This article is mentioned in:

Introduction

para-Phenylenediamine (p-PD) is used in hair dye formulations and it is estimated that two-thirds of the hair dye formulations that are marketed contain p-PD (1). Various azo dyes used by the industry also contain p-PD. Upon azo reduction of these compounds by environmental or intestinal microorganisms, p-PD is released. When p-PD is ingested, it is absorbed and redistributed to target sites to exert its effects (2,3). Epidemiological studies have indicated that increased usage of permanent hair dye may increase the risk of bladder cancer, non-Hodgkin’s lymphoma, multiple myeloma, and haematopoietic cancer (4,5). Besides personal usage, professional hairdressers and dye industry workers are frequently exposed to dyes containing p-PD, and epidemiological studies have demonstrated that those who are frequently exposed to this chemical compound have incurred a higher risk of various cancers (1,4). In addition, Sontag (6) demonstrated that the incidence of kidney tumours increased with exposure to p-PD in rats. Thus, in the current study, the mechanism by which p-PD induces apoptosis in normal rat kidney proximal tubular epithelial (NRK52E) cells was investigated. This cell line is a commonly used cell line for in vitro evaluation of apoptotic pathways. For example, it was demonstrated in NRK52E cells that Numb protects against puromycin aminonucleoside-induced apoptosis by inhibiting the Notch signalling pathway (7). In addition, urografin was demonstrated to induce apoptosis in NRK52E cells via upregulated glucose-regulated protein 78 (GRP78) and GRP94 expression, procaspase-12 cleavage and phosphorylation of protein kinase-like endoplasmic reticulum kinase and eukaryotic initiation factor 2α (8).

Focal adhesion kinase (FAK) phosphorylation is the pivotal integrin-mediated signalling event, since this cytoplasmic tyrosine kinase acts as the scaffold for several effector molecules, such as phosphoinositide 3-kinase (PI3K)/protein kinase B (Akt) and the Ras/Raf/c-Jun N-terminal kinase (JNK) cascades (9,10). FAK activation has been associated with survival signals through the activation of the PI3K/Akt and Ras/Raf/JNK pathways (11,12,13). PI3K/Akt is intimately involved in cell survival, as it regulates the activity of several Bcl-2 family members (14,15,16). Protein tyrosine kinases (PTKs) are activated by and form complexes with growth factor receptor-bound protein 2 and Son of sevenless, resulting in activation of Ras, and subsequent activation of the Raf and JNK cascade survival pathways (17,18). For example, evodiamine-induced oxidative stress and cell cycle arrest was demonstrated to act through the PTK/Ras/Raf/JNK pathway in HeLa human cervical carcinoma cells (19), and oridonin has been indicated to induce G2/M phase cell cycle arrest and apoptosis via inhibition of the PTK/Ras/Raf/JNK survival pathway in L929 murine fibrosarcoma cells (20).

Previous studies have demonstrated that p-PD induces apoptosis via p53 in addition to intrinsic and extrinsic pathways in MDCK cells (21,22). Huang et al (23) also demonstrated that p-PD induces DNA damage and the expression of mutant p53 and COX-2 proteins in SV-40 immortalized human uroepithelial cells. In the present study, the roles of the PTK/Ras/Raf/JNK and PI3K/Akt signalling pathways on p-PD-treated NRK-52E cells was investigated in relation to cell death.

Materials and methods

Materials

Dulbecco’s modified Eagle’s medium (DMEM), foetal bovine serum (FBS) and trypsin-EDTA were purchased from Gibco Life Technologies (Grand Island, NY, USA). p-PD, dimethyl sulfoxide (DMSO), Triton X-100, Tergitol NP-40, EDTA, Tris-HCl, trypan blue, phosphate-buffered saline (PBS), goat anti-rabbit Immunoglobulin G horseradish peroxidase (HRP)-conjugated polyclonal secondary antibodies, dithiothreitol (DTT), sodium dodecyl sulphate (SDS), ammonium acetate, Tris-borate-EDTA buffer, Bradford reagent and phenylmethyl sulfonyl fluoride were purchased from Sigma-Aldrich (St. Louis, MO, USA). The Annexin-V-FLUOS Staining kit was purchased from Roche Diagnostics GmbH (Mannheim, Germany). Proteinase K, ribonuclease A (RNase A) were obtained from BD Pharmingen (San Diego, CA, USA). Rabbit anti-rat monoclonal antibodies for Ras, SAPK-JNK, Akt, Bcl-2, Bcl-xL, Bad, tubulin and phospho SAPK-JNK and mouse anti-rat c-Raf were perchased from Cell Signaling Technology, Inc. (Danvers, MA, USA). Amersham ECL-Plus Western Blotting Reagents and polyvinylidine fluoride (PVDF) membranes were obtained from GE Healthcare Bio-Sciences (Pittsburgh, PA, USA). All of the chemicals were of the highest grade commercially available.

