 106 cells/ml in 12- or 6-well plates coated with poly-D-lysine. For the terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling (TUNEL) assay, CGNs were plated at a density of 1 x 106 cells/ml. After 24 h, cytosine arabinoside (10 µM) was added to the cultures to inhibit glial proliferation. Approximately 5% of the cells present in the cultures are astroglia; a few phase-bright microglia can be seen in each high power field, and the remaining cells are CGNs. CGNs were used for experiments on day 10.

Reverse Transcription-Polymerase Chain Reaction (RT-PCR)—For RT-PCR of GTPCH I, poly(A)+ mRNA was prepared from either cells or tissues using QuickPrep micro mRNA purification kit and cDNA was synthesized using the You-Prime-First-Strand Beads using oligo(dT) as a primer. The sense and antisense primers for rat GTPCH I were 5'-GGA TAC CAG GAG ACC ATC TCA-3' and 5'-TAG CAT GGT GCT AGT GAC AGT-3', respectively. These primers correspond to the nucleotide positions 422-442 and 773-793, respectively, and size of the expected product was 372 bp. An annealing temperature of 55 °C was used, and the cycle was repeated 30 times. For semiquantitative RT-PCR of TfR, total RNA from CGNs was extracted using TRIzol (Invitrogen). Two micrograms of total RNA was reverse transcribed using a first-strand cDNA synthesis kit (Amersham Biosciences) employing random hexamers. For RT-PCR, the number of cycles and the amount of cDNA are optimized to fall into the exponential phase of the polymerase chain reaction. The sense and antisense primers for rat transferrin receptor were 5'-AGT TTC CGC CAT CTC AGT C-3' and 5'-ACG TCC TGC ATT ATC TTC CC-3', respectively. These primers correspond to the nucleotide positions 546-564 and 1106-1125, respectively, and size of the expected product was 580 bp. An annealing temperature of 55 °C was used, and the cycle was repeated 30 times. All primers were designed employing Genetics Computer Group, Inc. (GCG) software. The PCR products were resolved on 2% agarose gel, and the bands were quantified by densitometry.

MTT Reduction Cytotoxicity Assay—MTT is converted in living cells to formazan, which has a specific absorption maximum at 562 nm. Treatment agents were added directly to the culture media. After treatment, the culture media were removed and cells were washed three times with control salt solution (CSS) (120 mM NaCl, 25 mM HEPES, pH 7.4, 25 mM KCl, 1.8 mM CaCl2, 4 mM MgCl2, and 15 mM glucose). Cells were incubated in CSS buffer containing 0.25 mg/ml MTT for 2 h at 37 °C. CSS buffer was removed, and cells were solubilized and mixed thoroughly in isopropanol, 0.08 N HCl (1:1). The absorption was measured at 562 nm with reference at 630 nm.

Measurement of Caspase-3-like Protease Activity—Caspase-3-like protease activity was assayed according to the protocol from the manufacturer (Caspase 3 Assay Kit, Sigma) with the following modification. CGNs were cultured in 6-well plates and exposed to MPP+ on day 10. Pretreatment drugs were added in culture media 2 h before MPP+ treatment. After a 24-h incubation, CGNs were washed, scraped, and pelleted with cold PBS. The cell pellet was lysed in 50 µl of lysis buffer (50 mM HEPES, pH 7.4, 5 mM CHAPS, and 5 mM DTT). Following homogenization and incubation on ice for 15-20 min, lysates were centrifuged at 16,000 x g at 4 °C for 20 min. Aliquots of 40 µl of lysate were incubated with 50 µl of assay buffer (20 mM HEPES, pH 7.4, 0.1% CHAPS, 5 mM DTT, 2 mM EDTA) and 10 µl of caspase-3 substrate (Ac-DEVD-p-nitroaniline, 200 µM) at 37 °C for 2 h. Protein concentration was determined according to the Lowry method.

Terminal Deoxynucleotidyltransferase-mediated dUTP Nick End Labeling (TUNEL) Staining—CGNs were cultured on chamber slips at a density of 1 x 106 cells/ml. Beginning on day 10, the cells were exposed to MPP+ for 30 h. Pretreatment drugs were added in culture media 2 h before MPP+ treatment. DNA fragmentation of individual cells was detected in situ by TUNEL with ApoAlert DNA fragmentation assay kit (Clontech). Briefly, cells were washed with PBS and fixed in formaldehyde solution (4% in PBS) for 25 min at 4 °C. Cells were washed and permeabilized in a solution containing 0.2% Triton X-100 in PBS for 5 min at 4 °C, followed by incubation in freshly prepared terminal deoxynucleotidyltransferase incubation buffer for 60 min at 37 °C in the dark. Cells were washed with PBS, stained with propidium iodide, mounted in an anti-fade solution, and visualized under a Nikon microscope. Fluorescein-dUTP incorporated into the fragmented nucleotides exhibited green fluorescence at an emission wavelength of 528 nm. All cells stained with propidium iodide exhibited red fluorescence at an emission wavelength of 617 nm. Cells were identified as apoptotic if they showed positive TUNEL staining (yellow color on overlay images). Apoptosis was assessed by counting the number of TUNEL-positive cells in at least three randomly selected fields and expressed as a percentage of the total number of cells counted.

Determination of TfR and nNOS Levels—CGNs were washed with ice-cold PBS and resuspended in 50 µl of radioimmune precipitation assay buffer (20 mM Tris-HCl, pH 7.4, 2.5 mM EDTA, 1% Triton X-100, 1% sodium deoxycholate, 1% SDS, 100 mM NaCl, 100 mM sodium fluoride). To a 10-ml solution of the above, 1 mM sodium vanadate and 1 tablet of protease inhibitor mixture (Roche Applied Science) were added. Cells were homogenized, and the lysate was centrifuged for 10 min at 12,000 x g at 4 °C. Protein was determined by the Lowry method, and 20 µg was used for the Western blot analysis. Proteins were resolved on 8% SDS-polyacrylamide gels and blotted onto nitrocellulose membranes. Low temperature SDS-PAGE (LT-PAGE) was performed according to a published method (42). Basically, equal amounts of protein were mixed with Laemmli sample buffer at 4 °C. Gel (6%) and buffer were equilibrated at 4 °C prior to electrophoresis and the buffer tank placed in 4 °C refrigerator during electrophoresis to keep the temperature below 10 °C. Subsequent to LT-PAGE, protein was transferred to a nitrocellulose membrane. Membranes were washed with Tris-buffered saline (140 mM NaCl, 50 mM Tris-HCl, pH 7.2) containing 0.1% Tween 20 and 5% skim milk to block nonspecific protein binding. Membranes were incubated with mouse anti-human transferrin receptor monoclonal antibody (Zymed Laboratories Inc. 1 µg/ml) or mouse anti-rat neuronal nitric-oxide synthase monoclonal antibody (BD Transduction Laboratories; 1 µg/ml) in Tris-buffered saline containing 0.1% Tween 20 for 1.5 h at room temperature. Following washing, the membranes were incubated with horseradish peroxidase-conjugated rabbit anti-mouse IgG (1:5000) for 1.5 h at room temperature. The TfR and nNOS bands were detected using the ECL method (Amersham Biosciences). Subsequently, blots were stripped and reprobed with an antibody against actin.

Measurement of Aconitase Activity—Aconitase activity was measured according to the published method with some modifications (43). CGNs were washed twice with cold phosphate-buffered saline (PBS), scraped, and centrifuged at 3,000 rpm for 10 min to pellet the cells and homogenized using a 25-gauge syringe needle (10 strokes) with 100 µl of lysis buffer (0.2% Triton X-100, 100 µM diethylenetriamine pentaacetic acid, and 5 mM citrate in PBS). The lysate was centrifuged at 12,000 rpm for 10 min, aconitase activity was measured in the supernatant, and protein was estimated by the Lowry method. The activity of aconitase was measured in 100 mM Tris-HCl (pH 8.0, 100 µl) containing 20 mM DL-trisodium isocitrate. The rate of change of absorbance was followed for 3 min at 240 nm in a UV-visible spectrophotometer (UV-1601PC; Shimadzu Corp.) at room temperature. An extinction coefficient for cis-aconitase of 3.6 mM-1 cm-1 at 240 nm was used to calculate the enzyme activity and the values were expressed as micromoles/min/mg of protein.

