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Cell Biology International (2008) 32, 1536–1545 (Printed in Great Britain)
2,4-Dinitrophenol induces apoptosis in As4.1 juxtaglomerular cells through rapid depletion of GSH
Yong Hwan Han, Sung Zoo Kim, Suhn Hee Kim and Woo Hyun Park*
Department of Physiology, Medical School, Institute for Medical Sciences, Center for Healthcare Technology Development, Chonbuk National University, Jeonju 561-180, Republic of Korea


Abstract

2,4-Dinitrophenol (DNP) is an uncoupler of oxidative phosphorylation in mitochondria. Here, we investigated the in vitro effect of DNP on apoptosis and the involvement of reactive oxygen species (ROS) in As4.1 juxtaglomerular cell death. Dose- and time-dependent induction of apoptosis was evidenced by flow cytometric detection of sub-G1 DNA content and annexin V binding assay. The intracellular H2O2 and O2 levels were markedly increased in DNP-treated cells. However, the reduction of intracellular H2O2 level by Tiron and catalase did not prevent apoptosis induced by DNP. Moreover, DNP rapidly reduced intracellular GSH content in As4.1 cells. Taken together, apoptosis in DNP-treated As4.1 cells is correlated with the rapid change of intracellular GSH levels rather than ROS levels.


Keywords: DNP, ROS, Calu-6, ROS scavenger, GSH, Mitochondria.

*Corresponding author. Tel.: +82 63 270 3079; fax: +82 63 274 9892.


1 Introduction

Reactive oxygen species (ROS) include hydrogen peroxide (H2O2), superoxide anion (O2) and hydroxyl radical (HO). ROS have recently been implicated in the regulation of various important cellular events, including transcription factor activation, differentiation and cell proliferation (Gonzalez et al., 2002; Baran et al., 2004; Shen and Liu, 2006). ROS are formed as by-products of mitochondrial respiration or precise oxidases including nicotine adenine dephosphate (NADPH) oxidase, xanthine oxidase (XO) and certain arachidonic acid oxygenases (Zorov et al., 2006) An alteration on the redox state of the tissue implies a change in ROS generation or metabolism. Principal metabolic pathway involves superoxide dismutase (SOD), which is expressed as extracellular, intracellular and mitochondrial isoforms, and metabolized O2 to H2O2 (Wilcox, 2002). Cells possess antioxidant systems to control the redox state, which is important for their survival. Excessive production of ROS gives rise to activation of events, which lead to death and survival in several types of cells (Simon et al., 2000; Chen et al., 2006; Dasmahapatra et al., 2006; Wallach-Dayan et al., 2006). The precise mechanisms involved in cell death induced by ROS remain an open question and the protective effect of some antioxidants on cell death is still controversial.

Mitochondria play a pivotal role in the control of most physiological processes, cell injury and programmed cell death (Cheng et al., 2006; Lakhani et al., 2006). The mitochondria of healthy cells maintain an electrochemical gradient across the mitochondrial inner membrane. The proton gradient and membrane potential are the proton motive force that is used to drive ATP synthesis. Coupling of electron transport through the respiratory chain and ATP generation can be disrupted by uncoupling agents such as cyanide m-chloropheylhydrazone, caybonyl cyanide p-(trifluoromethoxy) phenylhydroazone, and 2,4-dinitrophenol (Kuruvilla et al., 2003; Futakawa et al., 2006). Mitochondria are also one of the most important sources of ROS production at mitochondrial respiratory chain level (Nohl et al., 2005).

2,4-Dinitrophenol (DNP) is a cellular metabolic poison (Fig. 1A) which is known to uncouple electron transport and oxidative phosphorylation by carrying protons across the mitochondrial membrane (Hanstein, 1976; McFee et al., 2004). DNP stimulates respiration, increases O2 consumption and causes the loss of the pH gradient across the inner mitochondrial membrane. Therefore, it has been demonstrated that DNP has a potential for autoxidation, increases the production of ROS and reduces the mitochondrial membrane potential (ΔΨm) (Dryer et al., 1980; Kadenbach, 2003; Futakawa et al., 2006). It was also reported that DNP greatly enhanced activities of mitochondrial MnSOD in isolated rat liver cells (Dryer et al., 1980). In view of apoptosis, the enhancing capacity of death receptor-induced apoptosis by DNP was seen in some cell lines (Jukat, CEM, SKW6 and HeLa) (Linsinger et al., 1999; Vier et al., 2004). Recent studies showed that DNP failed to induce apoptosis in human dopaminergic cells (Watabe and Nakaki, 2007) but induce apoptosis by decreasing ATP levels in human fibroblasts and human male germ cell (Erkkila et al., 2006; Miyoshi et al., 2006). However, its properties as an antitumor agent have not yet been fully understood.