Cell culture and treatment

The NRK-52E normal rat renal tubular epithelial cell line was obtained from the American Type Culture Collection (Manassas, VA, USA). The cells were maintained as a monolayer in DMEM with 2.0 mM L-glutamine adjusted to contain 3.7 g/l sodium bicarbonate and 4.5 g/l glucose. The medium was supplemented with 1% penicillin (100 U/ml; Sigma-Aldrich), streptomycin (10,000 μg/ml; Sigma-Aldrich) and 10% FBS. Cells were cultured in 25-cm2 tissue culture-treated flasks at 37°C and 5% CO2 in humidified chambers.

The stock solution of p-PD (100 mg/ml) was dissolved in DMSO and different concentrations were prepared in the culture medium with a final DMSO concentration of 0.1%.

Cell viability assay

The NRK-52E cells were treated with various concentrations of p-PD (50, 100, 200 and 300 μg/ml), or 0.1% DMSO for the control, for 24 h at 37°C. A trypan blue exclusion protocol was used to determine the cell viability. Briefly, ~10 μl cell suspension in PBS was mixed with 40 μl trypan blue, and the numbers of stained (dead cells) and unstained cells (live cells) were examined under a Nikon Eclipse TS-100F inverted microscope (Nikon Corp., Tokyo, Japan) (24).

Cell cycle analysis

The NRK-52E cells were cultured in 25-cm2 culture flasks and treated with different concentrations of p-PD (50, 100, 200 and 300 μg/ml) for 24 h. Subsequent to exposure, the cells were collected, washed with PBS and fixed with ice-cold 70% ethanol overnight at 4°C. The cells were washed with PBS, stained with 1 ml fluorochrome solution from the Annexin-V-FLUOS Staining kit [containing 20 μg/ml propidium iodide (PI) and 10 μg/ml RNase A] for 15 min in dark conditions and analysed using a BD FACSCalibur flow cytometer (E97500679; BD Biosciences, Franklin Lakes, NJ, USA).

Annexin-V staining

The NRK52E cells were cultured in 60-mm tissue-culture dishes. The culture medium was replaced with fresh medium as cells reached 70% confluence, then different concentrations of p-PD (50, 100, 200 and 300 μg/ml) were added prior to 24-h culture. Levels of apoptosis were determined by staining with the Annexin-V kit (25). Following incubation, floating or adherent cells that were later trypsinised were pooled and centrifuged for 5 min at 1,000 × g. Pelleted cells were washed with PBS. Next, cells were centrifuged for 5 min at 1,000 × g and resuspended in 100 μl Annexin-V-Fluos and PI labelling solution (from the Annexin-V kit) for 10 min. The stained cells were analysed by flow cytometry, where the fluorescence emission was measured at 530 nm. The percentage cell apoptosis was calculated using BD Multiset™ 2.2, BD FACStation™ 5.2.1, ModFit LT 3.0 and CellQuest software (BD Biosciences).

Detection of intracellular reactive oxygen species (ROS)

The NRK52E cells were treated with 100 μg/ml p-PD for 1, 2 or 3 h, and controls were treated with 0.1% DMSO. All cells were stained with 10 μM 2′,7′-dichlorofluorescin diacetate (DCFH-DA; Sigma-Aldrich) for 30 min. Subsequent to washing with PBS, the fluorescence intensity was detected by a Tecan Infinite 200 PRO fluorescence plate reader (Tecan Group Ltd., Maennedorf, Switzerland) with excitation and emission wavelengths of 488 and 525 nm, respectively.

Western blot analysis

Western blot analysis was performed according to the methods of a previous study (26). The culture medium was replaced with fresh medium as cells reached 70% confluence, then different concentrations of p-PD (50, 100, 200 and 300 μg/ml) were added and cells were cultured for 24 h. Next, adherent and floating cells were collected and homogenised in a lysis buffer (10 mM Tris-HCl, pH 8.0; 0.32 mM sucrose; 5 mM EDTA; 2 mM DTT; 1 mM phenylmethyl sulfonyl fluoride; and 1% Triton X-100) and centrifuged at 10,621 × g (5427R; Eppendorf, Hamburg, Germany) for 10 min. The supernatants were collected and assayed for protein concentration using the Bradford protein assay method (27). An equal quantity of protein per sample was subjected to 10% SDS-polyacrylamide gel electrophoresis. Following electrophoresis, the proteins were transferred to the PVDF membranes by electroblotting and incubated with diluted primary antibodies for 1 h at 25°C. The membranes were washed, incubated for 30 min at 25°C with the HRP-conjugated secondary antibodies and subsequently washed extensively prior to detection by chemiluminescence with the ECL-Plus kit. The proteins were visualised by exposing the blots to film (Kodak, Rochester, NY, USA). The western blot data were quantified using Image J software (http://imagej.nih.gov/ij/).