Electrophoretic Mobility Shift Assay—IRP/IRE binding was measured by the electrophoretic mobility shift assay. 32P-Labeled IRE mRNA for the RNA band shift assay was prepared using as a template a 1000-base pair rat L ferritin pseudogene that contains the conserved IRE sequence. The plasmid (p66-L gene) containing this insert (which was generously provided by Dr. Elizabeth Leibold) was linearized with SmaI (Invitrogen) and used for in vitro transcription of IRE mRNA. Transcription was carried out with Sp6 RNA polymerase using a Ribo-probe transcription system from Promega.

Measurement of Oxidative Stress—The determination of intracellular oxidant production was based on the oxidation of carboxy-H2DCFDA, resulting in the formation of the fluorescent compound carboxy-2',7'-dichlorofluorescein (carboxy-DCF). Following treatment of CGNs with MPP+, the medium was aspirated, and cells were washed twice with PBS and incubated in 1 ml of cell culture medium without fetal bovine serum. Carboxy-H2DCFDA was added to a final concentration of 10 µM and incubated for 20 min. Cells were washed with PBS and maintained in 1 ml of culture medium. Fluorescence was monitored using a Nikon fluorescence microscope with wavelengths of 480/30 nm (excitation) and 535/40 nm (emission). The fluorescence intensity values from three different fields of view were calculated using the Metamorph software.

Hydroethidine (Dihydroethidium) Staining—The redox-sensitive, cell-permeable fluorophore hydroethidine (dihydroethidium, DHE) was used to evaluate the cellular production of superoxide in vivo. Following treatment of CGNs, culture medium was aspirated and cells were washed once with PBS and incubated in a fresh culture medium without fetal bovine serum. DHE (5 µM) was added to cells, and the incubation was continued for 20 min. Fluorescence images were obtained using a Nikon fluorescence microscope at excitation and emission wavelengths of 540/25 nm and 605/55 nm, respectively. The fluorescence intensity values from three different fields of view were calculated using the Metamorph software, and the average values presented.

Determination of Tetrahydrobiopterin Levels in CGNs—Tetrahydrobiopterin levels in CGNs were assayed using HPLC with fluorescence detection according to the method reported previously (44). Cells were washed three times with ice-cold PBS. For assays of total (reduced + oxidized) biopterin, 0.5 ml of ice-cold 0.1 N HCl was added to each well and cells were collected; for assays of oxidized biopterin, 0.5 ml of ice-cold 0.01 N NaOH was added to each well and cells were collected. After sonication for 1 min at 40 watts in ice, the cell suspension was centrifuged for 20 min at 12,000 x g at 4 °C. The supernatant (100 µl) was assayed for protein concentration by Bradford analysis (Bio-Rad) according to the instructions from the manufacturer. The supernatant (0.3 ml) was mixed with 50 µl of iodine solution (2%/1% KI/I2 in distilled water) for biopterin measurement. The mixture was incubated at room temperature for 1 h in the dark. After incubation, 1 µl of 5 N HCl was added only to the mixture for alkaline-oxidized biopterin. Excess iodine was reduced by adding 5 µl of 50% ascorbic acid solution. Biopterin levels were analyzed by HPLC (using a C18 reverse-phase column) with a fluorescence detector with excitation at 350 nm and emission at 440 nm using authentic biopterin as standard. The chromatographic mobile phase was 25 mM phosphate buffer, pH 6.4, and was delivered at a flow rate of 0.7 ml/min. The reduced biopterin was determined by subtracting the oxidized biopterin from the total biopterin.

Measurement of Nitrite and Nitrate—Nitrite and nitrate, oxidative metabolites of .NO, were measured by chemiluminescence according to a modified method (45). Briefly, after treatment of CGNs, culture medium was aspirated and cells were washed three times with Hanks' buffer (110 mM NaCl, 25 mM KCl, 20 mM HEPES, 5.5 mM glucose, 1.25 mM CaCl2,1mM MgCl2, pH 7.4). Cells were incubated with fresh Hanks' buffer containing 25 µM L-arginine. Nitrite/nitrate levels in the incubation buffer were first measured (sample I). Cells were incubated with the buffer for 30 min at 37 °C. Samples were analyzed for nitrite/nitrate content (sample II). Nitrite/nitrate present in sample I was subtracted from sample II to correct for the background nitrite/nitrate contamination. Protein concentration in each well was measured according to the Bradford method. For nitrite measurement, a solution containing KI and glacial acetic acid was used. For nitrate and nitrite measurement, a solution containing vanadium and HCl was used. .NO generated from sodium nitrite and sodium nitrate were used as standards.

Measurement of GTPCH I Activity in CGNs—GTPCH I activity in CGNs was assayed using HPLC with fluorescence detection. Cells were washed three times with ice-cold PBS and collected in 0.5 mM Tris-HCl, pH 7.4, containing protease inhibitor (Roche Applied Science). After sonication for 1 min at 40 watts in ice, the cell suspension was centrifuged for 20 min at 12,000 x g at 4 °C. Cell lysate was incubated with 1 mM GTP, 1 mM DTT, 1 mg/ml bovine serum albumin in 0.1 mM Tris-HCl, pH 7.4 for 2 h at 37 °C in the dark. The reaction product, dihydroneopterin, was oxidized to neopterin triphosphate using the iodine solution (2%/4% I2/KI) for 10 min in the dark. Excess iodine was reduced by ascorbic acid, and the pH was adjusted to alkalinity by adding the 2 M Tris-base. Neopterin triphosphate was dephosphorylated by the alkaline phosphatase (5 units/100 µl) at 37 °C for 1 h. The reaction was stopped by acidification with 1 M H3PO4, and the product neopterin was determined by a reverse-phase HPLC (a 5C18 reverse-phase column) using a fluorescence detector with an excitation wavelength at 350 nm and an emission wavelength at 440 nm using authentic neopterin as standard. The chromatographic mobile phase was 3% acetonitrile in distilled water and delivered at a flow rate of 0.1 ml/min.

Measurement of 55Fe Uptake in CGNs—CGNs were cultured in 6-well plates and treated with MPP+ for desired time points. Ferric chloride (0.2 µCi/ml 55Fe) was added to the media and incubated for another 4 h at 37 °C. After incubation, 200 µl of medium was taken to count 55Fe in media. Cells were washed in cold 1x PBS three times and lysed in 500 µl of lysis buffer (1x PBS, 0.1% Triton X-100, and protease inhibitor). After centrifuging at 6000 x g at room temperature for 10 min, 350 µl of supernatant was taken to count 55Fe in cell lysate. Supernatant (50 µl) was used to measure protein concentration by the Bradford method.

Proteasome Function Assays—Proteasome function was measured as reported previously (37, 46). Briefly, cells were lysed in 100 µl of ice-cold homogenization buffer (20 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, 2 mM DTT, 5 mM ATP, 20% (v/v) glycerol, and 0.04% (v/v) Nonidet P-40). Lysate (10 µl) was taken to measure protein concentration by Bradford method. Lysate (40 µl) was incubated at 37 °C with the fluorogenic substrate Boc-VLK-AMC (80 µM, trypsin-like) or Suc-LLVY-AMC (100 µM, chymotrypsin-like) in 500 µl of assay buffer (50 mM Tris-HCl, pH 7.4, 2 mM DTT, 5 mM MgCl2, and 2 mM ATP) for 30 and 60 min, respectively. Proteolytic activity was measured by monitoring the release of the fluorescent group 7-amido-4-methylocoumarin (AMC) at 380 nm excitation and 460 nm emission wavelengths using a Shimadzu spectrofluorophotometer (RF-5301PC). All readings were standardized using the fluorescence intensity of an equal volume of free AMC solution.

Data Analysis—Statistical significance was obtained using Student's t test employing the Sigmastat software.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
MPP+-induced Iron Signaling in CGNs—Cytosolic aconitase is an indicator of intracellular iron level and oxidant formation and has been shown to play an important role as an iron sensor (47-49). CGNs were treated with MPP+ (70 µM), and the total aconitase activity was measured in whole cell lysates at different time points. As shown in Fig. 1A, a steady decrease in aconitase activity with time was observed. The aconitase activity initially decreased after a 12-h incubation with MPP+, and further decreased with prolonged incubation of 48 h. Inactivation of aconitase by oxidative destruction of the [4Fe-4S] cluster is accompanied by an increase in its mRNA (IRE) binding activity, which is known as the "IRP1 activity" (49). As shown in Fig. 1B, treatment of cells with 70 µM MPP+ caused a time-dependent increase in IRP1 activity. At least a 10-fold increase in active IRP1 with respect to total IRP1 (active and inactive) was detected in MPP+-treated CGNs within 12 h (Fig. 1B, left panel). When lysates were treated with 1% {beta}-mercaptoethanol to completely activate IRP1 to the high affinity mRNA-binding form (50), differences in IRP1 binding to IRE were not observed between control and MPP+-treated cells (Fig. 1B, right panel). These data suggest that MPP+ treatment causes IRP1 activation, likely as a consequence of the inactivation of aconitase.