Fig. 1

Effects of DNP on apoptosis in As4.1 cells. Exponentially growing cells were treated with the indicated concentration of DNP for 48h. (A) Structure of DNP. (B) Sub-G1 cells were measured with a FACStar flow cytometer. (C) Effects of DNP on cell plasma membrane. Cells with annexin V-staining were measured with a FACStar flow cytometer. Graph shows the percentages of annexin-positive cells. (D) Annexin-positive cells were measured at the designated hours. *P<0.05 compared with the control group.


Juxtaglomerular cell tumors (JGCTs; also known as reninomas), first described in the late 1960s (Robertson et al., 1967; Kihara et al., 1968), are rare benign tumors of the kidney. About 100 cases have been described to date. Reninomas are understood to arise from juxtaglomerular cells. Clinically, the patients suffer from headaches, polyuria, nocturia and dizziness including other symptoms. Hypertension is a sign in almost all patients and laboratory findings of hyperreninemia, hyperaldosteronism and hypokalemia are characteristic. Recently, a malignant JGCT was described (Duan et al., 2004).

As4.1 cells have been used as a model for the JG cell. This cell line was isolated from kidney neoplasm in a transgenic mouse containing a rennin SV40T-antigen transgene (Sigmund et al., 1990). However, the role of mitochondria in kidney cell death, especially juxtaglomerular (JG) cell, has not been evaluated. In the present study, we investigated the effect of DNP on apoptosis in As4.1 juxtaglomerular cells, evaluated the involvement of ROS, such as H2O2, O2, and GSH in the DNP-treated As4.1 cells.

2 Materials and methods

2.1 Cell culture

As4.1 cells (ATCC No. CRL-2193) are a rennin-expressing clonal cell line derived from the kidney neoplasm of a transgenic mouse (Sigmund et al., 1990). Cell cultures were maintained in humidified incubator containing 5% CO2 in air at 37°C. As4.1 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin–streptomycin (GIBCO BRL, Grand Island, N.Y.). Cells were routinely grown in 100-mm plastic tissue culture dishes (Nunc, Roskilde, Denmark) and harvested with a solution of trypsin-EDTA while in a logarithmic phase of growth. Cells were maintained in these culture conditions for all experiments.

2.2 Reagents

Dulbecco's modified Eagle's medium (DMEM), fetal bovine serum (FBS), dimethylsulfoxide (DMSO), ribonuclease (RNAse), propidium iodide, and 2,4-dinitrophenol (DNP) were purchased from Sigma–Aldrich Chemical Company (St. Louis. MO, USA). DNP was dissolved in ethanol at 2.5×10−1M as a stock solution. The cell-permeable O2 scavengers, 4-hydroxy-TEMPO (4-hydroxyl-2,2,6,6-tetramethylpierydine-1-oxyl) (TEMPOL), the vitamin E analog (4,-dihydroxyl-1,3-benzededisulfonic acid) (Tiron), the nontoxic dietary glutathione precursor, (N-acetylcysteine) (NAC), dithiothreitol (DTT), and (1-[2,3,4-trimethoxibenzyl]-piperazine) (Trimetazidine) were obtained from Sigma. These were dissolved in designated solution buffer at 1×10−1M as a stock solution. All of the stock solutions were kept at 4 or −20°C. Unless indicated otherwise, reagents were purchased from Sigma.

2.3 Sub-G1 analysis

Sub-G1 distributions were determined by staining of DNA with PI as described previously (Han et al., 2007a). In brief, cells were incubated with the designated doses of DNP with or without ROS scavenger, SOD or catalase for 48h. Cells were then harvested, washed with PBS, fixed 70% ethanol, and stored at 4°C. Cells were washed again with PBS and were then incubated with PI (10μg/ml) with simultaneous treatment of RNase at 37°C for 30min. The number of cells in different phases of the cell cycle or having sub-G1 DNA content was measured with a FACStar flow cytometer (Becton Dickinson, San Jose, CA).