Statistical analysis

Results are expressed as the mean ± standard deviation from at least three independent experiments. Statistical analysis was performed using Student’s t-test. *P<0.05 was considered to indicate a statistically significant difference. The error bars denote standard deviation.

Results

p-PD alters cell morphology

The NRK-52E cells were treated with four different concentrations of p-PD (50, 100, 200 and 300 μg/ml) for 24 h, then observed under an inverted microscope. The control cells retained their normal, clear plasma membrane. There was a uniform cell distribution with neighbouring cells closely connected to each other and clear cell nuclei (Fig. 1A). Following p-PD treatment, the cells were enlarged, forming a variety of shapes and sizes. The cells lost contact with neighbouring cells and cytoplasmic vacuolisation occurred (Fig. 1B and C). In addition, cell shrinkage and blebbing of the plasma membrane were also observed at higher p-PD concentrations (Fig. 1D and E).

Figure 1 Effects of p-PD on cell morphology in NRK52E cells. NRK52E cells were treated with (A) DMSO or p-PD at a concentration of (B) 50, (C) 100, (D) 200 and (E) 300 μg/ml for 24 h. The results were observed under an inverted microscope; magnification, ×200. p-PD, para-phenylenediamine; DMSO, dimethyl sulfoxide.

p-PD reduces cell viability

The cell viability of p-PD-treated NRK-52E cells was examined with a trypan blue exclusion assay subsequent to a 24 h incubation period. The results demonstrated reduced cell viability in NRK-52E cells that were treated with p-PD; and as the concentration increased, cell viability reduced. Treatments of 50, 100, 200 and 300 μg/ml led to cell viabilities of 62.2, 52.5, 36.8 and 25.4% of the control (Fig. 2), respectively.

Figure 2 Effect of p-PD on cell viability in NRK52E cells, evaluated by a trypan blue exclusion assay. NRK52E cells were treated with different doses of p-PD (50, 100, 200 and 300 μg/ml) for 24 h. Data are presented as the mean ± standard deviation and are representative of three independent experiments. *P<0.05 vs. the control group. p-PD, para-phenylenediamine.

p-PD alters cell cycle progression and inhibits mitosis

p-PD was previously demonstrated to induce cell death. To determine the nature of the cell death (necrotic or apoptotic) changes in the DNA content of the p-PD-treated cells were detected using the PI staining method. The cell cycle results indicated that the cell cycle distribution of control cells was as follows: 0.64±0.01% in the sub-G1 phase; 53.04±1.52% in the G1 phase; 13.29±0.65% in the S phase; and 33.41±2.45% in the G2+M phase.

In cells exposed to 50 μg/ml p-PD, the percentage of cells in the sub-G1 phase increased to 20.99±0.89%, compared with 34.02±0.45% following exposure to 100 μg/ml p-PD. When the p-PD concentrations increased to 200 and 300 μg/ml, the percentages of cells in the sub-G1 phase further increased to 47.23±3.11 and 76.74±2.45%, respectively (Fig. 3).

Figure 3 Effect of p-PD on cell cycle distributions in NRK52E cells. Cells were treated with different doses of p-PD (50, 100, 200 and 300 μg/ml) for 24 h. Following staining with PI, the cell cycle was analysed by flow cytometry. The percentages of cells in the sub G1, G1, S and G2+M phases following treatment with various doses of p-PD are presented as cumulative data from three independent experiments. p-PD, para-phenylenediamine; PI, propidium iodide.

p-PD also induced a reduction in the numbers of cells in the G2+M phase compared with the control cells. Compared with 33.41±2.45% in the control cells, the cells treated with p-PD at concentrations of 50, 100, 200 and 300 μg/ml presented 14.92±0.98, 20.76±1.02, 20.84±0.87 and 9.46±0.06% of cells in the G2+M phase, respectively. This result demonstrates a reduction in mitosis in the treated cells compared with the control cells (Fig. 3).

Annexin-V staining and flow cytometry

The induction of apoptosis by p-PD was further confirmed by Annexin-V staining. Following incubation with p-PD at a concentration of 50, 100, 200 or 300 μg/ml for 24 h, the percentages of Annexin V+/PI+ cells increased to 8.47±0.04, 15.05±0.08, 25.01±0.14 and 29.52±0.31%, respectively, compared with the control group (2.13±0.03%). Additionally, Annexin V+/PI− cells were also increased to 12.95±0.02, 16.10±0.05, 15.87±0.05 and 36.01±0.29%, respectively, compared with the control group (6.98±0.01%) (Fig. 4).

Figure 4 Effect of p-PD on apoptosis in NRK52E cells. Cells were treated with different doses of p-PD (50, 100, 200 and 300 μg/ml) for 24 h. Following staining with Annexin-V and PI, levels of apoptosis were analysed by flow cytometry. The cumulative data are from three independent experiments. p-PD, para-phenylenediamine; PI, propidium iodide; ; FITC, fluorescien isothiocyanate.

p-PD induces intracellular ROS generation

The effects of p-PD on the production of intracellular ROS were examined following DCFH-DA staining. The control cells exhibited low-level ROS generation (Fig. 5), and the cells treated with 100 μg/ml p-PD for 1 h presented a ROS level twice as high as controls. Following 2-h treatment, the ROS level had increased four-fold compared with the control cells. The intracellular ROS level increased markedly in the cells that were treated with 100 μg/ml p-PD for 3 h (Fig. 5).

Figure 5 Effect of p-PD on ROS generation in NRK52E cells. NRK52E cells were treated with 100 μg/ml p-PD for 1, 2 and 3 h. ROS generation was indicated by DCFH-DA fluorescence. The fluorescence intensity levels were detected using a fluorescence plate reader at excitation and emission wavelengths of 488 nm and 525 nm, respectively. The fluorescence intensity in cells treated with various doses of p-PD is presented as cumulative data from three independent experiments (P<0.05 vs. control). p-PD, para-phenylenediamine.

Effects of p-PD on the protein expression levels of Ras, Raf, SAPK-JNK, Akt, Bcl-2, Bcl-xL and Bad

To assess the molecular mechanism underlying p-PD-induced apoptosis, the expression levels of the survival proteins Ras, Raf, Bcl-2 and Bcl-XL were assessed at 24 h following p-PD treatments of 50, 100, 200 and 300 μg/ml. The results demonstrated that the expression levels of the Ras and Raf survival proteins were reduced by p-PD in a dose-dependent manner (Fig. 6). However, Akt, Bcl-2 and Bcl-xL protein expression levels were not markedly altered compared with controls.

Figure 6 Effects of p-PD on signalling pathways in NRK52E cells. NRK52E cells were treated with different doses of p-PD (50, 100, 200 and 300 μg/ml) for 24 h. The protein expression levels of Ras, Raf, SAPK-JNK, Akt, Bcl-2, Bcl-xL and Bad were evaluated by western blot analysis. p-PD, para-phenylenediamine; SAPK, stress-activated protein kinase; JNK, c-Jun N-terminal kinase Akt, protein kinase B.

With regards to the apoptotic proteins, SAPK-JNK expression level was not markedly altered, and the phosphorylated SAPK-JNK expression levels markedly increased as p-PD concentrations increased (Fig. 6). In addition, p-PD had no effect on the Bad expression level with p-PD treatment, compared with the control cells. Tubulin was used as a loading control.

Discussion

Carcinogenesis is a process resulting from genetic alterations leading to mutations of oncogenes or tumour suppressor genes that drive the progressive transformation of normal cells into malignant cells (28,29). At the molecular level, genetic mutations are able to alter translated proteins and thereby disrupt downstream signalling pathways that are essential for apoptosis, cell cycle and other cellular processes (30,31). Cadmium, a causative agent in various types of cancer, elevates intracellular free calcium ion ([Ca2+]i) levels, leading to neuronal apoptosis partly by activating mitogen-activated protein kinases (MAPK) and mammalian target of rapamycin pathways (32). Additionally, cadmium exposure leads to the induction of the ERK signalling pathway, which alters gene expression in osteoblasts, and apoptotic death in Saos-2 cells (33). Chronic exposure to arsenic can lead to the development of various types of cancer; it downregulates Akt and c-Fos protein expression, and induces apoptosis in glutathione-deficient cells (34).