View larger version (26K):
[in this window]
[in a new window]
  		FIG. 1.
MPP+-induced changes in aconitase activity, IRP activity, TfR expression, and 55Fe uptake in CGNs. CGNs were treated with 70 µM MPP+ for different time periods as indicated, and the total aconitase activity was measured in whole cell lysates (A), the IRP1 activity was analyzed in cytoplasmic extracts by the gel shift assay with and without {beta}-mercaptoethanol (1%) (B), TfR protein levels were determined in cell lysates by Western blotting analysis using the anti-TfR antibody (C), TfR mRNA levels were determined by the RT-PCR method as described under "Experimental Procedures" (D), and CGNs were treated with 70 µM MPP+ for different time intervals as indicated and further incubated with 55Fe (0.2 µCi/ml) for 4 h (E). The 55Fe content in cell lysates was determined. The gels are representative of three independent experiments, and values are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; ***, p < 0.001 compared with control.

 
A result of increased IRP1/IRE binding is stabilization of TfR mRNA leading to increased TfR synthesis. The effects of MPP+ treatment on TfR protein and mRNA levels as a function of time were determined. As shown in Fig. 1 (C and D), MPP+ treatment induced the expression of TfR protein and mRNA in CGNs. TfR levels increased in MPP+-treated cells after 12 h and remained elevated up to 48 h. To examine whether increased cellular TfR expression results in increased cellular iron, we measured cellular 55Fe uptake in MPP+-treated cells. Fig. 1E shows that incubation with 70 µM MPP+ caused an increase in 55Fe uptake by cells. These results are consistent with the hypothesis that MPP+-induced iron uptake occurs through an IRP1-regulated, TfR-dependent transport mechanism.

MPP+-induced Intracellular BH4 Depletion and Inhibition of nNOS Activity and nNOS Dimer Levels in CGNs—We have shown previously that hydrogen peroxide-induced TfR iron uptake is critically dependent on the intracellular .NO concentration (37). As BH4 is one of the crucial regulators of NOS activity, we monitored intracellular BH4 level in MPP+-treated CGNs. Fig. 2A shows that 70 µM MPP+ treatment induced a time-dependent reduction in intracellular BH4 levels. After a 24-h incubation, intracellular BH4 was decreased significantly to ~50% of control. Intracellular BH4 further decreased after a 36-h incubation, and remained at that level after a 48-h incubation with MPP+. Under conditions when BH4 levels are insufficient, NOS activity decreases as a result of uncoupling of NOS (21). nNOS activity was monitored by measuring the nitrate/nitrite levels in MPP+-treated cells. Fig. 2B shows that 70 µM MPP+ treatment for 24 h significantly decreased nitrate/nitrite formation. To investigate whether the decrease in nitrate/nitrite formation was the result of altered nNOS protein levels, we determined the effect of MPP+ on the expression of cellular nNOS protein by Western blotting. BH4 is crucial for nNOS dimerization, and BH4 deficiency has been reported to cause a decrease in NOS dimer levels (42). LT-PAGE was used to examine the effect of MPP+ treatment on nNOS dimer formation in CGNs. As shown in Fig. 2 (C and D), MPP+ (70 µM) treatment decreased nNOS dimer in a time-dependent manner. However, there was also a decrease in nNOS monomer levels, the reasons for which remain unclear. Nonetheless, as reported in a recent study (51), it is conceivable that nNOS antibody did not fully recognize the monomer form of nNOS, which could account for decreased nNOS dimer levels without the concomitant increase in the monomer form. These findings show that MPP+ treatment decreases the BH4 levels and .NO generation in CGNs via decreased stabilization of the dimeric form of nNOS.



View larger version (17K):
[in this window]
[in a new window]
  		FIG. 2.
The effect of MPP+ on intracellular BH4 levels, nNOS activity, and nNOS dimer level in CGNs. A, CGNs were treated with 70 µM MPP+ for different time periods, and the intracellular BH4 levels were measured using the HPLC method as described under "Experimental Procedures." B, CGNs were treated with 70 µM MPP+ or 100 µM L-NMMA for 24 h, and the nNOS activity was determined by measuring nitrate/nitrite levels as described under "Experimental Procedures." C, CGNs were treated with 70 µM MPP+ for different time periods as indicated, and nNOS dimer levels were determined by LT-PAGE and Western blotting. D, the densitometric analyses of nNOS monomer and dimer levels were measured as shown in C. The gels are representative of three independent experiments, and values are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; ***, p < 0.001 compared with control.

 
Effect of BH4 Deficiency on MPP+-induced Superoxide Generation in CGNs—We and others have shown previously that nNOS generates superoxide in the absence of BH4 and that the ratio between .NO and superoxide is critically regulated by BH4 levels (21, 52). Intracellular BH4 levels were manipulated to clarify the role of BH4 and nNOS in MPP+-induced toxicity in cells. Intracellular BH4 content was reduced by treating cells with inhibitors of BH4 de novo synthesis. In agreement with a previous report (53), Fig. 3 (A and B) shows that inhibiting BH4 synthesis by incubating cells with DAHP (1 mM) and NAS (100 µM) for 24 h caused an increase in superoxide generation in CGNs. We then investigated superoxide generation in MPP+-treated cells. As shown in Fig. 3 (C and D), treatment of cells with 1 mM DAHP and 50 µM MPP+ caused a significant increase in superoxide generation, as compared with MPP+ or DAHP treatment alone.



View larger version (33K):
[in this window]
[in a new window]
  		FIG. 3.
The effect of depleting intracellular BH4 levels on superoxide formation in MPP+-treated and untreated CGNs. A, CGNs were treated with 1 mM DAHP and 100 µM NAS for 24 h, and superoxide formation measured by DHE staining as described under "Experimental Procedures." B, the mean fluorescence intensity values of DHE staining are shown as in A. C, CGNs were treated with 50 µM MPP+ in the presence and absence of 1 mM DAHP for different time periods as indicated, and superoxide formation was measured by DHE staining. D, mean fluorescence intensity values of DHE staining as in C. The fluorescence images are representative of three independent experiments, and the intensity values obtained using a Metamorph software are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; ***, p < 0.001 compared with control; #, p < 0.05 compared with MPP+ treatment.

 
Effect of BH4 Repletion on MPP+-induced ROS Generation and Apoptosis in CGNs—Sepiapterin treatment has been shown to increase intracellular BH4 levels (28, 54). Intracellular BH4 concentration in CGNs increased more than 10-fold after incubation with 40 µM sepiapterin for 24 h (Table I). This increase was completely reversed by pretreatment with 100 µM sepiapterin reductase inhibitor, NAS. Intracellular BH4 levels decreased by 50% after treatment of cells with 70 µM MPP+ for 24 h. In sepiapterin-treated cells, however, BH4 levels remained much higher even after MPP+ treatment. This suggests that MPP+ does not inhibit the activity of enzymes (e.g. sepiapterin reductase) involved in the de novo and salvage pathway in BH4 synthesis. MPP+-induced intracellular ROS formation was assessed in sepiapterin-treated cells. As shown in Fig. 4 (A and B), there was a significant decrease in superoxide generation by 70 µM MPP+ (70 µM) in the presence of 40 µM sepiapterin, as compared with MPP+ treatment alone. This decrease in superoxide formation was negated by co-incubation with 100 µM NAS, suggesting that BH4 formed from sepiapterin, and not sepiapterin per se, was responsible for decreasing MPP+-dependent superoxide formation. Interestingly, pretreatment of CGNs with iron chelators (deferoxamine and HBED) for 2 h significantly inhibited MPP+-induced superoxide formation. This result suggests a causal relationship between intracellular iron and BH4 levels.


View this table:
[in this window]
[in a new window]
  		TABLE I
The effect of sepiapterin on intracellular BH4 levels

CGNs were treated with MPP+ (70 µM), and sepiapterin (40 µM) with or without NAS (100 µM) for 24 h. Intracellular BH4 was determined by HPLC. Data obtained from three independent experiments are represented as means ± S.D. *, p < 0.05 compared with control. **, p < 0.001 compared with control. ***, p < 0.001 compared with MPP+ treatment.

