HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 281, Issue 39, 28518-28528, September 29, 2006

This Article Abstract Full Text (PDF) Supplemental Data All Versions of this Article: 281/39/28518    most recent M604359200v1 Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Kanda, M. Articles by Kondo, T. Search for Related Content PubMed PubMed Citation Articles by Kanda, M. Articles by Kondo, T. Social Bookmarking                      What's this?

Glutaredoxin Modulates Platelet-derived Growth Factor-dependent Cell Signaling by Regulating the Redox Status of Low Molecular Weight Protein-tyrosine Phosphatase^*

Munetake Kanda¹, Yoshito Ihara¹², Hiroaki Murata, Yoshishige Urata, Takaaki Kono, Junji Yodoi^¶, Shinji Seto, Katsusuke Yano, and Takahito Kondo

From the Department of Biochemistry and Molecular Biology in Disease, Atomic Bomb Disease Institute and Department of Third Internal Medicine, Nagasaki University Graduate School of Biomedical Sciences, Nagasaki 852-8523, Japan and ^¶Department of Biological Responses, Institute for Viral Research, Kyoto University, Kyoto 606-8507, Japan

Received for publication, May 8, 2006 , and in revised form, June 20, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Glutaredoxin (GRX) is a glutathione-disulfide oxidoreductase involved in various cellular functions, including the redox-dependent regulation of certain integral proteins. Here we demonstrated that overexpression of GRX suppressed the proliferation of myocardiac H9c2 cells treated with platelet-derived growth factor (PDGF)-BB. After stimulation with PDGF-BB, the phosphorylation of PDGF receptor (PDGFR) was suppressed in GRX gene-transfected cells, compared with controls. Conversely, the phosphorylation was enhanced by depletion of GRX by RNA interference. In this study we focused on the role of low molecular weight protein-tyrosine phosphatase (LMW-PTP) in the dephosphorylation of PDGFR via a redox-dependent mechanism. We found that depletion of LMW-PTP using RNA interference enhanced the PDGF-BB-induced phosphorylation of PDGFR, indicating that LMW-PTP works for PDGFR. The enhancement of the phosphorylation of PDGFR was well correlated with inactivation of LMW-PTP by cellular peroxide generated in the cells stimulated with PDGF-BB. In vitro, with hydrogen peroxide treatment, LMW-PTP showed decreased activity with the concomitant formation of dithiothreitol-reducible oligomers. GRX protected LMW-PTP from hydrogen peroxide-induced oxidation and inactivation in concert with glutathione, NADPH, and glutathione disulfide reductase. This strongly suggests that retention of activity of LMW-PTP by enhanced GRX expression suppresses the proliferation of cells treated with PDGF-BB via enhanced dephosphorylation of PDGFR. Thus, GRX plays an important role in PDGF-BB-dependent cell proliferation by regulating the redox state of LMW-PTP.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The redox status of sulfhydryl groups in proteins plays an important role in the regulation of cellular functions such as the synthesis and folding of proteins and regulation of the structure and activity of enzymes, receptors, and transcription factors. To maintain the cellular thiol-disulfide redox status under reducing conditions, living cells possess two major systems, the thioredoxin (TRX)³/thioredoxin reductase system and the glutathione (GSH)/glutaredoxin (GRX) system (1).

GRX, also known as thiol transferase, was first discovered as a GSH-dependent hydrogen donor for ribonucleotide reductase in Escherichia coli mutants lacking TRX (2). GRX catalyzes the reduction of protein disulfide via a disulfide exchange reaction by utilizing the active site Cys-Pro-Tyr-Cys with a dithiol mechanism involving both active-site thiols (3). In addition, GRX has a unique ability to reduce protein-S-S-glutathione mixed disulfide (deglutathionylation) or to participate in its formation (glutathionylation) through a monothiol mechanism, which requires only the more N-terminal active site Cys (3, 4). Oxidized GRX is selectively recycled to the reduced form by GSH with the formation of glutathione disulfide (GSSG) and regeneration of GSH through coupling with NADPH and GSSG reductase, a system termed the GSH-regenerating system (5, 6). These characteristic interactions with GSH distinguish GRX from TRX, which favors intramolecular disulfide substrates and is turned over by NADPH and thioredoxin reductase independent of GSH. Functional overlap or cross-talk between the two systems, however, has been indicated (7, 8).

Mammalian GRX is known to have two isoforms, GRX1 and GRX2 (4). GRX1 is a cytosolic form of GRX. GRX1 and S-glutathionylation are thought to be involved in a variety of cellular events such as signal transduction, stress response, and metabolic regulation by regulating the redox status of various cellular proteins including human immunodeficiency virus-1 protease (9), glyceraldehyde-3-phosphate dehydrogenase (10), nuclear factor I (11), ASK1 (12), EphA2 kinase (13), actin (14), Ras (15), tubulin (16), tau and microtubule-associated protein-2 (17), annexin A2 (18), and protein-tyrosine phosphatase 1B (4, 19). We also have reported that GRX plays an important role in protecting cells from apoptosis by regulating the redox status of Akt/protein kinase B (20). The other form of GRX, GRX2, is distributed in the mitochondria and nucleus and also functions as part of a cellular antioxidant defense system by regulating the redox status of mitochondrial proteins such as complex I protein (21–23).

Platelet-derived growth factor (PDGF) is a major mitogen for connective tissue cells and certain other cell types. PDGF is a family of homo- and heterodimers of disulfide-bonded polypeptide chains A, B, C, and D (i.e. PDGF-AA, BB, CC, DD, and AB) (24). PDGFs bind to two cell-surface receptor-tyrosine kinases, PDGF receptor (PDGFR) and . The PDGF molecules are bivalent, and PDGF-dependent activation of receptors causes a mitogenic signal transduction through the phosphorylation of specific tyrosines in the receptors. The in vivo function of PDGF signaling has been studied with gene targeting experiments (25, 26). PDGF-BB or PDGFR knock-out mice die during late gestation from cardiovascular complications (27, 28), suggesting the importance of PDGF-BB/PDGFR signaling in the myocardiac development of mice. Although the physiological functions of PDGF-BB/PDGFR signaling in myocardiac cells are not fully understood, PDGFR is expressed in adult rat heart (29) and rat cardiomyocyte-derived H9c2 cells (30), and PDGF-BB/PDGFR signaling also controls the proliferation of neonatal rat cardiomyocytes (31). These reports suggest that the PDGF-BB/PDGFR signaling pathway plays an important role in the cellular physiology of myocardiac cells, although the precise regulatory mechanism for the signaling pathway and its relation to cellular redox-dependent regulation are not yet clarified.

In the present study we explored the effect of overexpression of GRX on PDGF-BB-dependent signaling and cell proliferation in myocardiac H9c2 cells. We found that overexpression of GRX suppressed the tyrosine phosphorylation of PDGFR after stimulation with PDGF-BB, resulting in a suppression of the PDGF-BB-dependent cell proliferation. Furthermore, we describe a novel regulatory mechanism for PDGF-BB signaling involving the redox-dependent regulation of low molecular weight protein-tyrosine phosphatase (LMW-PTP) by GRX in a GSH-dependent manner.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Antibodies and Reagents—Rabbit anti-mouse GRX polyclonal antibody was prepared as described before (20). Rabbit antibodies against PDGFR and phospho-(Tyr-751)-PDGFR were obtained from Cell Signaling Technology. Mouse anti-glyceraldehyde-3-phosphate dehydrogenase monoclonal antibody and protein-tyrosine phosphatase inhibitor III (PTPI-III; -bromo-4-(carboxymethoxy)-acetophenone) were from Chemicon (Temecula, CA). Sheep anti-LMW-PTP polyclonal antibody for immunoprecipitation was from Exalpha Biologicals (Watertown, MA). Rabbit anti-Myc tag polyclonal antibody was purchased from Upstate%20Biotechnology">Upstate Biotechnology. Anti-c-Myc monoclonal antibody-conjugated agarose beads were obtained from BD Biosciences Clontech. ¹²⁵I-Labeled PDGF-BB (25 µCi/ml) was purchased from Amersham Biosciences. 4-Acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid (AMS) was purchased from Molecular Probes. PDGF-BB, GSH, GSSG, NADPH, 3-(4,5-dimethyl-thiazole-2-yl)-2,5-diphenyl tetrazolium bromide (MTT), p-nitrophenyl phosphate, and mouse anti-FLAG monoclonal antibody were from Sigma. GSSG reductase was from Roche Applied Science. H₂O₂, CdCl₂, and dithiothreitol (DTT) were from Wako Pure Chemicals (Osaka, Japan).

Cell Line and Culture—H9c2 cells, a clonal line derived from embryonic rat heart, were obtained from American Type Culture Collection (CRL-1446). H9c2 cells which had been transfected with the expression vector for mouse GRX1 cDNA have been described previously (20). Cell lines (GRX22 and -30) expressing high levels of FLAG-tagged GRX protein were used in the study. H9c2 cells and gene-transfected cells were cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum (FCS) containing 75 µg/ml G418 and 75 µg/ml hygromycin B in a humidified atmosphere of 95% air and 5% CO₂ at 37 °C.

Cell Proliferation—The proliferation of cultured cells was evaluated by measuring attached live cells photometrically after staining with crystal violet. The cells (5000) were placed in 100 µl of medium/well in 96-well plates and cultured in medium containing 0.5% FCS with or without 0.5 nM PDGF-BB for specific periods. Then the cells were fixed with 4% paraformaldehyde in phosphate-buffered saline (pH 7.5), washed, and stained with 0.01% crystal violet at room temperature for 20 min. Each well was extensively washed with water and dried. The stained cells were lysed by adding 100 µl of lysis buffer A (10% SDS and 0.1 N HCl), and the cell number was then estimated photometrically by measuring the absorbance at 570 nm using a microplate reader.

Immunoblot Analysis—Cultured cells were harvested and lysed for 20 min at 4 °C in lysis buffer B (20 mM Tris (pH 7.2), 150 mM NaCl, and 1% Nonidet P-40, including protease inhibitors (20 µM phenylmethylsulfonyl fluoride, 50 µM pepstatin, and 50 µM leupeptin)). The protein concentration was determined using a BCA assay kit (Pierce). Protein samples were electrophoresed on SDS-polyacrylamide gels (7.5–15%) under reducing conditions as described previously (32). The proteins in the gels were transferred onto a nitrocellulose membrane. The membranes were blocked in Tris-buffered saline (TBS, 10 mM Tris-HCl (pH 7.5) and 150 mM NaCl) containing 0.1% (v/v) Tween 20 (TBST) and 5% (w/v) nonfat dry milk and then reacted with primary antibodies in TBST containing 5% (w/v) bovine serum albumin or 5% (w/v) nonfat dry milk overnight with constant agitation at 4 °C. After several washes with TBST, the membranes were incubated with peroxidase-conjugated secondary antibodies. Proteins in the membranes were then visualized using the enhanced chemiluminescence (ECL) detection kit (Amersham Biosciences) according to the manufacturer's instructions.

Immunoprecipitation—Cultured cells were harvested and lysed in lysis buffer B. Cell lysates normalized for protein levels were immunoprecipitated using anti-c-Myc monoclonal antibody-conjugated agarose beads for 2 h at 4 °C. The beads were washed three times with cold lysis buffer B. For the immunoblot analysis, the beads were resuspended in 100 µl of SDS sample buffer containing DTT and boiled for 3 min. Then the supernatant was directly subjected to SDS-PAGE as described above.

Reverse Transcription-PCR—Total RNA was prepared from cultured cells using the standard protocol and was reverse-transcribed using a One Step RNA PCR kit (avian myoblastosis virus (AMV)) (TaKaRa Biomedicals, Japan) with AMV-derived reverse transcriptase XL according to the manufacturer's instructions. PCR was run for 25 cycles of 95 °C for 0.5 min at 65 °C and for 0.5 min at 72 °C for 1.5 min. Primer sequences were as follows: for rat LMW-PTP (33) (GenBank^TM accession number NM_021262 [GenBank] ), forward, 5'-CAT GGC AGA GGT TGG GTC CAA GTC AGT GC-3', and reverse, 5'-GCT AGT GAG TCT TCT CCA GGA AGG CCT TG-3'; for rat GRX (GenBank^TM accession number NM_022278 [GenBank] ), forward, 5'-CGG CAT GGC TCA AGA GTT TGT GAA CTG CAA AAT CC-3', and reverse, 5'-GTG GTT ACT GCA GAG CTC CAA TCT GCT TTA GCC GC-3'; for rat -actin (GenBank^TM accession number BC063166 [GenBank] ), forward, 5'-GAG CTA TGA GCT GCC TGA CG-3', and reverse, 5'-AGC ATT TGC GGT GCA CGA TG-3'.

Construction of LMW-PTP Gene Expression Vector—Fulllength human LMW-PTP cDNA was cloned from total RNA of human colon cancer HCT8 cells by reverse transcription-PCR using Super Script II RNase H reverse transcriptase (Invitrogen) and Advantage HF2 Taq polymerase (BD Biosciences Clontech) with the following primer pair, which was designed on the basis of reported nucleotide sequences for human LMW-PTP (34) (GenBank^TM accession number M83653 [GenBank] ): forward, 5'-CGT GGA TCC GAG AAG ATG GCG-3', and reverse, 5'-CCC GAA TTC TCA GTG GGC CTT-3'. The amplified DNA fragments were subcloned into a TA cloning vector (pCRII, Invitrogen), and the nucleotide sequences were confirmed by sequencing with an ALFexpress II system (Amersham Biosciences). The LMW-PTP cDNA was cloned into a C-terminal Myc-tagged expression vector, pcDNA3.1/Myc-His (Invitrogen) under the control of the CMV promoter for expression in mammalian cells and also was cloned into a bacterial expression vector, pGEX6p-1 (Amersham Biosciences). The gene transfection into mammalian cells was performed using Lipofectamine Plus reagent (Invitrogen) according to the manufacturer's directions.

RNA Interference and Transfections—Double-stranded small interfering RNAs (siRNAs) corresponding to rat GRX DNA sequences (GenBank^TM accession number NM_022278 [GenBank] ) (5'-ACU GCA AGA UUC AGU CUG GdTdT-3' (siRNA-GRX-1) and 5'-AAC GUG GUC UCC UGG AAU UdTdT-3' (siRNA-GRX-2)) and to rat LMW-PTP DNA sequences (GenBank^TM accession number NM_021262 [GenBank] ) (5'-CAC AUU GCA CGG CAG AUU AdTdT-3' (siRNA-LMW-PTP-1) and 5'-UGA GAG AUC UGA AUA GAA AdTdT-3' (siRNA-LMW-PTP-2)) were synthesized and annealed by Samchully Pharmaceuticals, Seoul, Korea. siRNAs were transfected into the cells using Lipofectamine2000 (Invitrogen) according to the manufacturer's protocol with a final siRNA concentration of 100 nM.

LMW-PTP Activity Assay—The activity of LMW-PTP was measured according to the methods of Chiarugi et al. (35) with a slight modification. Briefly, the LMW-PTP expression vector was transfected into control and GRX gene-transfected H9c2 cells. After 24 h of transfection, the cells were lysed, and the cell lysates normalized for protein levels were subjected to immunoprecipitation using anti-c-Myc monoclonal antibody-conjugated agarose beads at 4 °C for 2 h. In each experimental condition, duplicate immunoprecipitates were prepared and subjected to the PTP activity assay or to immunoblot analysis for estimation of the level of LMW-PTP obtained in each immunoprecipitation. After washing, the beads were incubated at 37 °C for 30 min in 0.1 M sodium acetate (pH 5.5), 1 mM EDTA, and 4 mM p-nitrophenyl phosphate, then the reaction was stopped by the addition of 6 N NaOH. Aliquots of the mixture were used to determine the phosphatase activity spectrophotometrically by measuring the absorbance at 410 nm. Each activity was standardized by comparing with the level of LMW-PTP in the immunoprecipitates densitometrically estimated in immunoblot analysis and expressed as the relative value to the activity for the untreated control (% of untreated control). Furthermore, LMW-PTP activity in vitro was also measured for recombinant LMW-PTP after incubation with or without H₂O₂. The aliquots were diluted 20-fold by adding 0.1 M sodium acetate (pH 5.5), 1 mM EDTA and subjected to the PTP activity assay as described above. Activity was quantified by comparison with a standard curve (0.01–0.2 mM p-nitrophenol).

Purification of Recombinant LMW-PTP and Generation of Antibody against LMW-PTP—LMW-PTP was purified with the glutathione S-transferase (GST) gene fusion system (Amersham Biosciences) according to the manufacturer's instructions. In brief, E. coli strain BL21 cells were transformed with pGEX6p-LMW-PTP, and protein expression was induced by isopropyl -D-1-thiogalactopyranoside. GST-fused LMW-PTP (GST-LMW-PTP) was affinity-purified from cell lysates using glutathione-Sepharose 4B (Amersham Biosciences) and then digested with PreScission protease. The cleaved GST was removed from glutathione-Sepharose 4B, and LMW-PTP was purified. The LMW-PTP was used to immunize rabbits to generate anti-LMW-PTP antibodies and also used for in vitro experiments concerning redox regulation of LMW-PTP as described below.

Determination of the Redox State of LMW-PTP in Vitro—The redox state of LMW-PTP was assessed by modifying free thiol with AMS (20). Briefly, recombinant LMW-PTP (2 µg) was reduced by incubation with 50 mM DTT for 30 min on ice. DTT was then removed by gel filtration on NAP-10 columns (Amersham Biosciences). Reduced LMW-PTP was incubated at room temperature for the periods indicated with H₂O₂ in buffer with or without the GRX (4–40 µg) and/or GSH-regenerating system as described (20). The GSH-regenerating system was composed of GSH/GSSG (1 mM GSH and 0.05 mM GSSG), NADPH (1 mM), and GSSG reductase (GR) (1.2 units). After incubation, LMW-PTP protein was incubated with buffer containing a final concentration of 50 mM Tris-HCl (pH 7.4), 1% SDS, and 15 mM AMS at 4 °C for 1 h. The reaction was stopped by the addition of 5x SDS sample buffer. Proteins were separated by 15% SDS-PAGE with or without DTT and blotted to a nitrocellulose membrane. Proteins in the membrane were then visualized by immunoblot analysis as described above.

Peroxide Quantification—Peroxide was quantified using the PeroXOquant quantitative peroxide assay kit (Pierce) according to the manufacturer's instructions. In brief, cells were incubated with PDGF-BB for given periods. Cells were then harvested and lysed on ice in 100 µl of distilled water by sonication using an ultrasonic applicator (Bioruptor UCD-200T, COSMO BIO, Tokyo, Japan). Working reagent (100 µl) was mixed with 10 µl of lysate and incubated at room temperature for 15–20 min. The purple product composed of Fe³⁺-xylenol orange complex was detected spectrophotometrically at 570 nm in the reaction mixture. Activity was quantified by comparison with a standard curve (1–50 µM H₂O₂). The protein concentration was determined using a BCA assay kit (Pierce).

Statistical Analysis—Statistical analysis was performed using Student's t test or analysis of variance (StatView software). Significance was set at p < 0.05.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Overexpression of the GRX Gene Inhibits PDGF-induced Proliferation of H9c2 Cells—Rat myocardiac H9c2 cells were transfected with mouse GRX1 gene expression vectors to obtain two cell lines overexpressing GRX. Fig. 1A shows that expression of GRX increased in the overexpressors (GRX22 and GRX30) and was immunologically undetectable in mock-transfected (control) H9c2 cells. Recently it has been reported that an antioxidant inhibited cell growth by blocking the phosphorylation of tyrosines in PDGFR elicited by PDGF-BB (36). In H9c2 cells PDGF-BB induces cell proliferation through PDGFR (30). We also examined the PDGF-BB-dependent proliferation of H9c2 cells in culture medium containing 0.5% FCS with different concentrations of PDGF-BB (0–0.5 nM) and found that the cells proliferated with PDGF-BB in a dose-dependent manner (Fig. 1B, inset). The result showed that 0.5 nM PDGF-BB could sufficiently stimulate the proliferation of H9c2 cells and was consistent with a previous report by Brostrom et al. (30). To investigate the functional role of overexpressed GRX in myocardiac cell proliferation, control and GRX gene-transfected cells were cultured in medium supplemented with 0.5% FCS with or without 0.5 nM PDGF-BB, then cell proliferation was analyzed as described under "Materials and Methods." In Fig. 1B, after 3 days of culture with 0.5 nM PDGF-BB, the cell count had doubled in control cells, but the increase was suppressed among the transfectants. On the other hand, without PDGF-BB, proliferation was apparently suppressed in all cells. To investigate whether PDGF-BB signaling is influenced by overexpression of GRX, the phosphorylation status of PDGFR was examined immunologically in the cells treated with 0.5 nM PDGF-BB (Fig. 1C). In control cells phosphorylation of PDGFR was markedly increased by PDGF-BB, but the increase was apparently suppressed in GRX gene-transfected cells treated with PDGF-BB. Together, these results indicate that PDGF-dependent cell proliferation was suppressed by overexpression of GRX in H9c2 cells through the suppressed phosphorylation of a specific receptor, PDGFR. Using ¹²⁵I-labeled PDGF-BB, the binding of PDGF-BB both to control and GRX gene-transfected cells was investigated as described in the supplemental material. The results showed that overexpression of GRX did not influence the PDGF-BB binding to the cells (supplemental Fig. S1, A and B). Furthermore, we examined cell surface expression of PDGFR in control and GRX gene-transfected cells after biotinylation of cell surface proteins and found that overexpression of GRX did not change the expression level of cell surface PDGFR (supplemental Fig. S1C). Together, these results indicate that overexpression of GRX does not influence the ligand-receptor binding mechanism of PDGF-BB in H9c2 cells.

View larger version (34K): [in this window] [in a new window]   FIGURE 1. Overexpression of GRX suppresses the PDGF-dependent cell growth. A, expression of GRX in mock-transfected (control) and GRX gene-transfected (GRX22 and GRX30) H9c2 cells. The expression levels of proteins were estimated by immunoblot (IB) analysis using antibodies against GRX (rabbit polyclonal) and FLAG (mouse monoclonal) as described under "Materials and Methods." Mouse anti-glyceraldehyde-3-phosphate dehydrogenase monoclonal antibody was used for the loading controls. B, proliferation of control and gene-transfected H9c2 cells cultured for 3 days in medium containing 0.5% FCS with or without PDGF-BB (0.5 nM). Cell proliferation was evaluated by measuring attached live cells photometrically after staining with crystal violet as described under "Materials and Methods." Dose-dependent proliferation with PDGF-BB is shown in control cells cultured in the medium containing 0.5% FCS with or without PDGF-BB (0–0.5 nM)(inset). Each value represents the mean of four independent experiments, and the S.D. was always within 10% of the mean. A.U., absorbance units. C, time course of phosphorylation of PDGFR in control and gene-transfected H9c2 cells stimulated with PDGF-BB. Cells were treated with 0.5 nM PDGF-BB for the period indicated. Phosphorylation of PDGFR was estimated by immunoblot analysis using rabbit antibodies against PDGFR and phospho-(Tyr-751)-PDGFR. The data represent three independent experiments.

View larger version (32K): [in this window] [in a new window]   FIGURE 2. Transfection of siRNAs for LMW-PTP and GRX up-regulates the phosphorylation of PDGFR in H9c2 cells treated with PDGF-BB. To down-regulate the expression of LMW-PTP (A) and GRX (B), specific siRNAs (100 nM) for LMW-PTP and GRX were designed for each gene and transfected into H9c2 cells as described under "Materials and Methods." After 48 h transfection the cells were lysed, and PCR analysis was performed to evaluate the transcriptional expressions of LMW-PTP (A) and GRX (B). siRNAs bearing scramble sequences were also transfected into the cells as controls. The expression of -actin was also shown as a control. Each value represents the mean ± S.D. of three independent experiments. C, after 48 h transfection with siRNAs, cells were serum-starved for 24 h and then stimulated with 0.5 nM PDGF-BB for the periods indicated. The phosphorylation status of PDGFR was examined by immunoblot (IB) analysis using rabbit antibodies against PDGFR and phospho (Tyr-751)-PDGFR. The data represent three independent experiments.

To further examine the involvement of LMW-PTP in the mechanism regulating PFGF-BB signaling, we prepared the cells transiently transfected with the expression vector for LMW-PTP, because the endogenous level of LMW-PTP protein in H9c2 cells was too low to investigate its biochemical characteristics. To clarify whether LMW-PTP regulates the phosphorylation of PDGFR in H9c2 cells, the level of tyrosine phosphorylation of PDGFR was examined in an immunoblot analysis using H9c2 cells transfected with two amounts of LMW-PTP gene expression vector, pcDNA-LMW-PTP. As shown in Fig. 3A, the level of phosphotyrosine in PDGFR was decreased by transfection with pcDNA-LMW-PTP. To investigate whether transient expression of LMW-PTP gene influences the expression level of cell surface PDGFR, we immunoprecipitated PDGFR from cell lysates of the cells transfected with or without pcDNA-LMW-PTP after biotinylation of cell surface proteins, and immunoprecipitates were examined by immunoblot analysis to detect PDGFR or biotinylated proteins. The results showed that the expression of exogenous LMW-PTP does not change the expression level of cell surface PDGFR (data not shown). To confirm that the decrease in the tyrosine phosphorylation of PDGFR was due to an increase in LMW-PTP activity caused by the gene transfection, the effect of PTP inhibitor III (PTPI-III) on the phosphorylation status of PDGFR was examined in the cells transfected with pcDNA-LMW-PTP. PTPI-III is a cell-permeable broad-range inhibitor of PTPs (40). In Fig. 3B, upper, the cells transfected with pcDNA-LMW-PTP were pretreated with various concentrations of PTPI-III for 30 min. Expressed Myc-tagged LMW-PTP was immunoprecipitated, then the activity for LMW-PTP was assayed in the immunoprecipitates as described under "Materials and Methods." The LMW-PTP activity was decreased by PTPI-III in a dose-dependent manner. In Fig. 3B, lower, the status of PDGFR was also examined by immunoblot analysis in the LMW-PTP gene-transfected cells treated with PTPI-III. Although the phosphorylation of PDGFR was apparently suppressed by transfection with pcDNA-LMW-PTP, the level gradually increased on treatment with PTPI-III in a dose-dependent manner. These results indicate that overexpression of the LMW-PTP gene causes a decrease in the phosphorylation of PDGFR simply through increased LMW-PTP activity.

View larger version (29K): [in this window] [in a new window]   FIGURE 3. Overexpression of LMW-PTP decreases the phosphorylation of PDGFR through the phosphotyrosine phosphatase activity. A, H9c2 cells were transfected with various concentrations of LMW-PTP expression vector (pcDNA-LMW-PTP) as described under "Materials and Methods." After 24 h the level of phosphorylated PDGFR was examined by immunoblot (IB) analysis using rabbit antibodies against PDGFR and phospho-(Tyr-751)-PDGFR. The expression of exogenous Myc-tagged LMW-PTP was detected with rabbit anti-Myc polyclonal antibody. The data represent three independent experiments. B, after 24 h of transfection with pcDNA-LMW-PTP, H9c2 cells were treated for 30 min with various concentrations of PTPI-III, an inhibitor for PTP. Expressed Myc-tagged LMW-PTP was immunoprecipitated using anti-c-Myc monoclonal antibody-conjugated agarose beads, and then the activity for LMW-PTP was assayed in the immunoprecipitates as described under "Materials and Methods" (upper). Each value represents the mean ± S.D. of three independent experiments. The phosphorylation status of PDGFR was examined by immunoblot analysis as described above (lower). The data represent three independent experiments. C, control and GRX gene-transfected H9c2 cells were serum-starved for 24 h, treated with the indicated concentration of PTPI-III for 30 min, and then stimulated with or without 0.5 nM PDGF-BB. The phosphorylation status of PDGFR was examined by immunoblot analysis as described above. The data represent three independent experiments.

The Activity of LMW-PTP Is Up-regulated by Overexpression of GRX—The PTP activity of LMW-PTP is dependent on the level of cellular reduced glutathione (GSH) (35). To know the effect of overexpression of GRX on LMW-PTP, the activity of LMW-PTP was examined in control and GRX-overexpressing cells after transfection with the LMW-PTP gene expression vector. Control and GRX-overexpressing cells were transiently transfected with pcDNA-LMW-PTP. After 24 h of transfection, the cells were treated with PDGF-BB for given periods, and the activity of LMW-PTP was estimated as described under "Materials and Methods." As shown in Fig. 4A, the activity was always stronger in GRX-overexpressing cells than controls during the treatment with PDGF-BB, indicating that the expression level of GRX affected the activity of LMW-PTP. To investigate whether the up-regulation of LMW-PTP activity was due to the effect of overexpressed GRX, the effect of cadmium (CdCl₂), an inhibitor for GRX (41), on the LMW-PTP activity was examined in control and GRX-overexpressing cells transiently transfected with pcDNA-LMW-PTP. After 24 h of transfection, the cells were treated with or without PDGF-BB and/or CdCl₂. Then the activity of LMW-PTP was estimated as described under "Materials and Methods." Treatment with 200 µM CdCl₂ reduced the thiol transferase activity of GRX by more than 60% in GRX-overexpressing H9c2 cells (20). As shown in Fig. 4B, the LMW-PTP activity was significantly suppressed by CdCl₂ in control and GRX-overexpressing cells with or without PDGF-BB, indicating that GRX activity contributed to the activation of LMW-PTP. In Fig. 4C, the effect of CdCl₂ on the phosphorylation of PDGFR was examined in control and GRX-overexpressing cells treated with PDGF-BB. The results showed that the phosphorylation was apparently increased by CdCl₂ in control and GRX-overexpressing cells treated with PDGF-BB. The phosphorylation levels were similarly increased in control and GRX-overexpressing cells treated with PDGF-BB. Collectively, these results indicate that the activity for LMW-PTP is up-regulated by overexpression of GRX through the increased thiol transferase activity of GRX. This also suggests that GRX plays an important role in regulating the redox state of LMW-PTP under PDGF-BB treatment.

View larger version (27K): [in this window] [in a new window]   FIGURE 4. Cadmium diminished the activity of LMW-PTP during PDGF-BB treatment. A, expression vectors for Myc-tagged LMW-PTP were transiently transfected into control and GRX gene-transfected cells as described under "Materials and Methods." After 24 h cells were serum-starved for 24 h and then stimulated with 0.5 nM PDGF-BB for the periods indicated. Expressed Myc-tagged LMW-PTP was immunoprecipitated, then the activity for LMW-PTP was assayed in the immunoprecipitates as described under "Materials and Methods." *, p < 0.05 versus untreated control cells (time 0). **, p < 0.05 versus same time point for control cells. B, cells were transfected with the LMW-PTP expression vectors and serum-starved as in A. Then the cells were pretreated with 200 µM CdCl2 for 1 h and stimulated with 0.5 nM PDGF-BB for the periods indicated. Expressed Myc-tagged LMW-PTP was immunoprecipitated, and the activity for LMW-PTP was assayed as described above. *, p < 0.05 versus untreated control cells (time 0). ***, p < 0.05 versus untreated GRX30 cells (time 0). C, control and GRX gene-transfected cells were serum-starved for 24 h. Then the cells were pretreated with 200 µM CdCl2 for 1 h and stimulated with PDGF-BB for the periods indicated. The phosphorylation status of PDGFR was examined by immunoblot analysis (IB) using specific antibodies.

View larger version (28K): [in this window] [in a new window]   FIGURE 5. PDGF-BB-induced peroxide production is involved in the phosphorylation of PDGFR. A, control and GRX gene-transfected H9c2 cells were treated with 0.5 nM PDGF-BB for the periods indicated. At each time point cells were harvested, and the intracellular peroxide concentration was measured using a peroxide assay kit (Pierce) as described under "Materials and Methods." Each value represents the mean ± S.D. of three independent experiments. *, p < 0.05 versus untreated control cells (time 0). **, p < 0.05 versus untreated GRX30 cells (time 0). B, control cells were pretreated with or without 500 µM Trolox (30 min) or 1 mM NAC (30 min) and stimulated with 0.5 nM PDGF-BB for 5 min. The phosphorylation status of PDGFR was examined by immunoblot (IB) analysis using specific antibodies. C, control cells were transfected with the LMW-PTP expression vectors and serum-starved as in Fig. 3A. Then the cells were pretreated with or without 500 µM Trolox (30 min) or 1 mM NAC (30 min) and stimulated with 0.5 nM PDGF-BB for 5 min. Expressed Myc-tagged LMW-PTP was immunoprecipitated, then the activity for LMW-PTP was assayed in the immunoprecipitates as described under "Materials and Methods." Each value represents the mean ± S.D. of three independent experiments. *, p < 0.05 versus untreated control cells (–)(time 0). **, p < 0.05 versus control cells (–) treated with PDGF-BB for 5 min (time 5).

GRX Requires the GSH-regenerating System for the Redox-dependent Regulation of LMW-PTP in Vitro—To further clarify the molecular mechanism by which GRX regulates the redox state of LMW-PTP, we examined whether GRX could protect LMW-PTP from oxidation-induced structural change in the presence of H₂O₂ in vitro. GRX was purified using the GST fusion protein expression system as described previously (20). GST-fused GRX was affinity-purified from E. coli cell lysates using glutathione-Sepharose 4B and then digested with PreScission protease. The cleaved GST was removed from glutathione-Sepharose 4B, and GRX was purified for the experiments. The LMW-PTP was incubated with or without 200 µM H₂O₂ under various conditions. The treatment with H₂O₂ decreased the PTP activity of LMW-PTP with or without GRX (Fig. 7A, left). However, in the presence of the GSH-regenerating system (GSH/GSSG and NADPH/GSSG reductase), GRX significantly suppressed the oxidation-induced inactivation of LMW-PTP, although the regenerating system itself also slightly suppressed the inactivation (Fig. 7A, right). Furthermore, in the presence of the GSH-regenerating system, GRX protected LMW-PTP from oxidation-induced inactivation in a dose-dependent manner (Fig. 7B). Then we examined the effect of GRX and/or the GSH-regenerating system on the redox state of LMW-PTP after treatment with H₂O₂ in vitro. In Fig. 7C, H₂O₂ caused oxidation-induced oligomer formation in LMW-PTP. The change was not suppressed solely by GRX. However, in the presence of the GSH-regenerating system, GRX could reduce the oxidation-induced change of LMW-PTP in a dose-dependent manner. On the other hand, when LMW-PTP was initially oxidized by H₂O₂, it was completely inactivated and not reactivated even in the presence of DTT or GRX with a GSH-regenerating system (data not shown). Taken together, these results indicate that GRX regulates the redox state of LMW-PTP in concert with the GSH-regenerating system to protect the activity against oxidative stress.

View larger version (42K): [in this window] [in a new window]   FIGURE 6. Effect of peroxide on LMW-PTP in vitro. A, recombinant LMW-PTP (2 µg) was reduced by incubation with 50 mM DTT for 30 min on ice. DTT was then removed by gel filtration on NAP-10 columns (Amersham Biosciences). Reduced LMW-PTP was incubated with the indicated concentration of H2O2 for 10 min at room temperature. The samples were modified with AMS as described under "Materials and Methods" and subjected to non-reducing and reducing SDS-PAGE followed by immunoblot analysis with anti-LMW-PTP antibody. B, reduced LMW-PTP was treated with the indicated concentration of H2O2 as in A. The activity for LMW-PTP in vitro was assayed as described under "Materials and Methods." Each value represents the mean ± S.D. of three independent experiments.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

View larger version (28K): [in this window] [in a new window]   FIGURE 7. GRX regulates the redox state of LMW-PTP in vitro under the conditions with peroxide. A, reduced LMW-PTP was prepared as in Fig. 6A and incubated at room temperature for the periods indicated with 200 µM H2O2 in buffer with or without the GRX (0–40 µg) and/or GSH-regenerating system as described under "Materials and Methods." The GSH regenerating system was composed of GSH/GSSG (1 mM GSH and 0.05 mM GSSG), NADPH (1 mM), and GSSG reductase (GR) (1.2 units). The activity for LMW-PTP in vitro was assayed as described under "Materials and Methods." Each value represents the mean ± S.D. of three independent experiments. *, p < 0.05 versus same time point for GSH system. B, reduced LMW-PTP was incubated for 10 or 30 min with or without 200 µM H2O2 in buffer containing the GSH-regenerating system and various concentrations of GRX. The activity for LMW-PTP in vitro was assayed as described under "Materials and Methods." Each value represents the mean ± S.D. of three independent experiments. *, p < 0.05 versus treatment with H2O2 for 10 min without GRX. **, p < 0.05 versus treatment with H2O2 for 30 min without GRX. C, reduced LMW-PTP was incubated for 10 min with 200 µM H2O2 in buffer with or without the GRX and/or GSH-regenerating system as in A. The samples were modified with AMS and separated by reducing and non-reducing SDS-PAGE as described under "Materials and Methods." Oxidation-induced conformational changes of LMW-PTP were examined by immunoblot analysis using anti-LMW-PTP antibody.

The PDGF pathway is a well characterized growth factor signaling pathway, and its importance is also recognized in the development of certain cell types. Gene knock-out experiments in mice emphasized the importance of PDGF-BB/PDGFR signaling in myocardiac development. Furthermore, PDGFR is expressed in adult rat heart (29) and rat myocardiac H9c2 cells (30), and PDGF-BB/PDGFR signaling also controls the proliferation of neonatal rat cardiomyocytes (31). These findings suggest that the PDGF-BB/PDGFR signaling pathway plays an important role in the cellular physiology of myocardiac cells, although the precise regulatory mechanism for the pathway has yet to be clarified.

PDGFR is a receptor-type tyrosine kinase, the activation of which is regulated by PDGF-dependent autophosphorylation and dephosphorylation by PTPs. On stimulation with PDGF, a number of tyrosine residues are phosphorylated in the cytosolic domain of PDGFR, leading to a site-specific recruitment of signal transduction molecules such as phosphatidylinositol 3-kinase, phospholipase C, Src, Grb2, SHP-2, GTPase-activating protein, and so forth (25). PTPs that were previously implicated in the control of PDGFR phosphorylation include SHP-1 (50), SHP-2 (51, 52), PTP-1B (19, 53), PTP-PEST (54), DEP-1 (55), TC-PTP (38), and LMW-PTP (56, 57).

Among PTPs that are involved in PDGF signaling we focused on LMW-PTP because the redox-dependent regulation of the molecule and its role in PDGF-BB/PDGFR signaling have been studied extensively (39). LMW-PTP was expressed in H9c2 cells, and the expression level was not affected by transfecting the GRX gene expression vector (supplemental Fig. S2). When the expression of LMW-PTP was suppressed by siRNA transfection, PDGF-BB-induced tyrosine phosphorylation was apparently enhanced compared with untransfected controls (Fig. 2C). This strongly suggests that LMW-PTP is responsible for the dephosphorylation of PDGFR in H9c2 cells, although other PTPs may also be involved in the mechanism. On the other hand, the PDGF-induced phosphorylation of PDGFR was apparently enhanced in GRX-overexpressing cells treated with tyrosine phosphatase inhibitor PTPI-III but was not enhanced in control cells treated with PTPI-III (Fig. 3C). In GRX-overexpressing cells, the enhancing effect of PTPI-III on the PDGF-induced phosphorylation of PDGFR was consistent with the data using siRNA for LMW-PTP. However, at present it is not clear why suppression of endogenous LMW-PTP by PTPI-III does not much influence on the PDGF-BB-induced phosphorylation of PDGFR in control cells. It is reported that -bromoacetophenone derivatives such as PTPI-III act as PTP inhibitors by covalently alkylating the conserved catalytic Cys residue in the PTP active site (58). In this respect it is possible that the susceptibility of endogenous LMW-PTP to PTPI-III could be changed when the redox state of endogenous LMW-PTP is different between control and GRX-overexpressing cells. This suggests that the effect of PTPI-III on endogenous LMW-PTP may be highly enhanced in GRX-overexpressing cells compared with controls.

LMW-PTP is an 18-kDa enzyme that is widely expressed (39, 59). Previous studies on the molecular biology of LMW-PTP in NIH3T3 cells demonstrated a well defined role for this enzyme in PDGF-BB-induced mitogenesis showing that activated PDGFR is a substrate for LMW-PTP (56). We showed that cellular LMW-PTP was inactivated by peroxide, which could be generated with PDGF-BB, and the inactivation was suppressed by antioxidants such as Trolox or NAC (Figs. 5, B and C). These results support previous findings that LMW-PTP is oxidized by ROS generated in cells stimulated with PDGF-BB, resulting in up-regulation of the phosphorylation of PDGFR. We also showed that GRX could up-regulate the activity of LMW-PTP even in the cells treated with PDGF-BB, resulting in an enhanced dephosphorylation of PDGFR. Furthermore, in vitro we found that treatment with H₂O₂ causes the formation of high molecular weight oligomers of LMW-PTP in SDS-PAGE under non-reducing conditions. The high molecular weight form disappeared and shifted to a single molecule of 25 kDa in SDS-PAGE under reducing conditions. This suggested that LMW-PTP forms DTT-reducible oligomers in vitro through oxidation-induced disulfide-bond formation between the molecules. In the presence of GRX- and GSH-regenerating systems, the H₂O₂-induced oligomer formation of LMW-PTP was apparently suppressed, and the activity of LMW-PTP was also protected from H₂O₂-induced inactivation. Collectively, these results indicate that GSH/GRX systems effectively work to protect against oxidation-dependent structural change and inactivation of LMW-PTP.

During PDGF signaling, LMW-PTP is regulated by a redox-dependent mechanism involving the two Cys residues of the catalytic site, which change reversibly from the reduced to oxidized state after PDGF treatment. The reversibility of in vivo LMW-PTP oxidation is glutathione-dependent (35). The additional catalytic Cys-17 retains an interesting role in the formation of the S-S intramolecular bond, which protects the catalytic Cys-12 from further and irreversible oxidation. The presence of an additional Cys near the catalytic one confers upon LMW-PTP the ability to rapidly recover its activity and finely regulate PDGF receptor activation. To investigate whether specific Cys residues of the catalytic site of LMW-PTP are involved in the oxidation-induced oligomer formation of LMW-PTP in vitro, the LMW-PTP mutant, in which Cys-17 was changed to Ser by site-directed mutagenesis, was produced and characterized by non-reducing SDS-PAGE after treatment with H₂O₂. The results showed that high molecular weight oligomers were formed by H₂O₂ even with the mutant LMW-PTP, indicating that Cys-17 had no particular contribution to oxidation-induced oligomer formation (data not shown). This suggests that there is no direct correlation between the inactivation status and formation of high molecular weight oligomers of LMW-PTP under conditions with H₂O₂ in vitro, although oxidation-induced oligomer formation was effectively suppressed by GRX- and GSH-regeneration systems. It is also noteworthy that the oxidation-induced oligomer formation of LMW-PTP was not detected in non-reducing SDS-PAGE of cell lysate samples from LMW-PTP gene-transfected cells treated with H₂O₂ (data not shown). Thus, further investigation is required to clarify precise mechanical correlation between the activation status and structure of LMW-PTP in cells treated with PDGF.

PTP-1B has been extensively studied in terms of its redox-dependent regulation related to the signaling receptors for epidermal growth factor (44, 60) and insulin (61, 62). The oxidation state of the active-Cys in PTP1B was also clearly characterized by structural analyses (63, 64). PTP1B has been reported to be involved in the redox-dependent dephosphorylation of PDGFR. Lee et al. (19) reported that oxidized PTP-1B is reduced by TRX rather than GSH/GRX systems. On the other hand, Barrett et al. (65) reported that GSH could recycle oxidized PTP1B, showing the involvement of GSH-related systems in the mechanism. Thus, the precise mechanism of the redox regulation of PTP-1B is still controversial. SHP-2 is also an example of a redox-regulated PTP involved in the PDGFR signaling (52). Recently, the tumor suppressor protein, PTEN (phosphate and tensin homolog deleted on chromosome ten), has also been reported to dephosphorylate PDGFR (66). Although the activity of PTEN is regulated in a redox-dependent manner, oxidized PTEN is reduced by TRX rather than GRX in vitro (67). In the present study the expression of TRX was slightly suppressed in the GRX-overexpressing cells compared with controls (data not shown), but its relation to the redox-dependent regulation of LMW-PTP was not elucidated. On the other hand, in the case of mouse embryonic fibroblasts in which the PTP-1B gene was knocked out, the phosphorylation status of PDGFR was not much influenced by stimulation with PDGF-BB, suggesting that other PTPs could compensate for the loss of PTP-1B function (68). Taken together, these findings suggest that several different PTPs are involved in the redox-dependent regulation of the dephosphorylation of phosphorylated PDGFR. Thus, to clarify the precise mechanism behind the regulation of PDGF signaling, further investigations are required. In conclusion, the present study shows that GRX and GSH-regenerating systems are involved in the regulation of PDGF-BB signaling through the redox-dependent regulation of LMW-PTP.

	FOOTNOTES

 
^* This work was supported in part by grants-in-aid for the 21st Century Center of Excellence (COE) program from the Ministry of Education, Science, Sports, Culture, and Technology of Japan and by grants from the Ministry of Health, Labor, and Welfare, Japan. 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.

The on-line version of this article (available at http://www.jbc.org) contains supplemental data.

¹ Both authors contributed equally to this work.

² To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology in Disease, Atomic Bomb Disease Institute, Nagasaki University Graduate School of Biomedical Sciences, 1-12-4 Sakamoto, Nagasaki 852-8523, Japan. Tel.: 81-95-849-7099; Fax: 81-95-849-7100; E-mail: y-ihara{at}net.nagasaki-u.ac.jp.

³ The abbreviations used are: TRX, thioredoxin; AMS, 4-acetamido-4'-maleimidylstilbene-2,2'-disulfonic acid; DTT, dithiothreitol; GRX, glutaredoxin; GST, glutathione S-transferase; LMW-PTP, low molecular weight-protein-tyrosine phosphatase; PTPI, PTP inhibitor; NAC, N-acetyl-L-cysteine; PDGF, platelet-derived growth factor; PDGFR, PDGF receptor; ROS, reactive oxygen species; FCS, fetal calf serum; siRNA, small interfering RNA; SHP, Src homology 2-containing protein-tyrosine phosphatase; DEP-1, density-enhanced protein-tyrosine phosphatase-1; TC-PTP, T-cell protein-tyrosine phosphatase.

	ACKNOWLEDGMENTS

 
We are grateful to Midori Ikezaki and Akiko Emura for technical assistance.

	REFERENCES

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1. Holmgren, A. (1989) J. Biol. Chem. 264, 13963–13966[Free Full Text]
2. Holmgren, A. (1976) Proc. Natl. Acad. Sci. U. S. A. 73, 2275–2279[Abstract/Free Full Text]
3. Gravina, S. A., and Mieyal, J. J. (1993) Biochemistry 32, 3368–3376[CrossRef][Medline] [Order article via Infotrieve]
4. Shelton, M. D., Chock, P. B., and Mieyal, J. J. (2005) Antioxid. Redox Signal. 7, 348–366[CrossRef][Medline] [Order article via Infotrieve]
5. Holmgren, A. (1979) J. Biol. Chem. 254, 3672–3678[Free Full Text]
6. Gan, Z.-R., and Wells, W. W. (1986) J. Biol. Chem. 261, 996–1001[Abstract/Free Full Text]
7. Prinz, W. A., Aslund, F., Holmgren, A., and Beckwith, J. (1997) J. Biol. Chem. 272, 15661–15667[Abstract/Free Full Text]
8. Casagrande, S., Bonetto, V., Fratelli, M., Gianazza, E., Eberini, I., Massignan, T., Salmona, M., Chang, G., Holmgren, A., and Ghezzi, P. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 9745–9749[Abstract/Free Full Text]
9. Davis, D. A., Newcomb, F. M., Starke, D. W., Ott, D. E., Mieyal, J. J., and Yarchoan, R. (1997) J. Biol. Chem. 272, 25935–25940[Abstract/Free Full Text]
10. Lind, C., Gerdes, R., Schuppe-Koistinen, I., and Cotgreave, I. A. (1998) Biochem. Biophys. Res. Commun. 247, 481–486[CrossRef][Medline] [Order article via Infotrieve]
11. Bandyopadhyay, S., Starke, D. W., Mieyal, J. J., and Gronostajski, R. M. (1998) J. Biol. Chem. 273, 392–397[Abstract/Free Full Text]
12. Song, J. J., Rhee, J. G., Suntharalingam, M., Walsh, S. A., Spitz, D. R., and Lee, Y. J. (2002) J. Biol. Chem. 277, 46566–46575[Abstract/Free Full Text]
13. Kikawa, K. D., Vidale, D. R., Van Etten, R. L., and Kinch, M. S. (2002) J. Biol. Chem. 277, 39274–39279[Abstract/Free Full Text]
14. Wang, J., Tekle, E., Oubrahim, H., Mieyal, J. J., Stadtman, E. R., and Chock, P. B. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 5103–5106[Abstract/Free Full Text]
15. Adachi, T., Pimentel, D. R., Heibeck, T., Hou, X., Lee, Y. J., Jiang, B., Ido, Y., and Cohen, R. A. (2004) J. Biol. Chem. 279, 29857–29862[Abstract/Free Full Text]
16. Landino, L. M., Moynihan, K. L., Todd, J. V., and Kennett, K. L. (2004) Biochem. Biophy. Res. Commun. 314, 555–560[CrossRef][Medline] [Order article via Infotrieve]
17. Landino, L. M., Robinson, S. H., Skreslet, T. E., and Cabral, D. M. (2004) Biochem. Biophy. Res. Commun. 323, 112–117[CrossRef][Medline] [Order article via Infotrieve]
18. Caplan, J. F., Filipenko, N. R., Fitzpatrick, S. L., and Waisman, D. M. (2004) J. Biol. Chem. 279, 7740–7750[Abstract/Free Full Text]
19. Lee, S.-R., Kwon, K.-S., Kim, S.-R., and Rhee, S. G. (1998) J. Biol. Chem. 273, 15366–15372[Abstract/Free Full Text]