p-PD is a potential carcinogen that is widely used in permanent hair dye (35,36), and it has been reported that incidences of kidney tumours increase in rats following exposure to p-PD (6). Therefore, in the present study, the molecular mechanism underlying p-PD-induced apoptosis was investigated in NRK-52E cells, and to the best of our knowledge, it was demonstrated for the first time that p-PD-induces cell death in a dose-dependent manner in NRK-52E cells (Fig. 2). It was confirmed that this cell death was due to apoptosis, as indicated in Fig. 3. Cell cycle analysis demonstrated a reduction in the number of cells in the G2+M phase in addition to an increase in the number of cells in the sub-G1 phase in the treated cells, when compared with the respective percentages in the control group. This finding indicates that p-PD induced cell cycle arrest. In addition, Annexin-V staining demonstrated that the number of apoptotic cells increased following p-PD exposure in a dose-dependent manner (Fig. 4). In previous studies, it has been established that oxidised p-PD induces the production of ROS, leading to an imbalance between production and the removal of ROS and overwhelming oxidative stress that eventually induces apoptosis (37,38).

ROS generated primarily by the mitochondria are highly reactive metabolites that are produced during normal cell metabolism (39). Curtin et al (40) reported that the increases in intracellular ROS levels may lead to apoptosis. The underlying mechanism may involve the direct interaction and destruction of cellular proteins, lipids and DNA, and/or indirect interference with normal cellular signalling pathways and gene regulation (41). Consistent with these findings, the present results demonstrated that intracellular ROS levels increased significantly in the p-PD-treated NRK-52E cells in a dose-dependent manner (Fig. 5). High levels of intracellular ROS cause disruption of the mitochondrial membrane potential, release of cytochrome c with subsequent activation of the caspase cascade and ultimately, programmed cell death (42,43). Additionally, intracellular ROS can catalytically inactivate protein tyrosine phosphatases through the oxidation of active-site cysteine residues, which negatively regulate receptor tyrosine kinase (RTK) activity and downstream signalling, and hence allow sustained PTK phosphorylation and activation (44).

PTKs serve a key role in the transmission of various signals from cell-surface receptors to the nucleus. PTKs can be divided into the transmembrane (T)RTKs and non-RTKs (45). Ras links RTKs and non-RTKs to downstream serine/threonine kinases, including the MAPKs (46). The activation of the Ras/Raf/MAPK pathway has been demonstrated to induce growth arrest in several cell types. Oridonin induces apoptosis in L929 cells through inhibition of the PTK/Ras/Raf/JNK pathway (20). In addition, PKT/Ras/Raf/JNK inhibition-derived ROS/NO production contributed to G2/M phase cell cycle arrest in evodiamine-treated human cervix carcinoma HeLa cells (19). Consistent with these findings, in the current study it was demonstrated that the Ras/Raf/JNK pathway is able to promote apoptosis by inducing p-PD in NRK52E cells. Additionally, anti-carcinogenic compounds, UV- and gamma-irradiation have previously been indicated to induce apoptosis via a JNK-dependent pathway (47–50).

Oxidative stress stimulates multiple intracellular signal transduction pathways such as Akt-Bad. Akt, which is downstream of PI3K, regulates mechanically driven and receptor-ligand signalling (51). Activation of the PI3K/Akt can lead to Bad phosphorylation at specific serine residues. Phosphorylated Bad binds 14-3-3ζ proteins in the cytosol that sequester and tag Bad for subsequent degradation (52). Alternatively, pro-apoptotic proteins can be retained in the cytosol by binding to anti-apoptotic proteins, such as Bcl-2 and Bcl-xL (53). An increase in Bcl-2 and Bcl-xL expression prevents cytochrome c release from the mitochondria, thereby inhibiting activation of caspases, such as caspase-9 and caspase-3, and preventing apoptosis (54,55). In the present study, it was demonstrated that there were no changes in the levels of Bcl-2, Bcl-xL and Bad proteins compared with controls (Fig. 7). These findings suggest that the molecular mechanism triggered by p-PD-induced cell death is independent of the PI3K/Akt/Bad pathway.

In conclusion, the results of the present study demonstrated that p-PD induced apoptosis in NRK52E cells; in addition, DCFH-DA staining confirmed that apoptosis was induced due to oxidative stress. Furthermore, the results indicated that p-PD induced apoptosis via the PTK-Ras-Raf-JNK pathway, which upregulated SAPK/JNK protein expression levels and downregulated Ras and Raf protein expression levels. However, p-PD was found to induce apoptosis independent of PI3K/Akt pathway, as Akt, Bcl-2, Bcl-XL and Bad protein expression levels were not significantly altered compared with the control. Future studies are required in order to further elucidate the role of p-PD in tumorigenesis.

Acknowledgements

The present study was supported by grants FRGS/2/2010/ST/IMU/03/1(SKK) from Jabatan Pengajian Tinggi Malaysia and BMS 102-2010 (10) from the International Medical University, Kuala Lumpur, Malaysia.

References