 



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 4.
The effect of varying intracellular BH4 levels and iron chelators on MPP+-induced oxidative stress in CGNs. A-D, CGNs were treated with 70 µM MPP+ alone or in the presence 40 µM sepiapterin or 10 µM iron chelators for 24 h. All compounds were preincubated for 2 h prior to the treatment with MPP+. 100 µM NAS was used to inhibit sepiapterin reductase. Intracellular ROS were determined by DHE-derived red fluorescence (for superoxide formation) (A) and DCFH-derived green fluorescence (C) as described under "Experimental Procedures." B, mean fluorescence intensity values of DHE red fluorescence obtained in A; D, mean fluorescence intensity values of DCF green fluorescence obtained in C. The fluorescence images are representative of three independent experiments, and the intensity values obtained using Metamorph software are means ± S.D. of three separate experiments. **, p < 0.01 compared with control; ***, p < 0.001 compared with control; #, p < 0.05 compared with MPP+ treatment; ##, p < 0.01 compared with MPP+ treatment.

 
Recently, we reported that the oxidation of DCFH, a non-fluorescent probe, to a fluorescent product, DCF, is dependent on the intracellular H2O2 and redox-active iron (55, 56). DCF staining in Fig. 4 (C and D) shows that pretreatment of cells with 40 µM sepiapterin significantly decreased MPP+-induced increased DCF staining, which was reversed by pretreatment with 100 µM NAS. Pretreatment of cells with iron chelators (deferoxamine and HBED) for 2 h greatly inhibited MPP+-induced DCF staining (Fig. 4, C and D). Based on these observations, we conclude that sepiapterin treatment inhibits MPP+-induced TfR iron accumulation in cells.

If depletion of BH4 content plays a role in MPP+-induced toxicity, then sepiapterin ought to protect against MPP+-induced apoptosis. Treatment of CGNs with 70 µM MPP+ increased the fraction of TUNEL-positive CGNs from 6 to 55% (Fig. 5, A and B, a and b). The fraction of TUNEL-positive nuclei was significantly decreased in cells treated with MPP+ and 40 µM sepiapterin (Fig. 5, A and B, b and c). As we reported previously (36), exposure of CGNs to 70 µM MPP+ for 24 h increased caspase-3-like protease activity by 3-fold (Fig. 5C). In the presence of 40 µM sepiapterin, MPP+-induced caspase-3-like protease activation was significantly inhibited. After incubating with MPP+ for 48 h, the cell viability was assayed by the MTT assay; 70 µM MPP+ treatment decreased cell viability to 47%, and sepiapterin increased the cell viability from 47 to 80% (Fig. 5D). The protective effects of sepiapterin in DNA fragmentation, caspase-3 activation, and cell viability were all reversed by pretreatment with 100 µM NAS. This further confirms our proposal that BH4, derived from sepiapterin, can reverse MPP+-induced toxicity in CGNs.



View larger version (24K):
[in this window]
[in a new window]
  		FIG. 5.
The effect of varying intracellular BH4 levels on MPP+-induced apoptosis in CGNs. A, CGNs were treated with 70 µM MPP+ in the presence or absence of 40 µM sepiapterin for 36 h. Cells were then fixed, stained for TUNEL-positive cells, and examined by fluorescence microscopy as described under "Experimental Procedures." B, percentage of apoptosis calculated using Metamorph software. C, CGNs were treated with 70 µM MPP+ in the presence or absence of 40 µM sepiapterin for 24 h. Cells were then collected, and the caspase-3 activity was measured. D, CGNs were treated with 70 µM MPP+ in the presence or absence of 40 µM sepiapterin for 48 h, and the cell viability was measured by the MTT assay. These experiments were also performed in the presence of 100 µM NAS, which inhibits sepiapterin conversion to BH4. The fluorescence images are representative of three independent experiments, and values shown are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; #, p < 0.05 compared with MPP+ treatment.

 
The Mechanism of BH4 Depletion in MPP+-treated CGNs—As data shown in Fig. 4 (A and B) suggest a causal relationship between intracellular iron and BH4 levels, the effects of cell-permeable iron chelators on intracellular BH4 levels in MPP+-treated cells were determined. Fig. 6A shows that preincubation of cells with deferoxamine or HBED inhibited BH4 depletion in MPP+-treated CGNs, indicating that intracellular redox-active iron might play an important role in BH4 depletion. To examine further the involvement of TfR-dependent iron in BH4 depletion, the monoclonal (IgA) anti-TfR antibody (42/6) was used to specifically bind to the extracellular domain of the TfR and block receptor endocytosis. In the presence of 42/6, iron can not enter the cell through TfR. Fig. 6B shows that the presence of 12 µg/ml 42/6 inhibited MPP+-induced BH4 depletion. These results suggest that MPP+-induced BH4 depletion likely occurs through a TfR-dependent iron transport mechanism.



View larger version (12K):
[in this window]
[in a new window]
  		FIG. 6.
The mechanism of MPP+-induced BH4 depletion in CGNs. A, CGNs were treated with 70 µM MPP+ and 10 µM iron chelator (with pretreatment for 2 h) for indicated time periods. B, CGNs were treated with 70 µM MPP+ and 12 µg/ml anti-TfR anti-body for 24 h. Cells were then collected, and intracellular BH4 levels were determined using the HPLC method described under "Experimental Procedures." C, CGNs expressed mRNA for rat GTPCH I as determined by the RT-PCR method. D, CGNs were treated with 70 µM MPP+ for different time periods as indicated, and the effect of MPP+ on GTPCH I mRNA level was determined by the semiquantitative RT-PCR method. E, CGNs were treated with 70 µM MPP+ for different time intervals, cells were collected, and GTPCH I activity determined using the HPLC method described under "Experimental Procedures." The gels are representative of three independent experiments. Data shown are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; #, p < 0.05 compared with MPP+ treatment.

 
The effect of MPP+ on BH4 synthesis was assessed by monitoring the expression of GTPCH I, the rate-limiting enzyme in de novo BH4 synthesis, in CGNs (Fig. 6, C and D). GTPCH I mRNA was expressed in CGNs, and the level of expression was not affected by 70 µM MPP+ treatment up to 36 h (Fig. 6D). The activity of GTPCH I in MPP+-treated CGNs was then monitored. Fig. 6E shows that 70 µM MPP+ treatment up to 48 h did not decrease GTPCH I activity.

Effect of BH4 Repletion on MPP+-induced Changes in nNOS Activity, Aconitase Activity, TfR Protein Expression, and nNOS Dimer Formation—MPP+ (70 µM) treatment for 24 h decreased the nNOS activity as determined by nitrite levels. Pretreatment with 40 µM sepiapterin restored the nNOS activity (Fig. 7A). Note that sepiapterin treatment itself did not increase nNOS activity, although sepiapterin significantly increased intracellular BH4 level (Table I). This suggests that, under normal conditions, nNOS is saturated by endogenous BH4. Similarly, pretreatment of CGNs with 40 µM sepiapterin inhibited aconitase inactivation caused by 70 µM MPP+ treatment for 24 h (Fig. 7B). Concomitantly, MPP+-induced increased cellular TfR protein level was markedly decreased by 40 µM sepiapterin pretreatment (Fig. 7, C and D). The effects of sepiapterin on aconitase activity and TfR protein expression were reversed in the presence of 100 µM NAS. These findings suggest that increased intracellular BH4 levels by sepiapterin down-regulate MPP+-induced TfR expression by inhibiting aconitase inactivation.



View larger version (28K):
[in this window]
[in a new window]
  		FIG. 7.
The effect of varying intracellular BH4 levels on MPP+-induced changes in nNOS activity, aconitase activity, TfR, and nNOS expression in CGNs. A, CGNs were treated with 70 µM MPP+ alone or in the presence of 40 µM sepiapterin, or 100 µM L-NAME for 24 h. nNOS activity was determined by nitrite measurements. B and C, CGNs were treated with 70 µM MPP+ in the presence or absence of 40 µM sepiapterin for 24 h. Cells were collected, and the total aconitase activity (B) and TfR protein levels (C) were measured, D, densitometric analyses of C. E, CGNs were treated with 70 µM MPP+ in the presence or absence of 40 µM sepiapterin for 24 h, and nNOS dimer levels were measured by LT-PAGE and Western blotting. F, densitometric analysis of nNOS dimer in E. G, CGNs were pretreated with sepiapterin (40 µM) or HBED (10 µM) for 2 h, and 70 µM MPP+ was then added and incubated for 24 h. The nNOS dimer was monitored by LT-PAGE and Western blotting. H, densitometric analysis of nNOS dimer in G. These experiments were also performed in the presence of 100 µM NAS, which was used to inhibit BH4 formation from sepiapterin. The gels are representative of three independent experiments. Values are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; #, p < 0.05 compared with MPP+ treatment.

