AJP - Renal Track the topics, authors and articles important to you
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Renal Physiol 293: F1889-F1897, 2007. First published September 19, 2007; doi:10.1152/ajprenal.00112.2007
0363-6127/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/6/F1889    most recent
00112.2007v1
Right arrow Alert me when this article is cited
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 Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ding, G.
Right arrow Articles by Yang, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ding, G.
Right arrow Articles by Yang, T.

ANG II induces c-Jun NH2-terminal kinase activation and proliferation of human mesangial cells via redox-sensitive transactivation of the EGFR

Guixia Ding,1,2,* Aihua Zhang,2,3,4,* Songming Huang,2,3 Xiaoqin Pan,2 Guo Zhen,4 Ronghua Chen,2 and Tianxin Yang5

1Department of Pediatrics, Second Affiliated Hospital, 2Center of Pediatric Nephrology, 3Department of Nephrology, and 4Department of Neurology, Nanjing Children's Hospital, Nanjing Medical University, Nanjing, China; and 5Division of Nephrology, University of Utah and Salt Lake Veterans Affairs Medical Center, Salt Lake City, Utah

Submitted 5 March 2007 ; accepted in final form 27 August 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We previously showed that ANG II induces mesangial cell (MC) proliferation via the JNK-activator protein-1 pathway. The present study attempted to determine the upstream mediators of JNK activation, with emphasis on reactive oxygen species (ROS) and the epidermal growth factor (EGF) receptor (EGFR). In cultured human MCs (HMCs), as early as 3 min, ANG II time dependently increased intracellular ROS production, which was sensitive to 10 µM diphenyleneiodonium sulfate and 500 µM apocynin, two structurally distinct NADPH oxidase inhibitors. In contrast, inhibitors of other oxidant-producing enzymes, including the mitochondrial complex I inhibitor rotenone, the xanthine oxidase inhibitor allopurinol, the cyclooxygenase inhibitor indomethacin, the lipoxygenase inhibitor nordihydroguiaretic acid, the cytochrome P-450 oxygenase inhibitor ketoconazole, and the nitric oxide synthase inhibitor NG-nitro-L-arginine methyl ester, were without effect. ANG II-induced ROS generation was inhibited by the angiotensin type 1 receptor antagonist losartan (10 µM) but not the angiotensin type 2 receptor antagonist PD-123319 (10 µM). ANG II induced translocation of p47phox and p67phox from the cytosol to the membrane. The antioxidants almost abolished the ANG II mitogenic response, as assessed by [3H]thymidine incorporation and cell number, associated with a remarkable blockade of the activation of EGFR (90% inhibition) and JNK (83% inhibition). The EGFR inhibitor AG-1478 was able to mimic the effect of antioxidants, in that it inhibited the mitogenic response and the JNK activation following ANG II treatment. Together, these data suggest that the ROS-EGFR-JNK pathway is involved in transducing the proliferative effect of ANG II in cultured HMCs.

glomerular disease; reactive oxygen species; cell growth

CHRONIC KIDNEY DISEASE (CKD) is an emerging top public health problem affecting 20,000,000 Americans. Approximately 1 in 9 adults in the United States have CKD, and another 20,000,000 more are at increased risk. CKD arises from a wide variety of etiologies and develops into a common manifestation of end-stage renal disease characterized by glomerulosclerosis and tubulointerstitial fibrosis and the complete loss of renal function, which require dialysis or kidney transplantation to maintain life (35). The use of inhibitors of the components of the renin-angiotensin axis, namely, angiotensin-converting enzyme inhibitors (ACEi) and angiotensin type 1 (AT1) receptor antagonists (ARBs), has become a standard clinical practice for treatment of CKD (27, 36, 41). Despite intensive investigation, the mechanism of the renal beneficial effects of ACEi and ARBs is incompletely understood. It is clear, however, that these effects of ACEi and ARBs cannot be entirely attributed to their hemodynamic effects (22, 24). Consistent with this notion, ANG II exerts a direct detrimental effect in the renal cells. In particular, ANG II stimulates proliferation of mesangial cells (MCs) (1, 18, 21, 31, 33, 34, 46), a hallmark of the pathological changes in CKD. Therefore, it is imperative to understand the signaling mechanism of the ANG II-elicited growth-promoting effect in the MCs.

MAPKs, a family of Ser/Thr protein kinases, are an important part of intracellular signaling pathways connecting extracellular signals to intracellular regulatory proteins. This family consists of three major members, ERK1/2, JNK, and p38. ERK1/2 is typically stimulated by growth factor acting via a tyrosine kinase receptor and is involved in regulation of cell proliferation, whereas JNK and p38 are more preferentially activated by cellular stress and implicated in the regulation of survival, apoptosis, and differentiation. Indeed, in MCs, ERK1/2 has been implicated in mediating the proliferative effect of basic fibroblast growth factor, one of the major players in MC proliferation (21). However, we recently reported an unexpected finding that MC proliferation in response to ANG II is mediated by JNK, but not ERK1/2 or p38, although all three MAPKs can be activated by ANG II (47). The following question arises: What signaling mechanism is responsible for ANG II-induced JNK activation in human MCs (HMCs).

The predominant signaling events upstream of MAPK activation in response to ANG II in MCs appear to involve generation of reactive oxygen species (ROS) and subsequent transactivation of the epidermal growth factor (EGF) receptor (EGFR). ROS have been identified as a major contributor of ANG II-induced mitogenic effects in vascular smooth muscle cells (VSMCs) as well as MCs (7, 16, 23). The sources of ROS production in response to ANG II include, but may not be limited to, NAD(P)H oxidase (12, 14). The precise mechanism of redox-dependent regulation of cell function, such as cell growth, is not clear given the wide range of proteins that can be targeted by ROS. However, of particular interest is redox-dependent activation of receptor tyrosine kinases (7). EGFR is a redox-sensitive tyrosine kinase that is transactivated by ANG II in VSMCs (8, 44) as well as MCs (29, 32, 42). The transactivation of EGFR by ANG II in MCs involves heparin-binding EGF shedding via metalloproteinase and leads to ERK1/2 activation (42). The goal of the present study was to determine whether redox-dependent EGFR transactivation is responsible for JNK activation and cell growth in response to ANG II treatment.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. ANG II, glucose oxidase (GO), diphenyleneiodonium (DPI), apocynin, rotenone (ROT), allopurinol (ALLO), indomethacin (INDO), NG-nitro-L-arginine methyl ester (L-NAME), nordihydroguiaretic acid (NDGA), ketoconazole (KETO), and 2',7'-dichlorofluorescein diacetate (DCFDA) were obtained from Sigma (St. Louis, MO); AG-1478 from Calbiochem (San Diego, CA); rabbit polyclonal stress-activated protein kinase/JNK, phosphorylated (Thr183/Tyr185) stress-activated protein kinase/JNK, phosphorylated (Ser63) c-Jun, and phosphorylated (Tyr1068) EGFR antibodies from Cell Signaling Technology (Beverly, MA); and polyclonal antibodies against EGFR, p47phox, and p67phox from Santa Cruz Biotechnology (Santa Cruz, CA). All other reagents were purchased from Sigma.

HMC isolation and culture. HMCs were established and characterized in our laboratory as previously reported. Briefly, cells were grown until confluent in RPMI 1640 medium buffered with 10 mmol/l HEPES to pH 7.4 and supplemented with 10% FCS, 5 µg/ml insulin and transferrin, 100 U/ml penicillin, and 100 mg/ml streptomycin. For passage, confluent cells were washed with PBS, removed with 0.025% trypsin-0.5 mM EDTA in PBS, and plated in RPMI 1640 medium. Experiments were performed on cells between passages 5 and 10.

Western blot analysis. At indicated time points, HMCs were rapidly washed with ice-cold PBS and lysed for 10 min on ice in lysis buffer (50 mM Tris, pH 7.5, 40 mM NaCl, 1% Triton X-100, 2 mM EDTA, 1 µg/ml leupeptin, 2 mM DTT, and 1 mM PMSF). Lysates were cleared by centrifugation at 14,000 g (4°C) for 10 min. Total proteins were quantified by the Bradford assay. Equal amounts of lysates were fractionated by 15% SDS-PAGE and electrotransferred to Bio-Blot nitrocellulose membranes (Bio-Rad). The membranes were blocked in TBST (20 mM Tris base, pH 7.6, 150 mM NaCl, and 0.1% Tween 20) containing 5% bovine serum albumin for 1 h at room temperature and incubated overnight with a primary antibody in the blocking solution at 4°C. The membranes were incubated for 1 h with a 1:1,000 dilution of horseradish peroxidase-conjugated secondary antibody at room temperature and visualized using an enhanced chemiluminescence kit (Amersham). The chemiluminescent signal was quantified using UVP software.

DNA synthesis and cell number. DNA synthesis was evaluated by a standard method, namely, [3H]thymidine incorporation (3, 6, 47). Briefly, HMCs were stimulated by GO or ANG II in the presence or absence of losartan, N-acetylcysteine (NAC), DPI, apocynin, or AG-1478 for 19 h and pulsed with 1 µCi/ml [3H]thymidine for 5 h. Cells were then washed twice with ice-cold PBS, incubated for 5 min in 5% TCA, washed with methanol, and dissolved in 99% formic acid. Incorporation of [3H]thymidine into TCA-insoluble material was measured by a liquid scintillation spectrophotometer. For assay of cell growth, HMCs in six-well plates were treated as described above, and cells were counted using a Coulter counter.

DCFDA measurement of ROS. The fluorogenic substrate DCFDA, a cell-permeable dye that is oxidized to highly fluorescent DCF by H2O2, has been commonly used to monitor intracellular generation of ROS (13, 17, 43, 48, 49). For measurement of ROS, cells were grown on glass coverslips. When the cells reached confluence, they were washed twice with PBS and incubated for 30 min with 50 µM DCFDA diluted in RPMI 1640 medium with 10% FCS. Then the cells were treated with ANG II in the presence or absence of DPI, apocynin, ROT, ALLO, INDO, L-NAME, NDGA, or KETO. At the end of the incubation period, the cells were washed twice with PBS and imaged using confocal laser microscopy. For quantitation of ROS levels, the cells were seeded onto a 96-well plate and treated as described above. Relative fluorescence was measured using a fluorescence plate reader (FLUOstar OPTIMA) at excitation and emission wavelengths of 485 and 528 nm, respectively. The following agents were used to assess potential sources of ROS production: DPI, a flavoprotein inhibitor (10 µM); apocynin, an NADPH oxidase inhibitor (500 µM); the complex I mitochondrial electron chain inhibitor ROT (10 µM); the xanthine oxidase inhibitor ALLO (30 µM); the cyclooxygenase inhibitor INDO (100 µM); the lipoxygenase inhibitor NDGA (10 µM); the cytochrome P-450 oxygenase inhibitor KETO (10 µM); and the nitric oxide synthase inhibitor L-NAME (100 µM).

Lucigenin chemiluminescence. Superoxide (O2bullet–) production in cell homogenates was measured with lucigenin-enhanced chemiluminescence in a microplate luminometer (Anthos Lucy 1). Briefly, HMCs were detached, washed in PBS, and resuspended in 400 µl of buffer (50 mmol/l KH2PO4, 1 mmol/l EGTA, and 150 mmol/l sucrose, pH 7.0) with a protease inhibitor cocktail. Cells were sonicated and distributed in triplicate (10 µg/well) onto a 96-well microplate. NADPH (300 µmol/l) and dark-adapted lucigenin (5 µmol/l) were added to wells just before they were read. O2bullet– production was expressed as arbitrary light units over 20 min.

Statistical analysis. Values are means ± SE from at least three separate experiments. Statistical comparisons between two groups and multiple groups were done by Student's t-test and one-way ANOVA, respectively, using SPSS 10.0. P < 0.05 was considered to represent statistical significance.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effect of ANG II on ROS production. Using DCFDA and fluorescence microscopy, we initially validated the time course of ROS production in response to ANG II treatment in cultured HMCs. As shown in Fig. 1, ANG II elevated the intensity of fluorescence (n = 8 for each) as early as 3 min, which gradually increased and peaked at 1 h.


Figure 1
View larger version (77K):
[in this window]
[in a new window]

  		Fig. 1. ANG II-induced reactive oxygen species (ROS) generation in human mesangial cells (HMCs). A–H: confluent HMCs in chamber slides were treated with vehicle (A-D) or 100 nM ANG II (E-H) in the presence of 2',7'-dichloro-fluorescein (DCF) for 15 (A and E), 30 (B and F), 60 min (C and G), and 120 (D and H) min, and images were obtained using a fluorescent microscope. I: quantitation of ROS production after 100 nM ANG II. Confluent HMCs in 96-well plates were exposed to vehicle or 100 nM ANG II in the presence of DCF. At 3, 15, 30, 60, and 120 min, fluorescence was quantified using FLUOstar OPTIMA. J: effect of angiotensin receptor (AT1R and AT2R) antagonists on ANG II-induced ROS production. Confluent HMCs in 96-well plates were pretreated for 30 min with losartan or PD-123319 and incubated for 60 min with ANG II in the presence of DCF. Values are means ± SE (n = 8). #P < 0.05; *P < 0.01 vs. corresponding control. {dagger}P < 0.01 vs. ANG II.

 
HMCs express AT1 and AT2 receptors. To examine the type of receptors involved, HMCs were treated with the AT1 antagonist losartan (10 µM) or the AT2 antagonist PD-123319 (10 µM) for 30 min and then with 100 nM ANG II for 1 h. ANG II-induced ROS production was inhibited by losartan, but not PD-123319 (Fig. 1J), indicating involvement of AT1, but not AT2, receptors.

Enzymatic sources of ANG II-induced ROS production. To determine the enzymatic sources of ROS, we measured ANG II-induced ROS production in the presence and absence of inhibitors of various oxidant-producing enzymes. As shown in Fig. 2, DPI and apocynin, two structurally distinct NADPH oxidase inhibitors, almost completely blocked ANG II-induced ROS generation in HMCs. In contrast, inhibitors of other oxidant-producing enzyme systems, including ROT, ALLO, INDO, NDGA, KETO, and L-NAME, were without effect.


Figure 2
View larger version (36K):
[in this window]
[in a new window]

  		Fig. 2. Enzymatic sources of ROS production in response to ANG II treatment. Confluent HMCs in chamber slides were pretreated for 30 min with 500 µM apocynin, 10 µM diphenyleneiodonium (DPI), 10 µM rotenone (ROT), 30 µM allopurinol (ALLO), 100 µM indomethacin (INDO), 10 µM nordihydroguiaretic acid (NDGA), 10 µM ketoconazole (KETO), or 100 µM NG-nitro-L-arginine methyl ester (L-NAME) and, for a further 60 min, with 100 nM ANG II in the presence of DCF. A: control; B: ANG II; C: ANG II + apocynin; D: ANG II + DPI; E: ANG II + ROT; F: ANG II + ALLO; G: ANG II + INDO; H: ANG II + NDGA; I: ANG II + KETO; J: ANG II + L-NAME. For quantification of ROS production, confluent HMCs in 96-well plates were pretreated for 30 min with apocynin, DPI, ROT, ALLO, INDO, NDGA, KETO, or L-NAME and incubated for 60 min with ANG II in the presence of DCF. K: quantification of fluorescence using FLUOstar OPTIMA. Values are means ± SE (n = 8). *P < 0.01 vs. ANG II.

 
Effect of ANG II on NADPH oxidase activity. Given the pharmacological evidence for involvement of NADPH oxidase in ANG II-induced ROS generation, we examined the effect of ANG II on NADPH oxidase activity in cultured HMCs. Using lucigenin chemiluminescence, we detected a very low level of O2bullet– in cultured HMCs in the absence of NADPH [0.52 ± 0.10 relative light units (RLU)·min–1·mg protein–1]. Addition of 100 µM NADPH resulted in a substantial increase in the level of O2bullet– (6.00 ± 0.69 RLU·min–1·mg protein–1). Treatment with 100 nM ANG II for 2 h elevated NADPH-dependent O2bullet– production (11.66 ± 0.80 RLU·min–1·mg protein–1). Stimulation of O2bullet– production was markedly attenuated by preincubation with losartan (6.72 ± 0.55 RLU·min–1·mg protein–1) and was abolished by apocynin and DPI (2.42 ± 0.39 and 2.52 ± 0.35 RLU·min–1·mg protein–1, respectively, n = 6 for each group; Fig. 3A).


Figure 3
View larger version (22K):
[in this window]
[in a new window]

  		Fig. 3. NADPH oxidase activity and membranous translocation of p47phox and p67phox in HMCs. A: NADPH oxidase activity. Confluent HMCs in 6-well plates were pretreated for 30 min with 10 µM losartan, 500 µM apocynin, or 10 µM DPI and, for a further 2 h, with 100 nM ANG II, and NADPH oxidase-dependent O2bullet– production by intact HMCs was measured by lucigenin-enhanced chemiluminescence. RLU, relative light units. B and C: membranous translocation of p47phox and p67phox. HMCs were treated as described in A, and fractionated membranous proteins and whole cell lysates were subjected to immunoblot analysis of p47phox and p67phox. Values are means ± SE (n = 3). *P < 0.01 vs. control. {dagger}P < 0.01 vs. ANG II.

 
Activation of NADPH oxidase requires translocation of the cytosolic components, including p47phox and p67phox, to the cell membrane. Therefore, using immunoblotting analysis of fractionated cellular proteins, we examined the effects of ANG II on membranous translocation of p47phox and p67phox in HMCs. As shown in Fig. 4, B and C, treatment with 100 nM ANG II for 2 h induced increased p47phox and p67phox protein levels in the membranous fraction by 3.72- and 3.86-fold, respectively, but had no effect on their total protein abundance. ANG II-induced membranous translocation of p47phox and p67phox was markedly attenuated by 10 µM losartan, 500 µM apocynin, or 10 µM DPI (Fig. 3, B and C).


Figure 4
View larger version (9K):
[in this window]
[in a new window]

  		Fig. 4. Effects of glucose oxidase (GO) on proliferation of HMCs. A: [3H]thymidine incorporation. Cells were grown to confluence, washed twice, and incubated in serum-free medium for 24 h. Then cells were incubated with new medium in the presence or absence of 1 mU/ml GO for 24 h, and [3H]thymidine incorporation into cells was determined. B: cells were treated as described in A, and cells were counted. Values are means ± SE of 4 independent experiments. *P < 0.01 vs. control.

 
Role of ROS in ANG II-induced HMC proliferation. To test the role of ROS in ANG II-induced HMC proliferation, we performed experiments to determine 1) whether exogenous ROS stimulated HMC proliferation, mimicking the effect of ANG II, and 2) whether inhibition of endogenous ROS production by antioxidants blocked the effect of ANG II. We first examined responsiveness of HMCs to exogenous ROS produced by the glucose-GO system, which is known to continuously produce H2O2 in a glucose-containing medium (10, 38). Addition of GO (1 mU/ml) significantly induced HMC proliferation, as determined by [3H]thymidine incorporation and cell number (Fig. 4). We subsequently examined the role of ROS in ANG II-induced HMC proliferation. HMCs were pretreated with losartan, NAC, or the NADPH oxidase inhibitor apocynin and DPI for 30 min and then incubated with 100 nM ANG II for 48 h. Losartan, NAC, apocynin, and DPI effectively inhibited ANG II-induced HMC proliferation (Fig. 5).


Figure 5
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 5. Role of reactive oxygen species (ROS) in ANG II-induced proliferative effect in HMCs. A: [3H]thymidine incorporation. Quiescent cells were treated with 100 nM ANG II in the presence or absence of 10 µM losartan, 5 mM N-acetylcysteine (NAC), 500 µM apocynin, or 10 µM DPI for 48 h. B: cells were treated as described in A, and cells were counted. Values are means ± SE of 4 independent experiments. *P < 0.01 vs. control. #P < 0.05; {dagger}P < 0.01 vs. ANG II.

 
Involvement of ROS in ANG II-induced transactivation of EGFR. Transactivation of EGFR is a major mechanism responsible for the growth-promoting effect of ANG II. We examined the role of ROS in ANG II-induced EGFR transactivation. We first examined the transactivation kinetics of EGFR after treatment with 100 nM ANG II. EGFR transactivation was determined by immunoblotting with use of an antibody recognizing the phosphorylated (Tyr1068) EGFR that is responsible for MAPK activation and EGFR activity. ANG II increased the level of phosphorylated EGFR at 5 min, with maximal stimulation at 30 min (Fig. 6A). Therefore, the 30-min time point was used in subsequent experiments for determination of the dose response of ANG II stimulation of EGFR. As shown in Fig. 6B, ANG II increased phosphorylation of EGFR in a dose-dependent manner, with maximal stimulation at 100 nM. Pretreatment with losartan significantly blocked ANG II-induced EGFR transactivation (Fig. 7). We then examined the effects of antioxidants on ANG II-induced EGFR transactivation. NAC, apocynin, or DPI completely blocked ANG II-induced phosphorylation of EGFR (Fig. 7).


Figure 6
View larger version (12K):
[in this window]
[in a new window]

  		Fig. 6. Transactivation of epidermal growth factor receptor (EGFR) by ANG II in HMCs. Cells were treated with 100 nM ANG II for 5, 15, and 30 min (A) or with 1–100 nM ANG II (B). At 5, 15, and 30 min, whole cell lysates were subjected to immunoblot analysis of phosphorylated (Tyr1068) EGFR (phospho-EGFR) and total EGFR. Values are means ± SE (n = 3). *P < 0.05; {dagger}P < 0.01 vs. control.

 

Figure 7
View larger version (15K):
[in this window]
[in a new window]

  		Fig. 7. Role of ROS in ANG II-induced EGFR transactivation. Confluent HMCs in 6-well plates were pretreated for 30 min with 10 µM losartan, 5 mM NAC, 500 µM apocynin, or 10 µM DPI and, for a further 30 min, with 100 nM ANG II. Then whole cell lysates were subjected to immunoblot analysis of phosphorylated (Tyr1068) EGFR and total EGFR. Values are means ± SE (n = 3). *P < 0.01 vs. control.

 
Dependence of ANG II-induced JNK activation and HMC proliferation on EGFR transactivation. Our previous study demonstrated an essential role of JNK in mediating the growth-promoting effect of ANG II in cultured HMCs. We examined a possible link between JNK and EGFR in ANG II signal transduction in these cells. Phosphorylation of JNK and c-Jun by immunoblotting was determined using antibodies against phosphorylated (Thr183/Tyr185) JNK and phosphorylated (Ser63) c-Jun, respectively. JNK activation was significantly inhibited by AG-1478, as well as losartan, NAC, apocynin, and DPI (Fig. 8). Consistent with this finding, ANG II-induced HMC proliferation was completely inhibited by AG-1478 (Fig. 9).


Figure 8
View larger version (14K):
[in this window]
[in a new window]

  		Fig. 8. JNK activation. HMCs were pretreated for 30 min with 10 µM losartan, 5 nM NAC, 500 µM apocynin, 10 µM DPI, or 250 nM AG-1478 and then treated with 100 nM ANG II for 30 min. Parallel immunoblots were performed on the same cell lysates using antibodies against phosphorylated (Thr183/Tyr185) JNK (pp54 and pp46), phosphorylated (Ser63) c-Jun (pc-Jun), and JNK. Anti-JNK antibody was used as loading control. Relative abundance of phosphorylated (Ser63) c-Jun band was evaluated by densitometry. Results are expressed as fold increase over control. Values are means ± SE of 4 independent experiments. *P < 0.01 vs. control.

 

Figure 9
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 9. Effects of EGFR antagonist AG-1478 on ANG II-induced HMC proliferation. A: [3H]thymidine incorporation. B: cell count. Values are means ± SE of 4 independent experiments. *P < 0.01 vs. ANG II.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
ANG II stimulates MC proliferation, but the underlying mechanism remains incompletely understood. Our previous study provides evidence for involvement of JNK, but not ERK1/2 or p38, in the ANG II-induced mitogenic effect in cultured HMCs (47). All three MAPKs were activated by ANG II, but only the JNK inhibitor SP-600125 blocked the mitogenic effect of ANG II. The present study extended this observation by identifying the upstream mediators of JNK activation, including ROS and EGFR. We found that ANG II, via AT1, but not AT2, receptors, increased intracellular ROS production in cultured HMCs associated with membranous translocation of p47phox and p67phox. ANG II-induced ROS release was almost completely blocked by DPI and apocynin, two structurally distinct NADPH oxidase inhibitors, but was unaffected by inhibitors of the other oxidant-producing enzyme systems. The NADPH oxidase inhibitors and NAC remarkably inhibited JNK activation, as well as MC proliferation, in response to ANG II treatment. These findings suggest involvement of NADPH oxidase-derived ROS in ANG II-induced JNK activation in cultured HMCs. ROS appeared to activate JNK through EGFR, inasmuch as antioxidants prevented the transactivation of EGFR and inhibition of EGFR by AG-1478 significantly inhibited JNK activation. Together, these observations suggest that the ROS-EGFR-JNK pathway transduces the signaling of the ANG II-induced proliferating effect in cultured HMCs.

Previously considered as only toxic by-products of metabolisms, ROS are now well appreciated as important signaling molecules involved in a wide variety of physiological and pathological processes (4, 20, 39). ROS have been shown to mediate MC contraction (40), hypertrophy (14), and apoptosis (26) in response to ANG II treatment. Therefore, it is of critical importance to clarify the source of ANG II-induced ROS generation in MCs. Additionally, no previous studies have directly addressed the role of ROS in the ANG II-induced mitogenic effect in these cells. Therefore, a major aim of the present study was to determine the source of ROS generation relevant to the ANG II-induced mitogenic effect in cultured HMCs. NADPH oxidase is a multicomponent enzyme consisting of the membrane-associated subunits p22phox and gp91phox and the cytosolic subunits p47phox, p67phox, and p40phox, and the activity of the enzyme requires translocation of the cytosolic complex to the membrane (9, 11, 19). In recent years, four homologs of gp91phox (Nox2), Nox1 (37), Nox3, Nox4, and Nox5 (25), have been identified. NADPH oxidase is a major O2bullet–-generating enzyme in phagocytes as well as nonphagocytic cells. After NADPH oxidase, there is a growing list of other oxidant-generating systems that contribute to ROS production under various conditions; these systems include the mitochondrial respiratory chain, xanthine oxidase, cyclooxygenase, lipoxygenase, cytochrome P-450, and nitric oxide synthase. We found that ANG II-induced ROS release was almost completely blocked by the NADPH oxidase inhibitors DPI and apocynin, but the mitochondrial complex I inhibitor ROT, the xanthine oxidase inhibitor ALLO, the cyclooxygenase inhibitor INDO, the lipoxygenase inhibitor NDGA, the cytochrome P-450 oxygenase inhibitor KETO, and the nitric oxide synthase inhibitor L-NAME were without effect. To our knowledge, this is the most comprehensive investigation of the respective contribution of various oxidant systems to ANG II-induced ROS production in cultured MCs. Not only did the NADPH oxidase inhibitors inhibit ROS production, but these compounds also blocked the proliferative effect of ANG II. In addition to the pharmacological evidence favoring NADPH oxidase as a major source of ANG II-induced ROS production, we also provided evidence for translocation of p47phox and p67phox to the membrane in response to ANG II treatment. These findings are in agreement with the previous observation that ANG II activates NADPH oxidase in cultured MCs (2, 15).

The molecular target of ROS in the ANG II-elicited signaling pathway in cultured HMCs appears to be EGFR. In support of this notion, exposure of HMCs to ANG II activated EGFR, as evidenced by a rapid increase in phosphorylation of the receptor. Activation of EGFR was completely prevented by antioxidants, including inhibitors of NADPH oxidase. Furthermore, the EGFR inhibitor AG-1478 was able to mimic the effect of antioxidants in attenuating ANG II-induced MC proliferation. It has been reported that the redox-dependent transactivation of EGFR is a major mechanism by which ANG II induces proliferative/hypertrophic effects in VSMCs (30, 44, 45). Our findings suggest that the same mechanism may be operative in cultured HMCs.

We have provided evidence that JNK represents a downstream mediator of ROS-dependent activation of EGFR in the ANG II-induced signaling pathway in cultured HMCs. This notion is based on the observation that ANG II-induced activation of JNK was inhibited by antioxidants and the EGFR inhibitor AG-1478. This is somewhat different from the observation in VSMCs (5) and rat neonatal cardiomyocytes (28) that transactivation of EGFR is required for ANG II-induced activation of ERK1/2 and p38, but not JNK. The reason for this discrepancy is unclear but may indicate a cell type-specific phenomenon.

In summary, our previous study demonstrated that JNK mediates ANG II-induced HMC proliferation, and the present study has extended this observation by examining ROS and EGFR as upstream mediators of the JNK activation. We found that ANG II, via AT1, but not AT2, receptors increased NADPH oxidase-dependent production of ROS, which activates EGFR, leading to activation of JNK. The elucidation the proliferative response to ANG II in HMCs of the ROS-EGFR-JNK pathway provides insights into mechanisms of MC proliferation and may also help identify new targets for treatment of glomerular diseases.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Natural Science Foundation of China Grant 30100081 (to A. Zhang), Natural Science Foundation of Jiangsu Province Grants BK2004144 (to A. Zhang), BS2003050 (to G. Ding), and BK2007259 (to G. Ding), and Nanjing Bureau of Health Grant ZKX0408 (to G. Zhen).


   FOOTNOTES
 

Address for reprint requests and other correspondence: T. Yang, Univ. of Utah and Veterans Affairs Medical Center, Division of Nephrology and Hypertension, 30 N 1900 E, Rm. 4R312, Salt Lake City, UT 84132 (e-mail: tianxin.yang{at}hsc.utah.edu; or R. Chen, Center of Pediatric Nephrology, Nanjing Medical Univ., Nanjing 210029, China (e-mail: rchen{at}njmu.edu.ch)

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.

* G. Ding and A. Zhang contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Ardaillou R, Chansel D, Chatziantoniou C, Dussaule JC. Mesangial AT1 receptors: expression, signaling, and regulation. J Am Soc Nephrol 10 Suppl 11: S40–S46, 1999.[CrossRef][Web of Science][Medline]
  2. Block K, Ricono JM, Lee DY, Bhandari B, Choudhury GG, Abboud HE, Gorin Y. Arachidonic acid-dependent activation of a p22phox-based NAD(P)H oxidase mediates angiotensin II-induced mesangial cell protein synthesis and fibronectin expression via Akt/PKB. Antioxid Redox Signal 8: 1497–1508, 2006.[CrossRef][Web of Science][Medline]
  3. Burton GF, Conrad DH, Szakal AK, Tew JG. Follicular dendritic cells and B cell costimulation. J Immunol 150: 31–38, 1993.[Abstract]
  4. Droge W. Free radicals in the physiological control of cell function. Physiol Rev 82: 47–95, 2002.[Abstract/Free Full Text]
  5. Eguchi S, Dempsey PJ, Frank GD, Motley ED, Inagami T. Activation of MAPKs by angiotensin II in vascular smooth muscle cells. Metalloprotease-dependent EGF receptor activation is required for activation of ERK and p38 MAPK but not for JNK. J Biol Chem 276: 7957–7962, 2001.[Abstract/Free Full Text]
  6. Evans SS, Ozer H. Enhancement of a human antibody response in vitro mediated by interaction of interferon-{alpha} with T lymphocytes. J Immunol 138: 2451–2456, 1987.[Abstract]
  7. Frank GD, Eguchi S. Activation of tyrosine kinases by reactive oxygen species in vascular smooth muscle cells: significance and involvement of EGF receptor transactivation by angiotensin II. Antioxid Redox Signal 5: 771–780, 2003.[CrossRef][Web of Science][Medline]
  8. Frank GD, Eguchi S, Inagami T, Motley ED. N-acetylcysteine inhibits angiotensin II-mediated activation of extracellular signal-regulated kinase and epidermal growth factor receptor. Biochem Biophys Res Commun 280: 1116–1119, 2001.[CrossRef][Web of Science][Medline]
  9. Gallin JI, Leto TL, Rotrosen D, Kwong CH, Malech HL. Delineation of the phagocyte NADPH oxidase through studies of chronic granulomatous diseases of childhood. Curr Opin Immunol 4: 53–56, 1992.[CrossRef][Web of Science][Medline]
  10. Garberg P, Hogberg J. Studies on Se incorporation in selenoproteins: effects of peroxisome proliferators and hydrogen peroxide-generating system. Chem Biol Interact 81: 291–306, 1992.[CrossRef][Web of Science][Medline]
  11. Geiszt M, Leto TL. The Nox family of NAD(P)H oxidases: host defense and beyond. J Biol Chem 279: 51715–51718, 2004.[Free Full Text]
  12. Gill PS, Wilcox CS. NADPH oxidases in the kidney. Antioxid Redox Signal 8: 1597–1607, 2006.[CrossRef][Web of Science][Medline]
  13. Gonzalez CF, Ackerley DF, Lynch SV, Matin A. ChrR, a soluble quinone reductase of Pseudomonas putida that defends against H2O2. J Biol Chem 280: 22590–22595, 2005.[Abstract/Free Full Text]
  14. Gorin Y, Block K, Hernandez J, Bhandari B, Wagner B, Barnes JL, Abboud HE. Nox4 NAD(P)H oxidase mediates hypertrophy and fibronectin expression in the diabetic kidney. J Biol Chem 280: 39616–39626, 2005.[Abstract/Free Full Text]
  15. Gorin Y, Ricono JM, Kim NH, Bhandari B, Choudhury GG, Abboud HE. Nox4 mediates angiotensin II-induced activation of Akt/protein kinase B in mesangial cells. Am J Physiol Renal Physiol 285: F219–F229, 2003.[Abstract/Free Full Text]
  16. Griendling KK, Ushio-Fukai M. Reactive oxygen species as mediators of angiotensin II signaling. Regul Pept 91: 21–27, 2000.[CrossRef][Web of Science][Medline]
  17. Hickson-Bick DL, Sparagna GC, Buja LM, McMillin JB. Palmitate-induced apoptosis in neonatal cardiomyocytes is not dependent on the generation of ROS. Am J Physiol Heart Circ Physiol 282: H656–H664, 2002.[Abstract/Free Full Text]
  18. Higueruelo S, Romero R. Angiotensin II requires PDGF-BB to induce DNA synthesis in rat mesangial cells cultured in an exogenous insulin-free medium. Nephrol Dial Transplant 12: 694–700, 1997.[Abstract/Free Full Text]
  19. Hoidal JR, Brar SS, Sturrock AB, Sanders KA, Dinger B, Fidone S, Kennedy TP. The role of endogenous NADPH oxidases in airway and pulmonary vascular smooth muscle function. Antioxid Redox Signal 5: 751–758, 2003.[CrossRef][Web of Science][Medline]
  20. Irani K. Oxidant signaling in vascular cell growth, death, and survival: a review of the roles of reactive oxygen species in smooth muscle and endothelial cell mitogenic and apoptotic signaling. Circ Res 87: 179–183, 2000.[Abstract/Free Full Text]
  21. Izevbigie EB, Gutkind JS, Ray PE. Angiotensin II and basic fibroblast growth factor mitogenic pathways in human fetal mesangial cells. Pediatr Res 47: 614–621, 2000.[Web of Science][Medline]
  22. Jafar TH, Schmid CH, Landa M, Giatras I, Toto R, Remuzzi G, Maschio G, Brenner BM, Kamper A, Zucchelli P, Becker G, Himmelmann A, Bannister K, Landais P, Shahinfar S, de Jong PE, de Zeeuw D, Lau J, Levey AS. Angiotensin-converting enzyme inhibitors and progression of nondiabetic renal disease. A meta-analysis of patient-level data. Ann Intern Med 135: 73–87, 2001.[Abstract/Free Full Text]
  23. Jaimes EA, Tian RX, Pearse D, Raij L. Up-regulation of glomerular COX-2 by angiotensin II: role of reactive oxygen species. Kidney Int 68: 2143–2153, 2005.[CrossRef][Web of Science][Medline]
  24. Kakinuma Y, Kawamura T, Bills T, Yoshioka T, Ichikawa I, Fogo A. Blood pressure-independent effect of angiotensin inhibition on vascular lesions of chronic renal failure. Kidney Int 42: 46–55, 1992.[Web of Science][Medline]
  25. Lambeth JD. NOX enzymes and the biology of reactive oxygen. Nat Rev Immunol 4: 181–189, 2004.[CrossRef][Web of Science][Medline]
  26. Lodha S, Dani D, Mehta R, Bhaskaran M, Reddy K, Ding G, Singhal PC. Angiotensin II-induced mesangial cell apoptosis: role of oxidative stress. Mol Med 8: 830–840, 2002.[Web of Science][Medline]
  27. Mackenzie HS, Ziai F, Omer SA, Nadim MK, Taal MW. Angiotensin receptor blockers in chronic renal disease: the promise of a bright clinical future. J Am Soc Nephrol 10 Suppl 12: S283–S286, 1999.[Web of Science][Medline]
  28. Nishida M, Tanabe S, Maruyama Y, Mangmool S, Urayama K, Nagamatsu Y, Takagahara S, Turner JH, Kozasa T, Kobayashi H, Sato Y, Kawanishi T, Inoue R, Nagao T, Kurose H. G{alpha}12/13- and reactive oxygen species-dependent activation of c-Jun NH2-terminal kinase and p38 mitogen-activated protein kinase by angiotensin receptor stimulation in rat neonatal cardiomyocytes. J Biol Chem 280: 18434–18441, 2005.[Abstract/Free Full Text]