2.4 Annexin V/PI staining

Apoptosis was determined by staining cells with annexin V-fluorescein isothiocyanate (FITC) and PI labeling as described previously (Han et al., 2007b). PI can also be used to differentiate necrotic, apoptotic, and normal cells. This agent is membrane-impermeant and is generally excluded from viable cells. In brief, cells were incubated with the designated doses of DNP with or without ROS scavengers, SOD or catalase for 48h. The cells were washed with cold PBS, and then resuspended in 500μl of binding buffer (10mM HEPES/NaOH pH 7.4, 140mM NaCl, 2.5mM CaCl2) at a concentration of 1×106cells/ml. Five microliters of annexin V-FITC (PharMingen, San Diego, CA) and PI (1μg/ml) were then added to these cells at 37°C for 30min, which were analyzed with a FACStar flow cytometer (Becton Dickinson).

2.5 Measurement of mitochondrial membrane potential (ΔΨm)

The mitochondrial membrane was monitored using the Rhodamine 123 fluorescent dye (from Sigma), a cell-permeable cationic dye that preferentially enters mitochondria based on a highly negative mitochondrial membrane potential (ΔΨm). Depolarization of mitochondrial membrane potential (ΔΨm) results in the loss of Rhodamine 123 from the mitochondria and a decrease in intracellular fluorescence. In brief, cells were incubated with the designated doses of DNP with or without ROS scavengers, SOD or catalase for 48h. Cells were washed twice with PBS and incubated with Rhodamine 123 (0.1μg/ml) at 37°C for 30min. Subsequently, PI (1μg/ml) was added and the intensity of fluorescence was then determined by flow cytometry.

2.6 Quantification of caspase-9 activity

The activity of caspase-9 was assessed using the caspase-9 Colorimetric Assay Kit (R&D Systems, Inc., Minneapolis, MN, USA), which is based on the spectrophotometric detection of the color reporter molecule p-nitroaniline (pNA) after cleavage from the labeled substrate LEHD-pNA (caspase-9) as an index. Briefly, cells were incubated with or without 2mM of DNP for 4 and 48h. The cells were then washed in PBS and suspended in 5 volumes of lysis buffer (20mM HEPES pH 7.9, 20% glycerol, 200mM KCl, 0.5mM EDTA, 0.5% NP40). The lysates were then collected and stored at −20°C until use. Protein concentration was determined by the Bradford methods. Supernatant samples containing 100μg of total protein were used for determination of caspase activity. These were added to each well in 96-well microliter plates (Nunc, Roskilde, Denmark) with each substrate at 37°C for 1–2h. The optical density of each well was measured at 405nm using a microplate reader (Spectra MAX 340, Molecular Devices Co., Sunnyvale, CA, USA). Each plate contained multiple wells of a given experimental condition and multiple control wells. Caspase activity was expressed as fold changes of arbitrary absorbance units (absorbance at a wavelength of 405nm).

2.7 Western blot analysis

Cells were incubated with the designated doses of DNP for 48h. The cells were then washed in PBS and suspended in 5 volumes of lysis buffer (20mM HEPES pH 7.9, 20% glycerol, 200mM KCl, 0.5mM EDTA, 0.5% NP40, 0.5mM DTT, 1% protease inhibitor cocktail (from Sigma)). The supernatant protein concentration was determined by the Bradford method. Supernatant samples containing 20μg of total protein were resolved by 8 or 15% SDS-PAGE gel depending on the target protein sizes, and were then transferred onto an Immobilon-P PVDF membrane (Millipore, Mass, USA) by electroblotting and probed with anti-p27 (BD Transduction Laboratories, Franklin Lakes, NJ, USA), anti-caspase-3 and anti-PARP (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The membranes were incubated with horseradish peroxidase-conjugated secondary antibodies. The blot was developed using an ECL kit (Amersham, Arlington Heights, IL).