 
We examined the effect of BH4 supplementation on nNOS dimer formation. Fig. 7 (E and F) shows that MPP+ (70 µM, 24 h) treatment decreased nNOS dimer, which was reversed by pretreatment of cells with 40 µM sepiapterin. Interestingly, sepiapterin treatment itself slightly increased nNOS dimer formation, although we did not investigate this in detail. The iron chelator, HBED, also moderately increased nNOS dimer formation (Fig. 7, G and H), consistent with its protective effect on BH4 depletion.

Effect of nNOS Specific Inhibitor on MPP+-induced Iron Uptake, Apoptosis, and BH4 Depletion in CGNs—7-Nitroindazole (7-NI) is a specific inhibitor for nNOS and was used to investigate the role of .NO in MPP+-induced toxicity in CGNs. Fig. 8A shows that treatment of CGNs with the combination of 7-NI (50 µM) and MPP+ (50 µM) significantly increased caspase-3 activity, compared with 7-NI or MPP+ alone. Treatment of CGNs with 50 µM MPP+ slightly decreased the cell viability after 48 h, whereas in the presence of 50 µM 7-NI, the percentage of cell death increased from 22 to 78% (Fig. 8B). The protective effects of sepiapterin on caspase-3 activity and cell viability in MPP+-treated cells were nullified by 7-NI. 7-NI (50 µM) itself was toxic to cells after 48 h, and this toxicity was not reversed by pretreatment of cell with 40 µM sepiapterin. These findings indicate that the protective effect of sepiapterin is not simply the result of the antioxidant effect of BH4. We examined the effect of 7-NI on TfR protein expression in MPP+-treated cells. Fig. 8 (C and D) shows that TfR protein expression was slightly but not significantly increased in the presence of 50 µM MPP+ or 50 µM 7-NI treatment alone, but was significantly increased by combined treatment. The DCF fluorescence analysis showed that co-treatment of cells with 50 µM MPP+ and 50 µM 7-NI for 24 h significantly increased DCF fluorescence when compared with the treatment with MPP+ or 7-NI alone (Fig. 8E). 7-NI treatment counteracted the reduction in both TfR protein level and DCF fluorescence in the presence of sepiapterin.



View larger version (33K):
[in this window]
[in a new window]
  		FIG. 8.
The effect of nNOS specific inhibitor on MPP+-induced iron uptake, apoptosis, and BH4 depletion in CGNs. A, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI in the presence or absence of 40 µM sepiapterin for 24 h. Cells were then collected and the caspase-3 like activity measured. B, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI in the presence or absence of 40 µM sepiapterin for 48 h. Cell viability was determined by the MTT assay. C, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI in the presence or absence of 40 µM sepiapterin for 24 h. Cell lyates were harvested and the TfR protein levels determined by Western blotting as described under "Experimental Procedures." D, densitometric analysis of TfR protein levels shown in C. E, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI in the presence or absence of 40 µM sepiapterin for 24 h. Intracellular oxidative stress was measured by DCF green fluorescence as described under "Experimental Procedures." F, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI for 24 h, and 0.2 µCi/ml 55Fe was then added and further incubated for 4 h. The 55Fe content in cell lysates was measured. G, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI for 24 h, and the intracellular BH4 levels were measured by HPLC as described under "Experimental Procedures." The fluorescence images are representative of three independent experiments. Values shown are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; **, p < 0.01 compared with control; ***, p < 0.001 compared with control; #, p < 0.05 compared with MPP+ treatment; ##, p < 0.01 compared with MPP+ treatment; ###, p < 0.001 compared with MPP+ treatment.

 
The effect of 7-NI on MPP+-induced 55Fe uptake was examined in CGNs. As shown in Fig. 8F, 55Fe uptake greatly increased in the presence of both 50 µM MPP+ and 50 µM 7-NI, when compared with MPP+ or 7-NI alone. These findings suggest that .NO plays an important role in maintaining the homeostasis of cellular iron levels. The effect of 7-NI on MPP+-induced alterations in intracellular BH4 was examined. As shown in Fig. 8G, intracellular BH4 concentration was significantly lower in cells treated with 7-NI (50 µM) and 50 µM MPP+ than in cells treated with either agent alone. These findings indicate a close correspondence between the intracellular iron, BH4 levels, and .NO, and that MPP+-induced toxicity in CGNs is governed by all of these factors.

Effect of MPP+ on Proteasome Activity in CGNs—We reported previously that oxidative stress decreased both chymotrypsin-like and trypsin-like activities of the 26 S proteasome (37). Impairment in the proteasome system has been implicated in the pathogenesis of PD (1). Thus, we sought to measure the proteasome activities in MPP+-treated CGNs. Fig. 9 (A and B) shows that 70 µM MPP+ treatment decreased the trypsin-like and chymotrypsin-like activities in a time-dependent manner. We then investigated the effect of sepiapterin on MPP+-induced changes in proteasome activity. As shown in Fig. 9 (C and D), 40 µM sepiapterin alone significantly increased both trypsin-like and chymotrypsin-like activities in CGNs. The enzyme activities remained elevated in cells during the combined treatment of MPP+ and sepiapterin. Although enzyme activities only slightly decreased in the presence of 50 µM MPP+ or 50 µM 7-NI treatment alone, they decreased significantly by both 50 µM MPP+ and 7-NI treatment. The protective effect of sepiapterin on proteasome activities in MPP+-treated cells is also antagonized by 7-NI treatment (Fig. 9, C and D). These data confirm our previous report that .NO plays an important role in regulating the proteasomal activity (37), indicating that the protective effect of sepiapterin is partially caused by its ability to increase the proteasome activity by increasing BH4 and nNOS activity and generating more .NO.



View larger version (24K):
[in this window]
[in a new window]
  		FIG. 9.
The effect of sepiapterin on proteasomal activities in CGNs treated with MPP+ A and B, CGNs were treated with 70 µM MPP+ for the time period indicated. Cells were collected, and the trypsin-like (A) and chymotrypsin-like (B) activities of the 26 S proteasome were measured as described under "Experimental Procedures." C and D, CGNs were treated with 50 µM MPP+ and/or 50 µM 7-NI in the presence or absence of 40 µM sepiapterin for 24 h, and the trypsin-like (C) and chymotrypsin-like (D) activities were measured. Values shown are means ± S.D. of three separate experiments. *, p < 0.05 compared with control; ***, p < 0.01 compared with control; #, p < 0.05 compared with MPP+ treatment; ##, p < 0.01 compared with MPP+ treatment.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we made the following key observations. (i) MPP+ generated mitochondrially derived ROS that stimulated TfR levels and iron signaling. (ii) MPP+-induced H2O2 and Tf-iron were responsible for degrading intracellular BH4 levels. BH4 deficiency stimulated nNOS uncoupling, leading to increased superoxide and H2O2 formation and decreased .NO formation. (iii) Pretreatment with anti-TfR antibody, cell-permeable iron chelators, or sepiapterin to increase intracellular BH4 all abrogated MPP+-induced oxidative stress and apoptotic cell death. Scheme 1 summarizes the involvement of ROS derived from both mitochondria and nNOS uncoupling mechanism in MPP+-mediated toxicity using CGNs as a cellular model.



View larger version (18K):
[in this window]
[in a new window]
  		SCHEME 1.
A proposed model for the role of nNOS in MPP+-induced toxicity in CGNs. MPP+-induced ROS inactivates the aconitase enzyme, which in turn increases the levels of non-iron-bound IRP1, enabling it to bind to the 3'-untranslated region of TfR mRNA, thereby enhancing its stability and level of expression. Elevation of TfR expression enhances the cellular uptake of iron, decreases BH4 levels and nNOS dimer form, resulting in more ROS and less .NO generation by NOS uncoupling, and ultimately results in apoptosis. Increasing intracellular BH4 levels by sepiapterin treatment lowers MPP+-induced intracellular ROS generation, increases .NO generation, and prevents the accumulation of TfR-Fe, thereby preventing apoptosis.

