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1Free Radical and Radiation Biology Program, Department of Radiation Oncology, 2Department of Internal Medicine, The University of Iowa, Iowa City, Iowa 52242
Submitted 12 December 2002 ; accepted in final form 28 April 2003
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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50%, inhibited PDGF-BB
(composed of the homodimer of B-polypeptide chain)-induced
[3H]thymidine incorporation compared with control
oligodeoxynucleotides. Ref-1 AODN inhibited PDGF-BB-induced S phase
entry by 63%, which was overcome by overexpression of Ref-1 by
adenoviral-mediated gene transfer. Overexpression of Ref-1 alone without PDGF
enhanced SMC entry into the S phase. Furthermore, decreasing Ref-1 protein by
treatment of SMCs with Ref-1 AODN, or by immunodepletion of Ref-1 from nuclear
extracts, inhibited PDGF-BB-induced activator protein-1 (AP-1) DNA binding
activity. Chemical reduction restored the AP-1 DNA binding in Ref-1-depleted
nuclear extracts. These results suggest that Ref-1 contributes to the
regulation of PDGF-BB-stimulated cell cycle progression from
G0/G1 to S in SMCs, with one of the possible steps being
redox-regulation of AP-1 by Ref-1 protein.
activator protein-1; cell cycle; antisense
Ref-1 [also designated as human apurinic/apyrimidinic endonuclease (APE,
HAP1, and APEX)], a multifunctional protein, has a DNA repair domain in its
COOH terminus, which is highly homologous to the sequence of Escherichia
coli exonuclease III
(36), and a redox regulation
domain in its NH2 terminus, which is involved in the regulation of
the DNA binding activity of a group of nuclear factors including activator
protein-1 (AP-1), NF-
B, p53, hypoxia-inducible factor (HIF)-1
,
HIF-like factor, and others via redox modification of a cysteine residue in
the target protein (12,
19,
23,
26,
46,
47). The redox activity of
Ref-1 is regulated by chemical reduction and oxidation in vitro
(8,
46,
47) and may involve Cys-65 and
Cys-93 in the NH2-terminal region of the protein
(43). In cultured cells, Ref-1
serves as an intermediary between thioredoxin and AP-1 in response to oxidant
stress induced by PMA (19) or
ionizing radiation (44) and
mediates AP-1 activation by heat shock
(8). However, whether Ref-1 is
involved in redox regulation of cell proliferation either in SMCs or in other
types of cells is currently unknown.
Previous studies (27,
34,
37,
39) from our lab and others
have shown that PDGF can activate NAD(P)H oxidase in SMCs to produce
. This growth factor-induced
NAD(P)H oxidase-dependent reactive oxygen species (ROS) generation leads to
activation of transcription factors such as AP-1 and, consequently, cell
proliferation (14,
22,
34,
39). Although Ref-1 has been
reported to be elevated in some cancers such as prostate, ovarian, and
cervical cancer (25,
30,
48), and to be involved in
inhibition of apoptosis (6), a
potential role of Ref-1 in SMC proliferation has not been examined.
Accordingly, this study was undertaken to test the role of Ref-1 in SMC
mitogenesis and cell cycle progression. PDGF-BB (composed of the homodimer of
B-polypeptide chain), which binds to both the - and 
-subunits of
the PDGF receptor and therefore has the most potent mitogenic activity among
the three isoforms (17), was
used to stimulate SMC growth.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Transfection with oligodeoxynucleotides. Three ref-1 antisense oligodeoxynucleotide (AODN) sequences that potentially hybridize to the rat ref-1 mRNA (NCBI nuclear sequence database accession no. D44495 [GenBank] ) were designed around the start codon: 1) 5'-CATCGCTGTAACGAAAGCC-3'; 2)5'-CCGCTTCGGCATCGCTGTAA-3'; and 3)5'-TCTTCCGCTGCCGCTCTCTT-3'. Western blots showed the first two AODNs had no or weak effects on decreasing the protein levels of Ref-1. The third sequence had the strongest inhibitory effects on Ref-1 protein levels. The control oligodeoxynucleotides (ODN) sequences for the third AODN sequence included sense Ref-1: 5'-AAGAGAGCGGCAGCGGAAGA-3'; mismatch sequence: 5'-TCTTCAGCTGCCACTCTCTC-3'; and scrambled sequence: 5'-CGCTTTCCTTCCTTCCGCTC-3'. To prevent digestion of ODN by nucleases, all the ODN were phosphorothioated at the first five and the last five bases. To avoid forming tetraplexes via Hoogsteen base pair formation, which may lead to sequence-nonspecific effects, none of the ODN had G quartets (38). All ref-1 AODN and control ODN were synthesized and HPLC purified by Integrated DNA Technologies (Coralville, IA).
Transfection was carried out by using an Effectene transfection reagent kit (Qiagen, Valencia, CA) following the manufacturer's supplied protocol. In brief, 1.35 or 2.7 µl of 100 µM of AODN or control ODN were diluted with DNA-condensation buffer (buffer EC)to90 µl, followed by mixing with 4.8 µl of Enhancer. After incubation at room temperature for 5 min, 15 µl of Effectene transfection reagent were added to the DNA-Enhancer mixture and incubated at room temperature for 10 min. The mixture was then diluted with serum-free DMEM and applied to cells. SMCs (50–60% confluent) were treated with 0.012 or 0.024 µM of AODN or control ODN in 11 ml of serum-free DMEM per 100-mm dish, or with 0.012 or 0.024 µM of AODN or ODN in 1 ml of serum-free DMEM per well of a 24-well plate, for 24 h. The incubation media were replaced by serum-free DMEM with or without 10 ng/ml recombinant human PDGF-BB (Intergen, Purchase, NY) as indicated in each experimental protocol.
[3H]thymidine incorporation. Cellular DNA synthesis was examined as described previously (5). In brief, cells were grown to 50% confluence in 24-well plates, treated with 0.012 µM AODN or control ODN in serum-free DMEM for 24 h, and then incubated with serum-free DMEM with or without PDGF-BB (10 ng/ml) for an additional 9 h. One µCi/ml [3H]thymidine (Amersham Pharmacia Biotech, Piscataway, NJ) was added, and the incubation was continued for 2 h. After the medium was removed, the attached cells were washed with cold PBS, incubated in 20% trichloroacetic acid for 30 min, washed, and then incubated in 0.25 M NaOH for 12 h. The cells were then lysed by vortexing and analyzed for radioactivity by liquid scintillation counting. The number of attached cells in each well was counted by using a Coulter particle counter (Beckman Coulter) in a parallel culture. The incorporated radioactivity [counts per minute (cpm)] per cell was calculated, and results were expressed as the ratio of PDGF-BB treated sample to control. Experiments were performed three times in triplicate 24-well plates.
Infection with adenoviral ref-1 constructs. Replication-deficient
adenoviral constructs containing human ref-1 cDNA (Adref-1)
(10) were generated by the
University of Iowa Gene Transfer Vector Core. The particle titer of
Adref-1 stock was 1.2 x 1012 DNA particles/ml, and
the functional titer was 4 x 1010 plaque-forming units
(pfu)/ml. The titer of adenoviral constructs containing -galactosidase
(AdlacZ, control vector) was 1.2 x 1010 pfu/ml.
Subconfluent SMCs in 100-mm dishes were incubated with Adref-1 or
AdlacZ at indicated MOI in 6 ml of serum-free DMEM for 24 h. Cells
were then incubated with serum-free DMEM for 12 h, followed by PDGF-BB
treatment if used.