  29. Nose A, Mori Y, Uchiyama-Tanaka Y, Kishimoto N, Maruyama K, Matsubara H, Iwasaka T. Regulation of glucose transporter (GLUT1) gene expression by angiotensin II in mesangial cells: involvement of HB-EGF and EGF receptor transactivation. Hypertens Res 26: 67–73, 2003.[CrossRef][Web of Science][Medline]
  30. Ohtsu H, Dempsey PJ, Frank GD, Brailoiu E, Higuchi S, Suzuki H, Nakashima H, Eguchi K, Eguchi S. ADAM17 mediates epidermal growth factor receptor transactivation and vascular smooth muscle cell hypertrophy induced by angiotensin II. Arterioscler Thromb Vasc Biol 26: e133–e137, 2006.[Abstract/Free Full Text]
  31. Orth SR, Weinreich T, Bonisch S, Weih M, Ritz E. Angiotensin II induces hypertrophy and hyperplasia in adult human mesangial cells. Exp Nephrol 3: 23–33, 1995.[Web of Science][Medline]
  32. Perlman A, Lawsin LM, Kolachana P, Saji M, Moore J Jr, Ringel MD. Angiotensin II regulation of TGF-β in murine mesangial cells involves both PI3 kinase and MAP kinase. Ann Clin Lab Sci 34: 277–286, 2004.[Abstract/Free Full Text]
  33. Ray PE, Aguilera G, Kopp JB, Horikoshi S, Klotman PE. Angiotensin II receptor-mediated proliferation of cultured human fetal mesangial cells. Kidney Int 40: 764–771, 1991.[Web of Science][Medline]
  34. Ray PE, Bruggeman LA, Horikoshi S, Aguilera G, Klotman PE. Angiotensin II stimulates human fetal mesangial cell proliferation and fibronectin biosynthesis by binding to AT1 receptors. Kidney Int 45: 177–184, 1994.[Web of Science][Medline]
  35. Schieppati A, Remuzzi G. Chronic renal diseases as a public health problem: epidemiology, social, and economic implications. Kidney Int Suppl: S7–S10, 2005.
  36. Strippoli GF, Bonifati C, Craig M, Navaneethan SD, Craig JC. Angiotensin converting enzyme inhibitors and angiotensin II receptor antagonists for preventing the progression of diabetic kidney disease. Cochrane Database Syst Rev: CD006257, 2006.
  37. Suh YA, Arnold RS, Lassegue B, Shi J, Xu X, Sorescu D, Chung AB, Griendling KK, Lambeth JD. Cell transformation by the superoxide-generating oxidase Mox1. Nature 401: 79–82, 1999.[CrossRef][Medline]
  38. Tenopoulou M, Doulias PT, Barbouti A, Brunk U, Galaris D. Role of compartmentalized redox-active iron in hydrogen peroxide-induced DNA damage and apoptosis. Biochem J 387: 703–710, 2005.[CrossRef][Web of Science][Medline]
  39. Thannickal VJ, Fanburg BL. Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 279: L1005–L1028, 2000.[Abstract/Free Full Text]
  40. Torrecillas G, Boyano-Adanez MC, Medina J, Parra T, Griera M, Lopez-Ongil S, Arilla E, Rodriguez-Puyol M, Rodriguez-Puyol D. The role of hydrogen peroxide in the contractile response to angiotensin II. Mol Pharmacol 59: 104–112, 2001.[Abstract/Free Full Text]
  41. Toto RD. Treatment of hypertension in chronic kidney disease. Semin Nephrol 25: 435–439, 2005.[CrossRef][Web of Science][Medline]
  42. Uchiyama-Tanaka Y, Matsubara H, Nozawa Y, Murasawa S, Mori Y, Kosaki A, Maruyama K, Masaki H, Shibasaki Y, Fujiyama S, Nose A, Iba O, Hasagawa T, Tateishi E, Higashiyama S, Iwasaka T. Angiotensin II signaling and HB-EGF shedding via metalloproteinase in glomerular mesangial cells. Kidney Int 60: 2153–2163, 2001.[CrossRef][Web of Science][Medline]
  43. Ueom J, Kwon S, Kim S, Chae Y, Lee K. Acquisition of heat shock tolerance by regulation of intracellular redox states. Biochim Biophys Acta 1642: 9–16, 2003.[Medline]
  44. Ushio-Fukai M, Griendling KK, Becker PL, Hilenski L, Halleran S, Alexander RW. Epidermal growth factor receptor transactivation by angiotensin II requires reactive oxygen species in vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 21: 489–495, 2001.[Abstract/Free Full Text]
  45. Ushio-Fukai M, Zuo L, Ikeda S, Tojo T, Patrushev NA, Alexander RW. cAbl tyrosine kinase mediates reactive oxygen species- and caveolin-dependent AT1 receptor signaling in vascular smooth muscle: role in vascular hypertrophy. Circ Res 97: 829–836, 2005.[Abstract/Free Full Text]
  46. Wolthuis A, Boes A, Rodemann HP, Grond J. Vasoactive agents affect growth and protein synthesis of cultured rat mesangial cells. Kidney Int 41: 124–131, 1992.[Web of Science][Medline]
  47. Zhang A, Ding G, Huang S, Wu Y, Pan X, Guan X, Chen R, Yang T. c-Jun NH2-terminal kinase mediation of angiotensin II-induced proliferation of human mesangial cells. Am J Physiol Renal Physiol 288: F1118–F1124, 2005.[Abstract/Free Full Text]
  48. Zhang A, Dong Z, Yang T. Prostaglandin D2 inhibits TGF-β1-induced epithelial-to-mesenchymal transition in MDCK cells. Am J Physiol Renal Physiol 291: F1332–F1342, 2006.[Abstract/Free Full Text]
  49. Zhang A, Wang MH, Dong Z, Yang T. Prostaglandin E2 is a potent inhibitor of epithelial-to-mesenchymal transition: interaction with hepatocyte growth factor. Am J Physiol Renal Physiol 291: F1323–F1331, 2006.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 J. Am. Soc. Nephrol.Home page
D. Schlondorff and B. Banas
The Mesangial Cell Revisited: No Cell Is an Island
J. Am. Soc. Nephrol., June 1, 2009; 20(6): 1179 - 1187.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Renal Physiol.Home page
S. Huang, A. Zhang, G. Ding, and R. Chen
Aldosterone-induced mesangial cell proliferation is mediated by EGF receptor transactivation
Am J Physiol Renal Physiol, June 1, 2009; 296(6): F1323 - F1333.
[Abstract] [Full Text] [PDF]

Home page
 HypertensionHome page
B. Shen, M. Hagiwara, Y.-Y. Yao, L. Chao, and J. Chao
Salutary Effect of Kallistatin in Salt-Induced Renal Injury, Inflammation, and Fibrosis via Antioxidative Stress
Hypertension, May 1, 2008; 51(5): 1358 - 1365.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/6/F1889    most recent
00112.2007v1
Right arrow Alert me when this article is cited
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 Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ding, G.
Right arrow Articles by Yang, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ding, G.
Right arrow Articles by Yang, T.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.