 
Oxidant-induced Iron Signaling—As alluded to in the Introduction, oxidant-induced iron signaling propagates free radical formation in cells. A major portion of cellular iron uptake is utilized for assembly of iron-sulfur clusters and cytochrome synthesis in mitochondria. Inactivation of aconitase stimulates cellular iron signaling mechanisms (47-49). A major pathway by which neuronal cells acquire iron is via TfR, which facilitates the uptake of iron-loaded transferrin (Tf-Fe). The cellular iron sensing mechanism is triggered by iron deficiency or when the 4Fe-4S cluster in aconitase is disassembled in the presence of ROS and RNS (49). Under conditions of iron deprivation or aconitase inactivation during oxidative stress, the IRPs bind with a high affinity to IRE present on 3'- and 5'-untranslated regions on TfR and ferritin mRNAs, respectively. The increased binding to TfR mRNA stabilizes the mRNA, leading to enhanced mRNA translation and increased TfR synthesis (49). As shown in Fig. 1, CGNs exposed to MPP+ elicit an increased TfR synthesis and Tf-Fe uptake via a similar mechanism. Pretreatment of CGNs with anti-TfR antibody inhibited MPP+-induced ROS and apoptosis, supporting a causative role for Tf-Fe in MPP+ neurotoxicity (36).

BH4 Regulates NOS Activity: Uncoupling of NOS—Results indicate that MPP+-induced ROS and Tf-Fe signaling are responsible for the depletion of intracellular BH4 that was accompanied by a decrease in the nNOS activity and an increase in superoxide formation because of nNOS uncoupling. As shown in Fig. 10, BH4, a critical co-factor for NOS activity, acts as a redox switch to the oxygenase domain of NOS (20, 57). The catalytic domain of nNOS consists of an NADPH-binding reductase, a heme-containing oxygenase domain that binds L-arginine, BH4, and a calmodulin-binding sequence (57, 58). In the "resting state," there is no flow of electrons from the reductase to the oxygenase domain. Upon binding of the calcium-calmodulin complex, electrons flow from the reductase domain to the oxygenase domain, resulting in the activation of the enzyme by reducing heme-iron (III) to iron (II) (Fig. 10). Molecular oxygen binds to the ferrous-heme group, forming the heme-iron (IV)-oxo complex, that oxidizes L-arginine to L-citrulline and .NO. BH4 is, however, essential for NOS activation as it stabilizes the oxoferryl state, preventing its disassociation to superoxide. In the presence of BH4, there is negligible superoxide formation from NOS.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 10.
Regulation of nNOS activity by BH4. In the presence of BH4 (+BH4), oxyferryl complex (Heme-OOH) oxidizes L-arginine to L-citrulline and .NO. In the absence of BH4 or BH4 deficiency (-BH4), mostly superoxide and reactive nitrogen species are formed.

 
The present results demonstrate that MPP+ treatment causes a time-dependent depletion of BH4 in CGNs because of the combined action of increased Tf-Fe and H2O2. Use of sepiapterin to selectively reverse the effect of MPP+ on BH4 content allowed us to conclude that MPP+-induced intracellular ROS generation, Tf-Fe signaling, and caspase-3 activation are a result of decreased BH4 content. Treatment with N-acetylserotonin, inhibitor of sepiapterin reductase, reversed the inhibitory effects of sepiapterin on the various aspects of MPP+ toxicity. Thus, the observed protection against MPP+ toxicity by sepiapterin is attributed to increased intracellular BH4. It is our current hypothesis that this is the result of prevention of NOS uncoupling, resulting in restoration of appropriate NOS activity and inhibition of superoxide-forming oxidase activity.

It has been previously reported that BH4 stabilizes the functionally active, dimeric form of nNOS (42, 59). BH4 also inhibits monomerization of nNOS, as well as the inactivation of the enzyme (60). We have shown that nNOS monomer generates ROS in lieu of .NO and L-citrulline in the absence of BH4 (21). Furthermore, the ratio between BH4 and oxidized BH4 was reported to control the formation of superoxide from endothelial NOS (61). Exposure of high glucose to bovine aortic endothelial cells was reported to increase the formation of superoxide (51). Under these conditions, endothelial NOS dimers were decreased (51). Increased formation in bovine aortic endothelial cells treated with high glucose was associated with increased formation of endothelial NOS monomers (51). Undoubtedly, the cofactor and substrate influence on NOS dimerization, stability, and regulation is more complex, and the degradation of nNOS in a cellular milieu is governed by many regulatory processes including protein-protein interactions and ubiquitin-proteasome pathways (62-65). The present work clearly shows that BH4 depletion in CGNs treated with MPP+ was accompanied by a decrease in nNOS homodimer, along with an increase in the nNOS monomer/dimer ratio. Sepiapterin treatment increased the dimeric form of nNOS in CGNs treated with MPP+. Thus, the protective effect of sepiapterin against MPP+-mediated neurotoxicity is presumably linked to its conversion to BH4, which stabilizes the nNOS dimer.

The present data show that cell-permeable iron chelators inhibit MPP+-mediated BH4 depletion in CGNs (Fig. 4E). Pretreatment of CGNs with TfR antibody also prevented BH4 degradation (Fig. 4F). These findings suggest that Tf-Fe is responsible for BH4 depletion. Previous reports indicate that both ROS and RNS can react with BH4 and initiate a free radical chain reaction mechanism forming dihydrobiopterin (two-electron oxidation product) and pterine (four-electron oxidation product) (44). It is conceivable that a higher oxidant (hydroxyl radical or perferryl iron) formed from intracellular iron and H2O2 is responsible for the depletion of BH4 in MPP+-treated CGNs.

.NO Controls Iron Homeostasis—The present data show that 7-NI (a selective inhibitor of nNOS) exacerbated MPP+-induced iron uptake, oxidative stress, BH4 depletion, and apoptosis in CGNs (Fig. 8). We have recently shown that .NO regulates oxidant-induced iron signaling, intracellular iron homeostasis, and oxidative stress through increased proteolytic activation (37). NOS inhibitors decreased the proteasomal activity and enhanced TfR-dependent iron signaling. Nitration of tyrosines present in TfR has been proposed to modulate iron signaling (37). An increase in protein nitration was detected in the presence of .NO, H2O2, and iron (66). Peroxynitrite formation decreased .NO levels and enhanced the iron-mediated toxicity while decreasing the proteasomal function. MPP+-mediated neurotoxicity is regulated by Tf-iron signaling that is dependent on intracellular BH4.

Implications in Neurodegenerative Diseases—Free radical damage has been implicated in age-related neurodegenerative diseases like PD and Alzheimer's disease (1, 9, 67, 68), which are marked by a progressive motor dysfunction resulting from selective loss of nigral cells. Iron plays a critical role in initiating and amplifying oxidative damage in tissues (69). Iron levels are elevated in the substantia nigra of PD patients, and recently Kaur et al. (70) demonstrated that iron chelation by overexpression of the iron binding protein ferritin or by administering an iron chelator (e.g. clioquinol) to mice significantly inhibited MPTP-induced oxidative damage in dopaminergic cells and behavioral deficits. The present results provide new insights into the mechanisms of iron accumulation and ROS formation in CGNs treated with a mitochondrial Parkinsonism-inducing toxin. PD is characterized by impaired nNOS activity and decreased levels of BH4 and dopamine (2, 15).

This study reveals an important link between Tf-iron uptake, BH4 depletion, and enhanced superoxide formation from uncoupled nNOS and demonstrates how nNOS could play a crucial role in eliciting the neuronal apoptosis by MPP+ through oxidant-induced iron signaling. Reactive nitrogen species have been implicated in the pathogenesis of age-related neurodegenerative mitochondrial diseases (1, 9). Recently, we reported that MPP+-induced {alpha}-synuclein aggregation in neuroblastoma cells might be the result of oxidants generated from H2O2 and TfR-dependent iron (71). It is likely that the novel molecular mechanism of toxicity proposed here for MPP+ and for other mitochondrial neurotoxins, in general, may lead to the development of new neuroprotective strategies for PD and other age-related chronic neurological disorders.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grants NS39958 and IPOIHL68769-01 and by the Parson's Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Dept. of Biophysics, Medical College of Wisconsin, 8701 Watertown Plank Rd., Milwaukee, WI 53226. Tel.: 414-456-4035; Fax: 414-156-6512; E-mail: balarama{at}mcw.edu.