Flow cytometric analysis of bromodeoxyuridine-labeled cells. Progression of the cell cycle was monitored by flow cytometric analysis of bromodeoxyuridine (BrdU) incorporation vs. DNA content following a previously published procedure (13). Briefly, after 24 h of treatments with AODN or control ODN in serum-free medium, SMCs grown in 100-mm dishes were treated with or without PDGF-BB (10 ng/ml) for 6, 9, or 12 h. Cells were pulse-labeled with 10 µM of BrdU (Sigma, St. Louis, MO) for 30 min. The attached cells were collected by trypsinization and fixed with cold 70% ethanol immediately after the labeling. Fixed cells were treated with HCl/pepsin, and indirect immunostaining was performed by using anti-BrdU monoclonal antibody (Becton Dickinson Immunocytometry Systems, San Jose, CA). Nuclei were then treated with FITC-conjugated anti-BrdU mouse monoclonal antibody (Becton Dickinson Immunocytometry Systems) and counterstained with propidium iodide (Sigma, St. Louis, MO). Fluorescence-activated cell sorting (FACS) analysis was performed by using a FACScan (Becton Dickinson). Twenty thousand cells were measured for each sample and data were analyzed by using CellQuest software. For BrdU pulse-chase experiments (18), cells were infected with 250 multiplicity of infection (MOI) of Adref-1 or AdlacZ in serum-free medium for 24 h. Infected cells were then cultured in DMEM containing 10% FCS for 24 h. Cells were pulse labeled with BrdU for 30 min, chased in BrdU-free DMEM containing 10% FCS for an additional 12 h and assayed for cell cycle phase distribution by flow cytometry.
EMSA for DNA protein binding. For preparation of nuclear extracts, cells were resuspended in 0.05 M potassium phosphate buffer containing 1% NP-40 (Armesco, Solon, Ohio), 1 mM PMSF (Armesco), 1.5 mM pepstatin (Roche Molecular Biochemicals), and 8.4 mM leupeptin (Roche Molecular Biochemicals) and incubated on ice for 30 min. The cells were then lysed by passing through a 25.5-gauge needle. The nuclei were pelleted by centrifugation at 4°C two times for 1 min at 2,000 g, and the supernatants were removed after each spin. The pelleted nuclei were resuspended in ice-cold buffer C (in mM: 20 HEPES, 0.42 NaCl, 1.5 MgCl2, 0.2 EDTA, 1 PMSF, 1.5 pepstatin, and 8.4 leupeptin and 25% glycerol), and placed on ice for 40 min. The suspensions were centrifuged at 16,400 g for 6 min, and the supernatants were collected and diluted with ice-cold buffer D (in mM: 20 HEPES, 0.1 KCl, 0.2 EDTA, 1 PMSF, 1.5 pepstatin, and 8.4 leupeptin and 20% glycerol). The nuclear protein concentration was determined before EMSA was performed.
For EMSA (20), 1–10
µg of nuclear proteins were incubated in a total volume of 20 µl
consisting of 1 µg of poly-(dIdC), 1x gel shift buffer, and 5 µl
of 32P-labeled double-stranded ODN (100,000 cpm) containing a
consensus AP-1 binding sequence (underlined)
(5'-AGCTTGTGAGTCAGCCGGATC-3') on ice for 45 min. The
5' overhangs of double-stranded ODN (100 ng) containing the AP-1
cis-element were labeled by a Klenow fill-in reaction in the presence of 20
µCi [32P]dCTP (New England Nuclear Life Science Products,
Boston, MA). For DTT recovery experiments, nuclear extracts were incubated
with 1x gel shift buffer and 0.5 mM DTT on ice for 45 min. Then, 1 µg
of poly(dIdC) and 5 µl of 32P-labeled double-stranded ODN
(100,000 cpm) containing a consensus AP-1 binding sequence were added to
the above mixture and incubated for another 45 min. For gel supershifts, 4
µg of nuclear protein derived from PDGF-BB-treated cells were incubated
with 6 µg of anti-Jun-B antibody at room temperature for 25 min and then 1
µg of poly(dIdC), 1x gel shift buffer, and 5 µl of
32P-labeled double-stranded ODN containing a consensus AP-1 binding
sequence were added to above solution and incubated for another 30 min at room
temperature. Samples were electrophoresed on 5% nondenaturing polyacrylamide
gel and exposed to Biomax MR film at –80°C overnight.
For experiments designed to test the effects of immunodepletion of Ref-1 from nuclear extracts, subconfluent SMCs were growth arrested in serum-free DMEM for 48 h and then treated with 10 ng/ml of PDGF-BB for 3 h. After nuclear extracts were collected, 50 µl (45–50 µg) of control or PDGF-treated nuclear extracts were incubated with 0.32 µg of nonimmune rabbit IgG (Santa Cruz Biotechnology, Santa Cruz, CA) or anti-Ref-1 antibody (Santa Cruz Biotechnology), and 35 µl of protein A-agarose (Santa Cruz Biotechnology), by shaking at 4°C for 2 h. After centrifugation, the supernatants were collected, and the protein concentration was determined. Equal amounts of protein from each sample were subjected to EMSA.
Western blot analysis. Cells were harvested by scraping. Cell pellets were resuspended in 0.05 M potassium phosphate buffer and lysed by sonicating on ice with four bursts of 30 s each by using a Vibra Cell Sonicator (Sonics and Materials). Equal amounts of protein from the cell lysates were denatured with protein solubilization buffer at 100°C for 5 min and electrophoresed in a 12.5% SDS-polyacrylamide running gel and a 5% stacking gel. The protein was then electrotransferred onto a nitrocellulose membrane. After being blocked in 5% nonfat milk at room temperature for 1 h, the blot was washed and incubated with antibody against Ref-1 (1:250) and Jun-B (1:200) (Santa Cruz Biotechnology) at 4°C overnight. The blot was then washed and incubated with horseradish peroxidase-conjugated antibody against IgG (Boehringer-Mannheim, Indianapolis, IN) at room temperature for 1 h. The washed blot was then treated with enhanced chemiluminescence solution and exposed to Biomax MR film. Quantifications of optical densities of the bands were performed by using AlphaImager 2200 v5.5 (Alpha Innotech). The protein loading of all gels was verified by Coomassie blue staining.
Statistical analyses. Data are presented as means ± SD from three independent experiments. Differences among mean values of multiple groups were analyzed by using SigmaStat 2.0 for Windows. ANOVA-Tukey was used to determine the significance of differences at P < 0.05.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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To investigate the role of Ref-1 in PDGF-BB-induced SMC growth, we used
ref-1 AODN to block Ref-1 protein synthesis. In
Fig. 1, cells were treated with
0.012 µM of ref-1 AODN, control (scrambled, mismatch, or sense)
ODN, or Effectene (vehicle) alone for 24 h. SMCs were then incubated with 10
ng/ml of PDGF-BB for 1 h (Fig.
1A) or 48 h (Fig.
1B). Ref-1 protein levels in SMCs were significantly
decreased (by 50%, lane 2 vs. lane 4) by ref-1
AODN, and the effect persisted for at least 48 h
(Fig. 1B); control
ODNs showed no inhibitory effect.
To test whether ref-1 AODN blocks PDGF-BB-induced mitogenesis, [3H]thymidine incorporation (an index of DNA synthesis) was examined. The experiments demonstrated that substantial [3H]thymidine incorporation began to occur 9 h after treatment of growth-arrested SMCs with 10 ng/ml of PDGF-BB, and maximal [3H]thymidine uptake was observed at 13 h after PDGF-BB treatment (data not shown). Therefore, SMCs were treated with 0.012 µM ref-1 AODN or control ODN in serum-free medium for 24 h, followed by PDGF-BB incubation for 9 h, and then coincubated with [3H]thymidine for 2 h. As shown in Fig. 2, exposure to PDGF-BB produced a 2.5-fold increase in [3H]thymidine uptake compared with control; this increased uptake was inhibited by pretreatment with ref-1 AODN. Neither control ODN nor vehicle had a significant inhibitory effect. Visual evidence of cytotoxicity (cells rounded up or detached from plates) was not detected in these experiments and the cell numbers among these seven groups were not significantly different (data not shown). Thus inhibition of PDGF-induced [3H]thymidine uptake by ref-1 AODN was likely due to the decrease in Ref-1 levels.
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Because [3H]thymidine uptake reflects cell proliferation and/or
DNA repair activity, we then tested whether ref-1 AODN inhibited cell
cycle progression from G0/G1 to S. SMCs were
synchronized by serum starvation in the presence of ref-1 AODN or
control ODN for 24 h, followed by stimulation with 10 ng/ml of PDGF-BB for 6,
9, or 12 h (Fig. 3). Cells
stimulated with PDGF-BB alone were used for comparison. Cells were then pulse
labeled with BrdU and harvested for FACS analysis immediately at the end of
the labeling period. Representative results presented in
Fig. 3A showed that
76% of the cells were in G0/G1 and only 5% cells were in
S in serum-starved control cells. Twelve hours after PDGF-BB-stimulation, 57%
of the cells incorporated BrdU, suggesting that majority of the cells entered
S during this time interval. However, PDGF-stimulated entry into S was
significantly inhibited (50%) in cells pretreated with 0.012 µM
ref-1 AODN. In these experiments, neither the transfection vehicle
nor the control ODN (sense, scrambled, or mismatch) had a significant
inhibitory effect on progression from G0/G1 to S.
Quantification of three independent experiments
(Fig. 3B) shows a
statistically significant inhibitory effect of ref-1 AODN on
PDGF-BB-induced S entry compared with PDGF-BB alone or treatment with sense
ODN. A time course experiment showed that S entry was delayed by
ref-1 AODN (Fig.
3C). We observed no sub-G1 fraction in these
experiments (data not shown), indicating that very few of the collected cells
were dead. Thus under the defined experimental conditions, ref-1 AODN
inhibited PDGF-BB-induced SMC proliferation without significantly decreasing
cell viability.
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We then utilized Adref-1 to determine the effects of
overexpression of Ref-1 protein on progression from
G0/G1 to S in SMCs. Infection with Adref-1
resulted in a dose-dependent increases in Ref-1 protein in SMCs
(Fig. 4A); total Ref-1
protein (endogenous and exogenous Ref-1) increased 2.2-fold after infection
with 200 MOI of Adref-1 compared with noninfected cells.
AdlacZ (a vector control) did not increase the level of Ref-1
protein. To examine the effect of overexpression of Ref-1 alone, cells were
infected with 250 MOI of Adref-1 or AdlacZ in serum-free
medium for 24 h (control cells were incubated in serum-free medium alone for
24 h), and pulse-chase experiments were performed
(Fig. 4, B and
C). Analysis of cell cycle phase distribution by flow
cytometry showed that overexpression of Ref-1 increased the fraction of
BrdU-negative S phase cells compared with control and AdlacZ infected
cells. At the end of a 12-h phase, the fraction of BrdU-negative S phase cells
in control and AdlacZ-infected cells were 18% and 17%,
respectively. In contrast, the fraction of BrdU-negative S phase cells in
Ref-1-overexpressing cells increased to 29%. Thus even without exposure to
PDGF-BB, overexpression of Ref-1 can affect cell cycle progression.
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To further examine the role of Ref-1 in PDGF-BB-induced SMC proliferation,
and to determine the specificity of the effects of ref-1 AODN on SMCs
progression from G0/G1 to S, cells were treated with
ref-1 AODN in serum-free medium for 24 h and then infected with 200
MOI of Adref-1 or AdlacZ in serum-free medium for 24 h. The
cells were subsequently incubated in serum-free medium for an additional 12 h,
followed by stimulation with PDGF-BB for 9 h. Most of the synchronized cells
(not treated with PDGF-BB or AODN) were in G0/G1 phase,
whereas 14% of the cells were in S phase at time 0
(Fig. 4D). After
stimulation with PDGF-BB for 9 h, only 24% of the ref-1 AODN-treated
cells were in S phase. However, infection of ref-1 AODN-treated cells
with Adref-1 accelerated the progression into S that had been
inhibited by ref-1 AODN: 39% of cells treated with both
Adref-1 and ref-1 AODN were located in S phase 9 h after
stimulation with PDGF-BB. With the MOI used, we did not see visual evidence of
cell killing after adenoviral infection. The specificity of this result was
demonstrated by the results that entry into S remained inhibited in cells
treated with ref-1 AODN + AdlacZ (fraction of S at 9 h of
PDGF treatment was 24%) compared with Ref-1-overexpressing cells pretreated
with ref-1 AODN. Taken together, these results suggest that Ref-1
plays an important role in SMC progression from G0/G1 to
S after PDGF-BB-stimulation and confirm that the effects of ref-1
AODN on PDGF-BB-induced cell cycle progression were due to depletion of Ref-1
protein.
Because Ref-1 has been shown to directly redox-regulate AP-1 activity in vitro (47), we tested whether ref-1 AODN would block PDGF-BB-induced AP-1 DNA binding. With the use of EMSA, we first determined the time course of induction of AP-1 DNA binding activity in SMCs after PDGF treatment. Growth-arrested SMCs were treated with 10 ng/ml of PDGF-BB for 0.5 to 6 h. As shown in Fig. 5A, AP-1 DNA binding began to increase within 0.5–1 h after PDGF-BB treatment, peaked at 2 h, and persisted for at least 6 h. Because PDGF-BB has been shown to primarily induce AP-1 complexes containing Jun-B in SMCs (34), we used anti-Jun-B antibody to supershift AP-1 complexes to confirm that the DNA binding was specifically due to AP-1 (Fig. 5B, lane 1). The arrow indicates the supershifted AP-1 complex. Then, subconfluent SMCs were incubated with ODN in serum-free medium for 24 h and treated with PDGF-BB for 3 h. Preincubation with Ref-1 AODN (Fig. 5, C and D, lanes 4), but not the controls (control ODN: lanes 5–7; vehicle: lane 3; or PDGF-BB alone: lane 2), decreased PDGF-stimulated AP-1 DNA binding activity. To further demonstrate that PDGF-BB-induced AP-1 DNA binding requires the presence of Ref-1 protein, anti-Ref-1 rabbit IgG was used to immunodeplete Ref-1 protein in the nuclear extracts (Fig. 6). As a control, nonimmune rabbit IgG was applied. PDGF-BB-induced AP-1 DNA binding was markedly decreased by treatment with anti-Ref-1 rabbit IgG compared with nonimmune IgG or PDGF-BB alone (Fig. 6, lanes 2, 4, and 6). After immunoprecipitation of nuclear extracts with anti-Ref-1 rabbit IgG, Western blotting of supernatants demonstrated that Jun-B levels were not decreased by the immunoprecipitation process (data not shown), suggesting that the decreased AP-1 DNA binding was due to loss of Ref-1 protein in the nuclear extracts. Furthermore, preincubation of PDGF-BB-treated nuclear extracts from which Ref-1 had been immunodepleted in the presence of 0.5 mM DTT restored the AP-1 DNA binding activity (Fig. 6, lane 6 vs. lane 8), suggesting that Ref-1 acts as a physiological reductant of AP-1, enabling its DNA binding activity in response to PDGF-BB. Collectively, these results indicate that Ref-1 contributes to the regulation of PDGF-BB-induced SMC growth, and part of the pathway could be through redox regulation of AP-1 by Ref-1.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Ref-1 is a ubiquitously expressed multifunctional protein involved in DNA
repair and redox regulation of transcription factors in response to oxidative
stress. Levels of Ref-1 were found to be elevated in several malignant tissues
(25,
30,
48), suggesting that Ref-1
might regulate the rate of tumor cell growth. To determine whether Ref-1
regulates the growth of SMCs in response to PDGF-BB, two different cell
proliferation assays (thymidine uptake and cell cycle analysis) were used to
test the effects of ref-1 AODN. Interestingly, 50% reduction by
ref-1 AODN was observed in both PDGF-induced thymidine uptake and S
entry. The incomplete inhibition observed in our studies may be due to
persistence of some Ref-1 protein in the cells
(Fig. 1), which may have
resulted from incomplete inhibition by AODN. On the other hand, PDGF mitogenic
signaling consists of both redox sensitive and redox independent signaling
pathways (5,
34,
41), suggesting that some
degree of cell proliferation could persist even in the absence of Ref-1
protein. The specificity of effects of ref-1 AODN was confirmed by
the observation that the S entry delay caused by ref-1 AODN was
recovered by Adref-1. We also observed that overexpression of Ref-1
by Adref-1 enhanced SMC entry into S phase in serum-treated cells, in
the absence of stimulation with PDGF-BB, suggesting that Ref-1 modulates SMC
growth induced by factors other than PDGF-BB. Whether overexpression of Ref-1
protein alone is sufficient to induce cell cycle progression, in the absence
of factors that stimulate mitogenic and/or redox signaling, remains to be
determined. Finally, we found that ref-1 AODN decreased the cyclin
D1 protein level, whereas overexpression of Ref-1 increased cyclin
D1 protein levels in SMCs (data not shown). Because cyclin
D1 is an important G1 regulatory protein
(31), these results supported
the fact that Ref-1 plays a critical role in regulating the progression from
the G0/G1 to S phase in SMCs.
AP-1, an inducible transcription factor that recognizes a palindromic
sequence (5'-TGACTCA-3', phorbol
12-O-tetradecanoate-13-acetate-responsive element), is a homodimeric
or heterodimeric leucine zipper complex containing the products of the
fos and jun protooncogenes
(45). AP-1 has been identified
as an intermediary in PDGF-induced proliferation of SMCs
(34). The activity of AP-1 is
controlled not only by transcriptional and posttranscriptional mechanisms via
increased rates of synthesis and decreased rates of degradation of
fos and jun mRNA species, but also by translational and
posttranslational (phosphorylation and redox regulation) mechanisms
(7,
40,
45). The molecule thought to
be directly responsible for redox-activating AP-1 DNA binding is Ref-1
(47), although the glutathione
system and the thioredoxin-Ref-1 pathway have both been shown to participate
in AP-1 transactivation (11,
32). Redox regulation of AP-1
is mediated through a conserved cysteine, Cys-154 in Fos or Cys-272 in Jun,
which is located in the DNA-binding domain of both proteins
(1). Because PDGF has been
shown to stimulate NAD(P)H oxidase-dependent production of
leading to cell growth
(34,
39), and because Ref-1 has
been shown to regulate AP-1 activity during oxidant stress
(8,
44), we asked whether Ref-1
mediates PDGF-induced AP-1 DNA binding activity. Ref-1 protein levels were
reduced in cells by the use of ref-1 AODN or in nuclear extracts by
immunodepletion of Ref-1 with anti-Ref-1 antibody. In both conditions, EMSA
showed that depletion of Ref-1 decreased AP-1 DNA binding induced by PDGF-BB.
These results strongly suggest that Ref-1 participates in PDGF-BB-induced AP-1
transactivation. Moreover, the reduction in AP-1 DNA binding resulting from
immunodepletion of Ref-1 protein in nuclear extracts from PDGF-BB stimulated
cells was recovered by the reducing agent DTT
(Fig. 6, lanes 7 and
8), similar to results reported in response to heat shock in NIH 3T3
cells (8). These findings
suggest that PDGF-BB-induced DNA binding activity of AP-1 complex is dependent
on redox regulation by Ref-1. AP-1 has been shown to downregulate the
cyclin-dependent kinase inhibitor p21Cip1/WAF1 (5a), induce cyclin
D1 (2), and augment
cyclin E expression (28).
Ref-1 may affect these cell cycle regulatory proteins by activating AP-1
activity, which in turn promotes G0/G1-S transition;
future work will be necessary to examine the role of Ref-1 in regulation of
these cell cycle proteins.
We observed that Ref-1 protein levels in whole cell homogenates did not change within 48 h after PDGF-BB treatment, indicating that Ref-1 protein synthesis is most likely not affected by PDGF-BB within 48 h of treatment. Together with the results of the EMSA experiments, it is suggested that PDGF-BB changes the redox status of Ref-1 rather than inducing de novo protein synthesis. In fact, the thioredoxin-Ref-1 signaling pathway has been shown to be independent of de novo protein synthesis; rather, the redox signal is delivered by translocation of thioredoxin and Ref-1 from the cytoplasm to the nucleus to activate transcription factors in response to oxidative stress (19, 42, 44). With the use of immunohistochemistry techniques, Ref-1 has been detected in the cytoplasm in unstimulated SMCs (24). Further studies are required to test whether PDGF-BB causes Ref-1 nuclear translocation in SMCs.
In summary, the results from our studies indicate that Ref-1 has a regulatory role in PDGF-BB stimulated SMC progression from the G0/G1 to S phase and that Ref-1 is an important intermediary in redox regulation of SMC mitogenic signaling.
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DISCLOSURES |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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  G.-M. Zou, M.-H. Luo, A. Reed, M. R. Kelley, and M. C. Yoder Ape1 regulates hematopoietic differentiation of embryonic stem cells through its redox functional domain Blood, March 1, 2007; 109(5): 1917 - 1922. [Abstract] [Full Text] [PDF]
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 D. L. Basi, N. Adhikari, A. Mariash, Q. Li, E. Kao, S. V. Mullegama, and J. L. Hall Femoral artery neointimal hyperplasia is reduced after wire injury in Ref-1+/- mice Am J Physiol Heart Circ Physiol, January 1, 2007; 292(1): H516 - H521. [Abstract] [Full Text] [PDF]
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 A. N. Lyle and K. K. Griendling Modulation of vascular smooth muscle signaling by reactive oxygen species. Physiology, August 1, 2006; 21: 269 - 280. [Abstract] [Full Text] [PDF]
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 R. E. Clempus and K. K. Griendling Reactive oxygen species signaling in vascular smooth muscle cells Cardiovasc Res, July 15, 2006; 71(2): 216 - 225. [Abstract] [Full Text] [PDF]
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 S. Yang, K. Irani, S. E. Heffron, F. Jurnak, and F. L. Meyskens Jr. Alterations in the expression of the apurinic/apyrimidinic endonuclease-1/redox factor-1 (APE/Ref-1) in human melanoma and identification of the therapeutic potential of resveratrol as an APE/Ref-1 inhibitor Mol. Cancer Ther., December 1, 2005; 4(12): 1923 - 1935. [Abstract] [Full Text] [PDF]
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 H. Liu, R. Colavitti, I. I. Rovira, and T. Finkel Redox-Dependent Transcriptional Regulation Circ. Res., November 11, 2005; 97(10): 967 - 974. [Abstract] [Full Text] [PDF]
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 A. Pines, L. Perrone, N. Bivi, M. Romanello, G. Damante, M. Gulisano, M. R. Kelley, F. Quadrifoglio, and G. Tell Activation of APE1/Ref-1 is dependent on reactive oxygen species generated after purinergic receptor stimulation by ATP Nucleic Acids Res., August 2, 2005; 33(14): 4379 - 4394. [Abstract] [Full Text] [PDF]
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 T. Lu, X.-L. Wang, T. He, W. Zhou, T. L. Kaduce, Z. S. Katusic, A. A. Spector, and H.-C. Lee Impaired Arachidonic Acid-Mediated Activation of Large-Conductance Ca2+-Activated K+ Channels in Coronary Arterial Smooth Muscle Cells in Zucker Diabetic Fatty Rats Diabetes, July 1, 2005; 54(7): 2155 - 2163. [Abstract] [Full Text] [PDF]
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