2.8 Detection of intracellular H2O2 and O2 level

Intracellular H2O2 level was detected by means of an oxidation-sensitive fluorescent probe dye, 2',7'-dichlorodihydrofluorescein diacetate (H2DCFDA) (Invitrogen Molecular Probes, Eugene, OR). H2DCFDA was deacetylated intracellularly by nonspecific esterase, which was further oxidized by cellular peroxides, yielding 2,7-dichlorofluorescein (DCF), a fluorescent compound. Dihydroethidium (DHE) (Invitrogen Molecular Probes) is a fluorogenic probe that is highly selective for the detection of superoxide anion radicals. DHE is cell-permeable and reacts with superoxide anion to form ethidium which, in turn, intercalates in deoxyribonucleic acid, thereby exhibiting a red fluorescence. In brief, cells were incubated with the indicated does of DNP with or without ROS scavengers, SOD or catalase for 48h. Cells were then washed in PBS and incubated with 5μM DHE at 37°C for 30min according to the instructions of the manufacturer. Red fluorescence was detected using a FACStar cytometer (Becton Dickinson) (Bass et al., 1983). Ten thousand events were collected for each sample. H2O2 and O2 production were expressed as mean fluorescence intensity (MFI) which was calculated by CellQuest software.

2.9 Detection of the intracellular glutathione (GSH)

Cellular GSH levels were analyzed using 5-chloromethlfluorescein diacetate (CMFDA, Molecular Probes). CMFDA is a useful and membrane-permeable dye for determining levels of intracellular glutathione (Hedley and Chow, 1994; Macho et al., 1997). In brief, cells were incubated with the indicated dose of DNP with or without ROS scavenger, SOD or catalase for 48h. Cells were then washed with PBS and incubated with 5μM CMFDA at 37°C for 30min according to the instructions of the manufacturer. Cytoplasmic esterases convert nonfluorescent CMFDA to fluorescent 5-chloromethylfluorescein, which can then react with the glutathione. CMF fluorescence intensity was determined using a FACStar flow cytometer (Becton Dickinson) and calculated by CellQuest software. Ten thousand events were collected for each sample.

2.10 Statistical analysis

Results represent the mean of three independent experiments; bar, SD. Student's t-test or one-way analysis of variance (ANOVA) with post hoc analysis using Tukey's multiple comparison test was used for parametric data. P value (<0.05) was considered to be statistically significant.

3 Results

3.1 Induction of apoptosis by DNP in As4.1 cells

We performed an in vitro apoptosis detection assay to determine whether DNP-treatment could induce apoptosis in As4.1 cells. Interestingly, the percent of As4.1 cells having the sub-G1 cells was elevated to &007E;40% at 48h following the treatment of 2mM DNP, and these were dose-dependent (Fig. 1B). To characterize the cell death induced by DNP, we examined the nuclear morphologies of dying cells using the fluorescent DNA-binding agent, DAPI. As4.1 cells treated with DNP displayed typical morphological features of apoptosis cells, i.e. condensed nuclei (data not shown). To further confirm and evaluate the induction of apoptosis, we stained cells with annexin V and PI. As with the percentages found in the sub-G1 cells by flow cytometry, the proportion of annexin V-staining cells in the DNP-treated cells was increased in a dose- and time-dependent manner (Fig. 1C and D).

3.2 Effects of DNP on mitochondrial membrane potential (ΔΨm) and apoptotic-related proteins in As4.1 cells

To evaluate the effect of DNP on mitochondrial membrane potential (ΔΨm), changes in mitochondrial membrane potential (ΔΨm) was monitored using Rhodamine 123. Cells were treated with indicated doses of DNP for 48h or treated with 2mM of DNP for designated times. The number of cells stained negatively for Rhodamine 123 showed a similar pattern to those of the cells staining positive for annexin V (Fig. 2A and B). Since DNP is a mitochondrial damaging agent, we tested the activation of caspase-9 after treatment with DNP. Caspase-9 was activated by DNP at 4h (Fig. 2C). However, caspase-9 inhibitor did not block the cell death induced by DNP (data not shown).


Fig. 2

Effects of DNP on mitochondrial membrane potential (ΔΨm) and apoptosis-related proteins in As4.1 cells. (A) Mitochondrial membrane potential (ΔΨm). Cells were treated with the indicated concentration of DNP for 48h, stained with Rhodamine 123, and analyzed by flow cytometry. Graph shows the percentages of Rhodamine 123-negative cells. (B) Rhodamine 123-negative cells were measured at the indicated hours. (C) Graph shows the activity of caspase-9. (D) Exponentially growing cells were treated with the indicated concentration of DNP for 48h. Aliquots of 20μg of protein extracts were resolved by 8 or 15% SDS-PAGE gel, transferred onto the PVDF membrane, and immunoblotted with the indicated antibodies, p53, Bcl-2, PARP, ß-tubulin. *P<0.05 compared with the control group.


Concerning the apoptotic-related proteins during DNP-induced apoptosis, anti-apoptotic protein, Bcl-2, was decreased (Fig. 2D) and pro-apoptotic protein, Bax, was not changed (data not shown). In addition, the Bcl-2 is transcriptional targets for the tumor suppressor protein, p53, which induces cell cycle arrest or apoptosis in response to DNA damage (Coutts and La Thangue, 2006). As shown in Fig. 2D, the expression of p53 was increased in As4.1 cells treated with DNP. In regard to poly (ADP-ribose) polymerase (PARP) protein which is a hall mark of apoptosis, the result showed that the intact 116kDa moiety of PARP was degraded in DNP-treated As4.1 cells (Fig. 2D). These results indicate conclusively that DNP-induced apoptosis in As4.1 cells was accompanied by Bcl-2 decrease, loss of mitochondrial membrane potential membrane (ΔΨm), and PARP protein degradation.

3.3 Effect of DNP on ROS and GSH production in As4.1 cells

To assess the production of intracellular H2O2 in DNP-treated As4.1 cells, we used H2DCFDA fluorescence dye. An intracellular H2O2 level was significantly increased in a dose-dependent manner at 48h (Fig. 3A and B). When we treated As4.1 cells with several doses of DNP during the short time, the decreased pattern of H2O2 levels by this drug was clearly detected in less than 2h (1mM and 2mM of DNP) (Fig. 3B). And 3mM of DNP during the short time incubation increased the intracellular H2O2 levels. Since the increasing DCF fluorescence in the H2O2-treated As4.1 cells under our experimental condition was obvious (data not shown), our method of detecting the level of H2O2 in DNP-treated As4.1 cells using the H2DCFDA fluorescence was considered appropriate.


Fig. 3

Effects of DNP on ROS production in As4.1 cells. Exponentially growing cells were treated with the indicated doses of DNP for 48h. (A, B) Intracellular H2O2 levels. (C, D) Intracellular O2 levels. The graphs show the fold changes of fluorescence. *P<0.05 compared with the control group. (B, D) The graph shows the fold changes of fluorescence for the designated concentrations of DNP and times. ●, control; ○, DNP 500μM; ▼, DNP 1mM; ▽, DNP 2mM; ■, DNP 3mM; □, DNP 4mM.


We subsequently checked the change of intracellular O2 in DNP-treated As4.1 cells. Red fluorescence derived from DHE, reflecting O2 accumulation, was significantly increased in DNP-treated As4.1 cells. In particular, As4.1 cells treated with 3mM of DNP showed a more than fourfold increase in DHE fluorescence (Fig. 3C). Accumulation of O2 was observed on the time of 40min at the exposure of DNP (Fig. 3D). Maximum O2 in As4.1 cells treated with designated doses of DNP occurred at about 80min.

Cellular GSH is crucial for regulation of cell proliferation, cell cycle progression and apoptosis (Poot et al., 1995; Schnelldorfer et al., 2000). We therefore analyzed the changes of GSH level of As4.1 cells by using CMF fluorescence. The M1 population of As4.1 cells showed a lower level of intracellular GSH content. DNP significantly elevated the percentage of cells residing in the M1 population (GSH depleted cells) at 48h in a dose-dependent manner (Fig. 4A and B), indicating the depletion of intracellular GSH content of As4.1 cells by DNP. The noteworthy depletion changes of intracellular GSH content was observed at the about 1–4mM concentration of DNP. The decrease of CMF fluorescence was observed within the early time of 20min at the exposure to DNP (2–4mM) (Fig. 4C).


Fig. 4

Effects of DNP on GSH production in As4.1 cells. (A) Exponentially growing cells were treated with the indicated DNP for 48h. Intracellular GSH level determined by FACStar flow cytometer as described in Section 2. (B) The graph shows the levels of mean of CMF fluorescence of (A). *P<0.05 compared with the control group. (C) The graph shows the mean CMF fluorescence for the designated concentration of DNP and times. ●, control; ○, DNP 500μM; ▼, DNP 1mM; ▽, DNP 2mM; ■, DNP 3mM; □, DNP 4mM.


3.4 Effects of ROS scavengers, SOD and catalase on ROS production, GSH depletion, and apoptosis in DNP-treated As4.1 cells

To determine whether ROS production and GSH depletion in DNP-treated As4.1 cells were changed by ROS scavengers, cells were pretreated with cell-permeable ROS scavengers, Tempol or Tiron, or a well-known antioxidant, NAC (Zafarullah et al., 2003) for 30min, and then treated with 2mM of DNP. An anti-ischemic and metabolic agent, Trimetazidine was also used as an indirect antioxidant (Tikhaze et al., 2000; Stanley and Marzilli, 2003). To measure the accurate intracellular fluorescence level of ROS, forward light scatter (FSC) characteristics were used to exclude cell debris from the analysis. In contrast to our expectation, the scavengers did not reduce the increased O2 level by 2mM DNP. The scavenger, Tiron, significantly decreased the H2O2 levels in DNP-treated cells. In addition, none of the scavengers prevent the depletion of GSH content in DNP-treated As4.1 cells (Fig. 5).


Fig. 5

Effects of ROS scavengers, SOD, and catalase on intracellular ROS and GSH production in DNP-treated As4.1 cells. Exponentially growing cells were treated with the ROS scavengers, SOD and catalase in addition to 2mM of DNP for 8h. (A) Graph shows the fold changes of DCF fluorescence. (B) Graph shows the fold changes of DHE fluorescence. (C) Graph shows the mean CMF fluorescence. *P<0.05 compared with the control group. **P<0.05 compared with the only DNP-treated cells.


To see whether ROS production and GSH depletion in DNP-treated As4.1 cells were changed by SOD and catalase, As4.1 cells were treated with DNP in the presence or absence of SOD (30unit/ml) or catalase (30unit/ml) for 48h. Catalase significantly decreased the level of H2O2 in DNP-treated As4.1 cells and SOD did not increase the intracellular H2O2 (Fig. 5A). In regard to the O2 levels by SOD and catalase, SOD and catalase slightly decreased the O2 levels in DNP-treated cells (Fig. 5B). Also, SOD and catalase did not significantly inhibit the depletion of GSH content in As4.1 cells treated with DNP (2mM) (Fig. 5C).

We also examined whether ROS scavengers prevent DNP-induced As4.1 cell death. None of the scavengers significantly decreased the sub-G1 cell population in DNP-treated As4.1 cells (Fig. 6A). In view of annexin V positive staining, only NAC slightly decreased the As4.1 cells having annexin V positive stain by DNP (Fig. 6B). However, none of the scavengers inhibit the loss of mitochondrial membrane potential (ΔΨm) (Fig. 6C). Additionally, high concentration (1mM) of these ROS scavengers could not significantly change apoptosis parameters (data not shown).


Fig. 6

Effects of ROS scavengers, SOD, and catalase on DNP-induced apoptosis. Exponentially growing cells were treated with the ROS scavengers, SOD, and catalase in addition to 2mM of DNP. Sub-G1 cells, annexin-positive cells and Rhodamine 123-negative cells were measured by flow cytometry. Graphs show the percentage of sub-G1 population (A), annexin-positive cells (B), and Rhodamine 123-negative cells (C). *P<0.05 compared with the control group.


Following tests to see whether SOD and catalase could prevent DNP-induced As4.1 cell death, we found they did not alter the sub-G1 cells and annexin V positive stained cells induced by DNP (Fig. 6A and B). Interestingly, catalase efficiently reduced the loss of mitochondrial membrane potential (ΔΨm) in DNP-treated As4.1 cells (Fig. 6C).

4 Discussion

We focused here on the apoptotic role of DNP, and on the involvement of ROS such as H2O2, O2 and GSH in DNP-induced As4.1 cell death. In addition, we also investigated whether ROS scavengers could rescue As4.1 cell death and its mechanisms. The results show that DNP potently induces apoptosis in a dose- and time-dependent manner in As4.1 juxtaglomerular cells. This observation was verified by morphological change of nucleus (data not shown), the increase of sub-G1 DNA content, phosphatidylserine externalization, and PARP protein cleavage. To gain insight into the molecular mechanism involved in apoptosis caused by DNP, expression of apoptotsis-related proteins and changes in mitochondrial membrane potential (ΔΨm) were assessed in As4.1 cells. We predicted that Bax to Bcl-2 ratio would be increased, since many apoptotic agents increase Bax protein and/or decrease Bcl-2 protein during apoptotic process in their target cells. Similarly, we showed that induction of apoptosis was accompanied by an elevation in Bax to Bcl-2 ratio. p53 regulates Bax and Bcl-2 protein expression (Coutts and La Thangue, 2006). In our experiment, the expression of p53 was increased in DNP-treated As4.1 cells. It is conceivable that the p53 induction by DNP upregulates the ratio of Bax to Bcl-2, resulting in triggering of apoptosis. These results imply that apoptosis by DNP is a p53-dependent manner in As4.1 cells. Notably, there was similar pattern between annexin V positive staining cells and Rhodamine 123-negative staining cells. But the levels of Rhodamine 123-negative staining cells by DNP were higher than those of annexin V positive staining cells. Because uncoupling of oxidative phosphorylation carry protons across the inner mitochondrial membrane (Linsinger et al., 1999), these results support idea that DNP primarily damages the mitochondria of target cells, and then progression to next step of apoptosis such as phosphatidylserine exposure is more efficient.

The mechanism of apoptosis involves mainly two signaling pathways, including mitochondrial pathway and cell death receptor pathway (Ashkenazi and Dixit, 1998; Budihardjo et al., 1999; Shi, 2002). The key element in the mitochondrial pathway is the efflux of cytochrome c from mitochondria to cytosol, where it subsequently forms a complex (apoptosome) with Apaf-1 and caspase-9, leading to the activation of the caspase-3 (Mehmet, 2000). The cell death receptor pathway is characterized by binding cell death ligands and cell death receptors, and subsequently activates caspase-8 and caspase-3 (Hengartner, 2000; Liu et al., 2004). To determine which pathway is required for the induction of apoptosis by DNP, we examined the activity of caspase-9 in early time point (4h). We also incubated DNP-treated As4.1 cells with caspase inhibitors which are pan-caspase, caspase-3, caspase-8, and caspase-9 inhibitor. Interestingly, the activity of caspase-9 was significantly increased in DNP-treated As4.1 cells (Fig. 2C) and caspase-8 activity was not (data not shown). This result indicates that DNP induces apoptosis via the mitochondrial pathway. However, DNP-induced apoptosis did not attenuate in the presence of these inhibitors in view of sub-G1 cells and annexin V positive staining (data not shown). These results suggest that the induction of apoptosis by DNP did not require caspases. The modes of caspase activation during apoptosis by DNP may be dependent on cell types, because these caspase inhibitors were effective to rescue the DNP-induced apoptosis in Calu-6 lung cancer cells (Han et al., 2008).

Our data show that the intracellular H2O2 level was increased, depending on the concentration and incubation time of DNP in As4.1 cells. One interpretation is that the activity of SOD was decreased in DNP-treated cells, resulting in slow conversion from O2 to H2O2, reduction of the H2O2 level, and accumulation of O2. This is supported by the fact that the activity of SOD was significantly decreased in DNP-treated cells (data not shown). In addition, as we expected, the intracellular O2 level was significantly increased in a dose-dependent manner at 48h and incubation time of DNP. However, the level of O2 was decreased at an early time point of 20min in 0.5–4mM of DNP-treated As4.1 cells. In cells treated with 0.5–4mM of DNP, the production of O2 was augmented at &007E;80min. The level of O2 was then reduced after &007E;80min. The data suggest that the increase or decrease of O2 levels may result from the production of itself or/and changes of SOD activity by DNP, and be also time- and dose-dependent, which probably results in an alteration of the levels of intracellular H2O2 by affecting the activities of catalase or/and GSH peroxidase. Therefore, the exact mechanisms of cell death in relation to the levels of intracellular ROS in DNP-treated cells must be defined further.

We attempted to determine whether ROS scavengers prevent DNP-induced cell death throughout the reduction of the intracellular ROS level. In contrast to our expectation, none of the scavengers used in this experiment could reduce the level of O2 in As4.1 cells treated with 2mM of DNP. However, Tiron an ROS scavenger showing a reduction in the intracellular H2O2 level in DNP-treated As4.1 cells, did not affect the level of apoptosis. Despite using a high dose of Tiron (1mM) dramatically decreased the intracellular H2O2 level and O2 level in 2mM DNP-treated As4.1 cells, it did not prevent apoptosis induced by 2mM of DNP in view of sub-G1 cells and annexin V positive staining (data not shown). Also, high dose of other scavengers did not alter the intracellular ROS level and apoptosis parameters (data not shown). Moreover, Tiron and catalase did not potentiate the activity of catalase and reduce that of SOD attenuated by DNP (data not shown). These results suggest that the apoptotic effects of DNP are not comparable to the intracellular ROS levels. This notion could be supported by the finding that catalase and Tiron could not inhibit apoptosis in DNP-treated As4.1 cells with significant changes of intracellular H2O2 level. However, we cannot rule out the possibility that the increased ROS levels following treatment with DNP trigger apoptosis in As4.1 cells at early time points. Interestingly, catalase could partially prevent the loss of mitochondrial membrane potential (ΔΨm). Increased H2O2 induces collapse of the mitochondrial membrane potential (ΔΨm) (Arita et al., 2006). This suggest that the intracellular H2O2-induced by DNP plays a role in loss of mitochondrial membrane potential (ΔΨm) rather than apoptosis resulting from entire cell damage. However, Tiron significantly decreased the production of intracellular H2O2 induced by DNP but could not prevent the loss of mitochondrial membrane potential in our study. Tiron is known to be a widely used antioxidant to rescue ROS-evoked cell death and Tiron mediated-cell death results from disturbance of iron metabolism (Kim et al., 2006). Tiron might impair the mitochondrial membrane potential (ΔΨm) by chelating the iron in cytochrome c independent of H2O2 level.

With regard to intracellular GSH, a main nonprotein antioxidant in the cell, it is able to clear away the superoxide anion free radical and provide electrons for enzymes such as glutathione peroxidase, which reduce H2O2 to H2O (Rhee et al., 2005). Intracellular GSH content has a decisive effect on anticancer drug-induced apoptosis, indicating that apoptotic effects are inversely proportional to GSH content (Higuchi, 2004; Estrela et al., 2006). Likewise, our result indicated that DNP significantly depleted intracellular GSH at 48h. And the time course studies revealed that the intracellular GSH depletion occurred prior to the onset of DNP-induced apoptosis. These results support the notion that the rapid depletion of intracellular GSH might play a critical role in DNP-induced apoptosis and intracellular GSH levels are closely related to DNP-induced cell death. This is supported by the result that GSH synthesis inhibitor (BSO) and GSH reductase inhibitor (Carmustine) intensified DNP-induced cell death (data not shown). Interestingly, NAC, the precursor of GSH (Zafarullah et al., 2003), did not affect ROS production and DNP-induced apoptosis. Probably, this is that DNP prevented the GSH synthesis from NAC in As4.1 cells or the uptake of NAC since we have detected that NAC significantly rescued uncoupling agent (carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP))-induced As4.1 cell death (unpublished data).

It has been considered that the kidney and juxtaglomerular apparatus (JGA) contain an ROS generating system responsive to angiotensin II (Wilcox, 2002, 2003). The ROS in the JGA-related cells is related to regulation of blood pressure (Wilcox, 2002, 2003). However, the role of ROS in kidney cell death, especially JG cells, has not been evaluated for apoptosis. Therefore, understanding the molecular mechanism of kidney cell death by an ROS generator such as DNP remains an important task.

In summary, DNP generated ROS and induced the depletion of GSH content in As4.1 cells. Treatment with Tiron and catalase to reduce increased intracellular H2O2 level did not attenuate DNP-induced apoptosis in As4.1 cells, and catalase inhibits the loss of mitochondrial membrane potential (ΔΨm). Taken together, apoptosis in DNP-treated As4.1 cells is correlated with the rapid change of intracellular GSH levels rather than ROS levels.

Acknowledgements

This research was supported by the Korean Science and Engineering Foundation (R01-2006-000-10544-0) and a Korea Research Foundation Grant funded by the Government of the Republic of Korea (MOEHRD).

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Received 20 February 2008/10 July 2008; accepted 19 August 2008

doi:10.1016/j.cellbi.2008.08.023


ISSN Print: 1065-6995
ISSN Electronic: 1095-8355
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