1 The abbreviations used are: PD, Parkinson's disease; BH4, tetrahydrobiopterin; CGN, cerebellar granule neuron; DAHP, 2,4-diamino-6-hydroxypyrimidine; DCF, 2',7'-dichlorofluorescein; DCFH, 2',7'-dichlorodihydrofluorescein; DHE, dihydroethidium; GTPCH I, GTP cyclohydrolase I; HBED, N,N'-bis(2-hydroxybenzyl ethylenediamine-N,N'-diacetic acid; IRP, iron regulatory protein; IRE, iron-responsive element; L-NAME, NG-nitro-L-arginine-methyl ester; L-NMMA, NG-monomethyl-L-arginine monoacetate; MPP+, 1-methyl-4-phenylpyridinium; 7-NI, 7-nitroindazole; MTT, 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl tetrazolium bromide; NAS, N-acetylserotonin; nNOS, neuronal nitric-oxide synthase; TfR, transferrin receptor; NOS, nitric-oxide synthase; ROS, reactive oxygen species; RNS, reactive nitrogen species; MPTP, 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine; HPLC, high performance liquid chromatography; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; LT, low temperature SDS; RT, reverse transcription; carboxy-H2DCFDA, carboxy-2',7'-dichlorodihydrofluorescein diacetate; TUNEL, terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling; Boc, butoxycarbonyl; Suc, succinyl; AMC, 7-amido-4-methylcoumarin; CSS, control salt solution; PBS, phosphate-buffered saline; DTT, dithiothreitol. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Jeannette Vásquez-Vivar for providing valuable assistance on the measurement of BH4 and GTPCH I activity.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Dawson, T. M., and Dawson, V. L. (2003) Science 302, 819-822[Abstract/Free Full Text]
  2. Lovenberg, W., Levine, R. A., Robinson, D. S., Ebert, M., Williams, A. C., and Calne, D. B. (1979) Science 204, 624-626[Abstract/Free Full Text]
  3. Spillantini, M. G., Crowther, R. A., Jakes, R., Hasegawa, M., and Goedert, M. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 6469-6473[Abstract/Free Full Text]
  4. Bonn, D. (1996) Lancet 347, 1614[CrossRef][Medline] [Order article via Infotrieve]
  5. Lagnston, J. W., Ballard, P., Tetrud, J. W., and Irwin, I. (1983) Science 219, 979-980[Abstract/Free Full Text]
  6. Burns, R. S., Chiueh, C. C., Markey, S. P., Ebert, M. H. E., Jacobowitz, D. M., and Kopin, I. J. (1984) Proc. Natl. Acad. Sci. U. S. A. 80, 4546-4550
  7. Langston, J. W., Irwin, I., Langston, E. B., and Forno, L. S. (1984) Neurosci. Lett. 48, 87-92[Medline] [Order article via Infotrieve]
  8. Javitch, J. A., D'Amato, R. J., Strittmatter, S. M., and Snyder, S. H. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 2173-2177[Abstract/Free Full Text]
  9. Metodiewa, D., and Koska, C. (2000) Neurotox. Res. 1, 197-233[Medline] [Order article via Infotrieve]
  10. Matthews, R. T., Beal, M. F., Fallon, J., Fedorchak, K., Huang, P. L., Fishman, M. C., and Hyman, B. T. (1997) Neurobiol. Dis. 4, 114-121[CrossRef][Medline] [Order article via Infotrieve]
  11. Stewart, V. C., and Heales, S. J. R. (2003) Free Radical Biol. Med. 34, 287-303[CrossRef][Medline] [Order article via Infotrieve]
  12. Mohanakumar, K. P., Thomas, B., Sharma, S. M., Muralikrishnan, D., Chowdhury, R., and Chiueh, C. C. (2002) Ann. N. Y. Acad. Sci. 962, 389-401[Medline] [Order article via Infotrieve]
  13. Muramatsu, Y., Kurosaki, R., Mikami, T., Michimata, M., Matsubara, M., Imai, Y., Kato, H., Itoyama, Y., and Araki, T. (2002) Metab. Brain Dis. 17, 169-182[CrossRef][Medline] [Order article via Infotrieve]
  14. Barc, S., Page, G., Barrier, L., Piriou, A., and Fauconneau, B. (2001) Neurosci. Lett. 314, 82-86[CrossRef][Medline] [Order article via Infotrieve]
  15. Kuiper, M. A., Visser, J. J., Bergmans, P. L., Scheltens, P., and Wolters, E. C. (1994) J. Neurol. Sci. 121, 46-49[CrossRef][Medline] [Order article via Infotrieve]
  16. Nagatsu, T., Yamaguchi, T., Kato, T., Sugimoto, T., Matsuura, S., Akino, M., Nagatsu, I., Iizuka, R., and Narabayashi, H. (1981) Clin. Chim. Acta 109, 305-311[CrossRef][Medline] [Order article via Infotrieve]
  17. Nagatsu, T., Yamaguchi, T., Rahman, M. K., Trocewicz, J., Oka, K., Hirata, Y., Nagatsu, I., Narabayashi, H., Kondo, T., and Iizuka, R. (1984) Adv. Neurol. 40, 467-473[Medline] [Order article via Infotrieve]
  18. Tayeh, M. A., and Marletta, M. A. (1989) J. Biol. Chem. 264, 19654-19658[Abstract/Free Full Text]
  19. Schmidt, K., Werner, E. R., Mayer, B., Wachter, H., and Kukovetz, W. R. (1992) Biochem. J. 281, 297-300[Medline] [Order article via Infotrieve]
  20. Hemmens, B., and Mayer, B. (1998) Methods Mol. Biol. 100, 1-32[Medline] [Order article via Infotrieve]
  21. Vasquez-Vivar, J., Hogg, N., Martasek, P., Karoui, H., Pritchard, K. A., Jr., and Kalyanaraman, B. (1999) J. Biol. Chem. 274, 26736-26742[Abstract/Free Full Text]
  22. Xia, Y., Tsai, A. L., Berka, V., and Zweier, J. L. (1998) J. Biol. Chem. 273, 25804-25808[Abstract/Free Full Text]
  23. Cho, S., Volpe, B. T., Bae, Y., Hwang, O., Choi, H. J., Gal, J., Park, L. C. H., Chu, K., Du, J., and Joh, T. H. (1999) J. Neurosci. 19, 878-889[Abstract/Free Full Text]
  24. Tiefenbacher, C. P., Chilian, W. M., Mitchell, M., and DeFily, D. V. (1996) Circulation 94, 1423-1429[Abstract/Free Full Text]
  25. Tiefenbacher, C. P., Lee, C. H., Kapitza, J., Dietz, V., and Niroomand, F. (2003) Pflugers Arch. 447, 1-7[CrossRef][Medline] [Order article via Infotrieve]
  26. Delgado-Esteban, M., Almeida, A., and Medina, J. M. (2002) J. Neurochem. 82, 1148-1159[CrossRef][Medline] [Order article via Infotrieve]
  27. Ihlemann, N., Rask-Madsen, C., Perner, A., Dominguez, H., Hermann, T., Kober, L., and Torp-Pedersen, C. (2003) Am. J. Physiol. 285, H875-H882
  28. Thony, B., Auerbach, G., and Blau, N. (2000) Biochem. J. 347, 1-16[CrossRef][Medline] [Order article via Infotrieve]
  29. Nichol, C. A., Smith, G. K., and Duch, D. S. (1985) Annu. Rev. Biochem. 54, 729-764[CrossRef][Medline] [Order article via Infotrieve]
  30. Bishop, G. M., Robinson, S. R., Liu, Q., Perry, G., Atwood, C. S., and Smith, M. A. (2002) Dev. Neurosci. 24, 184-187[CrossRef][Medline] [Order article via Infotrieve]
  31. Sipe, J. C., Lee, P., and Beutler, E. (2002) Dev. Neurosci. 24, 188-196[CrossRef][Medline] [Order article via Infotrieve]
  32. Lan, J., Jiang, D. H. (1997) J. Neural Transm. 104, 649-660[CrossRef][Medline] [Order article via Infotrieve]
  33. Mochizuki, H., Imai, H., Endo, K., Yokomizo, K., Murata, Y., Hattori, N., and Mizuno, Y. (1994) Neurosci. Lett. 168, 251-253[CrossRef][Medline] [Order article via Infotrieve]
  34. Powers, K. M., Smith-Weller, T., Franklin, G. M., Longstreth, W. T., Swanson, P. D., and Checkoway, H. (2003) Neurology 60, 1761-1766[Abstract/Free Full Text]
  35. Qian, Z. M., Pu, Y. M., Tang, P. L., and Wang, Q. (1998) Neurosci. Lett. 251, 9-12[CrossRef][Medline] [Order article via Infotrieve]
  36. Kalivendi, S. V., Kotamraju, S., Cunningham, S., Shang, T., Hillard, C. J., and Kalyanaraman, B. (2003) Biochem. J. 371, 151-164[CrossRef][Medline] [Order article via Infotrieve]
  37. Kotamraju, S., Tampo, Y., Keszler, A., Chitambar, C. R., Joseph, J., Haas, A. L., and Kalyanaraman, B. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 10653-10658[Abstract/Free Full Text]
  38. Ryu, S. Y., Jeong, K. S., Kang, B. N., Park, S. J., Yoon, W. K., Kim, S. H., and Kim, T. H. (2000) Anticancer Res. 20, 3331-3338[Medline] [Order article via Infotrieve]
  39. Mulero, V., and Brock, J. H. (1999) Blood 94, 2383-2389[Abstract/Free Full Text]
  40. Shang, T., Uihlein, A. V., Van Asten, J., Kalyanaraman, B., and Hillard, C. J. (2003) J. Neurochem. 85, 358-367[Medline] [Order article via Infotrieve]
  41. Hillard, C. J., Edgemond, W. S., Jarrahian, A., and Campbell, W. B. (1997) J. Neurochem. 69, 631-638[Medline] [Order article via Infotrieve]
  42. Klatt, P., Schmidt, K., Lehner, D., Glatter, O., Bachinger, H. P., and Mayer, B. (1995) EMBO J. 14, 3687-3695[Medline] [Order article via Infotrieve]
  43. Kennedy, M. C., Emptage, M. H., Dreyer, J. L., Beinert, H. (1983) J. Biol. Chem. 258, 11098-11105[Abstract/Free Full Text]
  44. Vasquez-Vivar, J., Whitsett, J., Martasek, P., Hogg, N., and Kalyanaraman, B. (2001) Free Radical Biol. Med. 31, 975-985[CrossRef][Medline] [Order article via Infotrieve]
  45. Weikert, S., Freyer, D., Weih, M., Isaev, N., Busch, C., Schultze, J., Megow, D., and Dirnagl, U. (1997) Brain Res. 748, 1-11[CrossRef][Medline] [Order article via Infotrieve]
  46. Canu, N., Barbato, C., Ciotti, M., T., Serafino, A., Dus, L., and Calissano, P. (2000) J. Neurosci. 20, 589-599[Abstract/Free Full Text]
  47. Hausladen, A., and Fridovich, I. (1996) Methods Enzymol. 269, 37-41[Medline] [Order article via Infotrieve]
  48. Konorov, E. A., Kennedy, M. C., and Kalyanaraman, B. (1999) Arch. Biochem. Biophys. 368, 421-428[CrossRef][Medline] [Order article via Infotrieve]
  49. Paraskeva, E., and Hentze, M. W. (1996) FEBS Lett. 389, 40-43[CrossRef][Medline] [Order article via Infotrieve]
  50. Hentze, M., Rouault, T. A., Harford, J. B., and Klausner, R. D. (1989) Science 244, 357-359[Abstract/Free Full Text]
  51. Zou, M. H., Shi, C., and Cohen, R. A. (2002) J. Clin. Invest. 109, 817-826[CrossRef][Medline] [Order article via Infotrieve]
  52. Rosen, G. M., Tsai, P., Weaver, J., Porasuphatana, S., Roman, L. J., Starkov, A. A., Fiskum, G., and Pou, S. (2002) J. Biol. Chem. 277, 40275-40280[Abstract/Free Full Text]
  53. Nakamura, K., Bindokas, V. P., Kowlessur, D., Elas, M., Milstien, S., Marks, J. D., Halpern, H. J., and Kang, U. J. (2001) J. Biol. Chem. 276, 34402-34407[Abstract/Free Full Text]
  54. Madsen, J. T., Jansen, P., Hesslinger, C., Meyer, M., Zimmer, J., and Gramsbergen, J. B. (2003) J. Neurochem. 85, 214-223[Medline] [Order article via Infotrieve]
  55. Kotamraju, S., Chitambar, C. R., Kalivendi, S. V., Joseph, J., and Kalyanaraman, B. (2002) J. Biol. Chem. 277, 17179-17187[Abstract/Free Full Text]
  56. Tampo, Y., Kotamraju, S., Chitambar, C. R., Kalivendi, S. V., Keszler, A., Joseph, J., and Kalyanaraman, B. (2003) Circ. Res. 92, 56-63[Abstract/Free Full Text]
  57. Vasquez-Vivar, J., Kalyanaraman, B., and Martasek, P. (2003) Free Radical Res. 37, 121-127[CrossRef][Medline] [Order article via Infotrieve]
  58. Griffith, O. W., and Stuehr, D. J. (1995) Annu. Rev. Physiol. 57, 707-736[CrossRef][Medline] [Order article via Infotrieve]
  59. Habisch, H. J., Gorren, A. C., Liang, H., Venema, R. C., Parkinson, J. F., Schmidt, K., and Mayer, B. (2003) Mol. Pharmacol. 63, 682-689[Abstract/Free Full Text]
  60. Reif, A., Frohlich, L. G., Kotsonis, P., Frey, A., Bommel, H. M., Wink, D. A., Pfleiderer, W., and Schmidt, H. H. (1999) J. Biol. Chem. 274, 24921-24929[Abstract/Free Full Text]
  61. Vasquez-Vivar, J., Martasek, P., Whitsett, J., Joseph, J., and Kalyanaraman, B. (2002) Biochem. J. 362, 733-739[CrossRef][Medline] [Order article via Infotrieve]
  62. Panda, K., Rosenfeld, R. J., Ghosh, S., Meade, A. L., Getzoff, E. D., and Stuehr, D. J. (2002) J. Biol. Chem. 277, 31020-31030[Abstract/Free Full Text]
  63. Kone, B. C., Kuncewicz, T., Zhang, W., and Yu, Z. Y. (2003) Am. J. Physiol. 285, F178-F190
  64. Song, Y., Cardounel, A. J., Zweier, J. L., and Xia, Y. (2002) Biochemistry 41, 10616-10622[CrossRef][Medline] [Order article via Infotrieve]
  65. Osawa, Y., Lowe, E. R., Everett, A. C., Dunbar, A. Y., and Billecke, S. S. (2003) J. Pharmacol. Exp. Ther. 304, 493-497[Abstract/Free Full Text]
  66. Thomas, D. D., Espey, M. G., Vitek, M. P., Miranda, K. M., and Wink, D. A. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 12691-12696[Abstract/Free Full Text]
  67. Perry, G., Nunomura, A., Hirai, K., Zhu, X., Perez, M., Avila, J., Castellani, R. J., Atwood, C. S., Aliev, G., Sayre, L. M., Takeda, A., and Smith, M. A. (2002) Free Radical Biol. Med. 33, 1475-1479[CrossRef][Medline] [Order article via Infotrieve]
  68. Behl, C., and Moosmann, B. (2002) Free Radical Biol. Med. 33, 182-191[CrossRef][Medline] [Order article via Infotrieve]
  69. Thomas, C. E., Morehouse, L. A., and Aust, S. D. (1985) J. Biol. Chem. 260, 3275-3280[Abstract/Free Full Text]
  70. Kaur, D., Yantiri, F., Rajagopalan, S., Kumar, J., Mo, J. Q., Boonplueang, R., Viswanath, V., Jacobs, R., Yang, L., Beal, M. F., DiMonte, D., Volitaskis, I., Ellerby, L., Cherny, R. A., Bush, A. I., and Andersen, J. K. (2003) Neuron 37, 899-909[CrossRef][Medline] [Order article via Infotrieve]
  71. Kalivendi, S. V., Cunningham, S., Kotamraju, S., Joseph, J., Hillard, C. J., and Kalyanaraman, B. (January 23, 2004) J. Biol. Chem. 10.1074/jbc. M400101200

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 Proc. Natl. Acad. Sci. USAHome page
H. Plun-Favreau and J. Hardy
PINK1 in mitochondrial function
PNAS, August 12, 2008; 105(32): 11041 - 11042.
[Full Text] [PDF]

Home page
 J BiochemHome page
F. Urano, N. Hayashi, F. Arisaka, H. Kurita, S. Murata, and H. Ichinose
Molecular mechanism for pterin-mediated inactivation of tyrosine hydroxylase: formation of insoluble aggregates of tyrosine hydroxylase.
J. Biochem., April 1, 2006; 139(4): 625 - 635.
[Abstract] [Full Text] [PDF]

Home page
 StrokeHome page
T. Nakamura, G. Xi, J.-W. Park, Y. Hua, J. T. Hoff, and R. F. Keep
Holo-Transferrin and Thrombin Can Interact to Cause Brain Damage
Stroke, February 1, 2005; 36(2): 348 - 352.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/18/19099    most recent
M400101200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Shang, T.
Right arrow Articles by Kalyanaraman, B.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Shang, T.
Right arrow Articles by Kalyanaraman, B.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement