2019 Volume 44 Issue 6 Pages 415-424
Polyhexamethylene guanidine phosphate (PHMG-p) is an active ingredient of humidifier disinfectants and causes severe lung injury resulting in pulmonary fibrosis. Current evidence indicates that pulmonary fibrosis is initiated as a result of epithelial damage, which can lead to an inflammatory response and fibrotic cell infiltration; however, the toxic mechanism of PHMG-p on the epithelium is still unknown. In this study, the toxic response of PHMG-p on human lung epithelial cells was evaluated, and its mechanisms associated with reactive oxygen species (ROS), DNA damage, and its relationship with p53 activation were investigated. The toxic responses of epithelial cells were assessed by flow cytometry analysis and western blot analysis. The results revealed that PHMG-p induced G1/S arrest and apoptosis in A549 cells. Interestingly, p53 was activated by PHMG-p treatment and p53 knockdown suppressed PHMG-p-induced apoptosis and cell cycle arrest. PHMG-p promoted ROS generation and consequently increased the expression of DNA damage markers such as ATM and H2AX phosphorylation. The antioxidant N-acetylcysteine reduced the expression of phosphorylated ATM and H2AX, and the ATM inhibitor, caffeine, inhibited p53 activation. Taken together, our results demonstrate that PHMG-p triggered G1/S arrest and apoptosis through the ROS/ATM/p53 pathway in lung epithelial cells.
Polyhexamethylene guanidine phosphate (PHMG-p) was used as a major component of humidifier disinfectants to clean humidifiers and remove microbial growth in South Korea. However, epidemiological and toxicological studies have revealed that PMHG-p induces pulmonary inflammation and fibrosis (Park et al., 2014; Song et al., 2014). Exposure to PHMG-p via inhalation has been previously found to cause the release of cytokines, the infiltration of inflammatory cells, and the formation of fibroblast foci in rodents (Kim et al., 2016).
Chronic lung epithelial damage is considered a key event in pulmonary fibrogenesis. Repeated damage to lung epithelial cells can cause collapse of the epithelial basement, which leads to the infiltration of inflammatory and fibrotic cells, resulting in the accumulation of excessive extracellular matrix (Wuyts et al., 2013). Epithelial cell cycle arrest and apoptosis have been suggested as the initial events of fibrogenesis (Uhal, 2003; Uhal, 2008; Yang et al., 2010). Sisson et al. (2010) demonstrated that type II pneumocyte injury initiated the development of pulmonary fibrosis. In addition, DNA damage was observed in the lung epithelium of patients with idiopathic pulmonary fibrosis (Kuwano et al., 1996).
When these cells are injured, DNA damage checkpoints delay cell cycle progression to get opportunities for DNA repair before replication (Enoch and Norbury, 1995). The cell cycle is regulated by the complex of a regulatory subunit known as cyclin and cyclin-dependent kinases (CDKs). When cells become damaged, CDK2 complexes with cyclin E and cyclin A are down-regulated, and CDK4 complexes with cyclin D are inhibited, which leads to G1/S cell cycle arrest (Lim and Kaldis, 2013). DNA damage is recognized by proteins that contain both signaling and repair activity. An important player in this process is ataxia-telangiectasia mutated (ATM). Induction of ATM induces G1/S cell cycle arrest and apoptosis by stabilizing p53 (Maréchal and Zou, 2013; Lavin and Kozlov, 2007). DNA damage induces cell cycle arrest to repair the DNA damage; however, failure to repair DNA results in apoptosis by activating p53 signaling (Chen, 2016; O’Reilly, 2001). In p53-dependent apoptosis, pro-caspase 3 is cleaved, thereby inactivating PARP, which repairs DNA lesions, and eventually leads to apoptosis (Green and Reed, 1998).
In a previous study, the functional analysis of mRNA microarray found that PHMG-p was associated with cellular responses including cell cycle, apoptosis, p53 activation, and DNA damage (Shin et al., 2018). However, the relationship between PHMG-p and epithelial damage has not yet been elucidated. In this study, we aimed to elucidate the toxic mechanisms of PHMG-p on epithelial damage. By flow cytometry and western blotting, cell cycle arrest and apoptosis were observed in A549 cells exposed to PHMG-p. Moreover, reactive oxygen species (ROS) production, DNA damage, and p53 activation were investigated.
Human lung epithelial cells, A549 cells, were obtained from the Korean Cell Line Bank (Seoul, Korea) and cultured in RPMI supplemented with 5% fetal bovine serum (Biotechnics Research Inc., Lake Forest, CA, USA), penicillin (100 units/mL), and streptomycin (100 mg/mL) at 37°C in an atmosphere of 5% CO2/95% air under saturating humidity.
Cell viability was assessed using the WST-1 assay. PHMG-p was provided by the Korea Institute of Toxicology (Jeongeup, Korea). Cells were seeded into 96-well plates at a density of 5 × 104 cells/well, cultured for 24 hr, and treated with PHMG-p at concentrations of 0-4 μg/mL for 12, 24, and 36 hr. Referring to the manufacturer’s instructions, 10 μL WST-1 reagent (Roche Diagnostics, Montclair, NJ, USA) was added to each well and the plates were incubated in an atmosphere of 5% CO2 at 37°C for 1 hr. Cell viability was quantified by measuring the absorbance at 440 nm and 690 nm using a VERSAmax microplate reader (Molecular Devices, Sunnyvale, CA, USA).
A549 cells (3 × 104 cells/well) were plated into 96-well plates and treated with transforming growth factor (TGF)-β1 (5 ng/mL) or PHMG-p using Incucyte ZOOM (Essen Biosciences, Ann Arbor, MI, USA). Scratches were formed using a wound maker. After creating wounds of equal width, the cells were washed twice with media to remove debris. Wound images were automatically acquired and recorded for 36 hr every 4 hr using a 10 × phase-contrast objective.
A549 cells were seeded into a 100-mm dish (5 × 105 cells/dish), incubated for 24 hr at 37°C, and treated with PHMG-p at concentrations of 0-4 μg/mL for 36 hr. After exposure to PHMG-p, cells in the culture media and attached cells were harvested by collecting the media and trypsinization, followed by centrifugation at 2,000 rpm at 24°C for 5 min. After centrifugation, the cells were washed once with ice-cold phosphate-buffered saline (PBS) and then centrifuged at 2,000 rpm at 4°C for 5 min. For cell fixation, the cells were resuspended in 300 μL distilled water and 700 μL ice-cold 95% EtOH was added dropwise while vortexing the cells. Cells were incubated for at least 30 min on ice, followed by centrifugation at 12,000 rpm at 4°C for 5 min. After washing the cells with ice-cold PBS and centrifugation at 12,000 rpm at 4°C for 5 min, the cells were resuspended in PBS. To remove any RNA remaining in the cell suspension, 10 mg/mL RNase A stock solution (#R6513; Sigma-Aldrich, St. Louis, MO, USA) was added to the samples at a final concentration of 100 μg/mL and incubated for 10 min at 24°C. After incubation, 1 mg/mL propidium iodide (PI) solution (#P4170; Sigma-Aldrich) was added to the samples at a final concentration 10 μg/mL and incubated in the dark until analysis by flow cytometry. PI staining was analyzed by flow cytometry (Guava easyCyteTM System, Millipore, Hayward, CA, USA).
A549 cells were seeded in a 100-mm dish (50 × 104 cells/dish), incubated for 24 hr at 37°C, and treated with PHMG-p at concentrations of 0-4 μg/mL for 36 hr. After exposure, the cells in the culture media and the attached cells were harvested by collecting the media, trypsinization, followed by centrifugation at 900 × g at 24°C for 5 min. After centrifugation, the cells were washed with ice-cold 1X PBS and then centrifuged at 900 × g at 24°C for 5 min. The cell pellets were lysed with radioimmunoprecipitation assay buffer (#89901; Thermo Scientific, Waltham, MA, USA) supplemented with mixed cocktail protease inhibitor and protein phosphatase inhibitor. The lysates were collected into e-tubes and incubated on ice for 30 min. After incubation, the cells were centrifuged at 13,000 × g for 15 min at 4°C. The lysates were quantified by using the Micro BCA Protein Assay Kit (Pierce, Rockford, IL, USA). The samples were denatured with buffer containing 2% SDS, 6% 2-mercaptoethanol, 40% glycerol, 0.004% bromophenol blue, and 0.06 M Tris-HCl at 90-100°C for 6 min, then cooled to 24°C for 5 min. Denatured total proteins (50 μg) were loaded onto 8-12% acrylamide gels. After electrophoresis at 80 V for 20 min and 120 V for 1 hr, proteins were transferred onto 0.2 mm polyvinylidene fluoride membranes (170-4156; Bio-Rad Laboratories, Hercules, CA, USA) using the Trans-Blot1 Turbo system (Bio-Rad Laboratories). The membrane was blocked in with 5% skim milk/TBS-T for 1 hr at 24°C and incubated with primary and secondary antibodies. The following primary antibodies were diluted in 5% bovine serum albumin in TBS-T: rabbit anti-Cyclin D1 antibody (sc-8390; Santa Cruz Biotechnology, CA, USA), mouse anti-CDK2 antibody (sc-6248; Santa Cruz Biotechnology), rabbit anti-CDK4 antibody (sc-260; Santa Cruz Biotechnology), rabbit anti-caspase 3 antibody (#9665; Cell Signaling Technology, Danvers, MA, USA), rabbit anti-PARP antibody (#9542; Cell Signaling Technology), rabbit anti-p-p53 (Ser 15) antibody (#9284; Cell Signaling Technology), rabbit anti-p-ATM antibody (#13050; Cell Signaling Technology), rabbit anti-p-H2AX (Ser 139) antibody (#9718; Cell Signaling Technology), rabbit anti-p-Chk1 (S345) antibody (#2348; Cell Signaling Technology), rabbit anti-p-Chk2 (T68) antibody (#2197; Cell Signaling Technology), and rabbit anti-β-actin antibody (#4970; Cell Signaling Technology). The secondary antibody used was horseradish peroxidase conjugate (AAC10P; Serotec, Oxford, UK) diluted 1:10000 in 5% bovine serum albumin in TBS-T. The membrane was incubated with ECL (170-5060; Bio-Rad Laboratories) for 5 min and developed with an automatic X-ray film processor (JP-33; JPI Healthcare, Seoul, Korea). ImageJ software was used for quantification (NIH, Bethesda, MD, USA). The densities of each band were normalized to those of β-actin or GAPDH.
A549 cells were seeded into a 100-mm dish (50 × 104 cells/dish), incubated for 24 hr at 37°C, and treated with PHMG-p concentrations of 0-4 μg/mL for 36 hr. After exposure to PHMG-p, cells in the culture media and the attached cells were harvested by collecting the media and trypsinization, followed by centrifugation at 2000 rpm at 24°C for 5 min. Referring to the manufacturer’s instructions, the cells were washed twice with ice-cold PBS and then centrifuged at 2000 rpm at 4°C for 5 min. After centrifugation, the cells were resuspended in binding buffer (51-66121E; BD Pharmingen, San Jose, CA, USA) and then 100 μL of the solution containing the cells was transferred into a 5-mL culture tube. Next, 5 μL of FITC Annexin V (51-65874X; BD Pharmingen) and 5 μL PI (5-66211E; BD Pharmingen) were added for staining. The solutions were gently vortexed and incubated for 15 min at 24°C in the dark. A binding buffer was added to each sample and the samples were analyzed by flow cytometry within 1 hr.
Cells that were 70% confluent were exposed to a mixture of Opti-MEM, Lipofectamine RNAiMAX (Invitrogen, Carlsbad, CA, USA) and siRNAs in accordance with the manufacturer’s guidelines. Control siRNA (siCONTROL Non-targeting siRNA Pool, D-001206-13-05) and p53 siRNA (ON-TARGETplus SMARTpool p53, L-003329-00-0005) were purchased from Dharmacon (Lafayette, CO, USA). After 24 hr of transfection, PHMG-p was treated to A549 cells for 36 hr.
A549 cells were seeded into a 6-well plate and incubated for 24 hr at 37°C. Cells were pretreated with p53 siRNA for 24 hr and exposed to PHMG-p for 36 hr. Total RNA was extracted using RNA iso reagent (Takara, Shiga, Japan). cDNA was synthesized by using a RT Premix kit (Bioneer, Daejeon, Korea) with 0.5 μg/mL random primers. PCR was performed using 0.1 μM of the sense and antisense oligonucleotide primers, 10 μL 2 × SYBR Premix Ex Taq (Takara), and 1 μL template DNA in a final volume of 20 μL. PCR amplification was conducted using a CFX Connect Real-Time system (Bio-Rad Laboratories). The primer sequences used in the experiment were as follows: p53, sense, 5′-AGGCCTTGGAACTCAAGGAT-3′; antisense, 5′-CCCTTTTTGGACTTCAGGTG-3′; GAPDH, sense, 5′-AGATCATCAGCAATGCCTCC-3′; antisense, 5′-ATGGCATGGACTGTGGTCATG-3′. The results are shown as relative fold-changes compared to GAPDH.
To measure intracellular ROS generation, the dichlorodihydrofluorescein diacetate (DCFH-DA) method was used. A549 cells were seeded into 48-well plates (3 × 104 cells per well) and incubated for 24 hr at 37°C. After incubation, the cells were treated with PHMG-p (0-4 μg/mL) for 6 hr. Cells were treated with hydrogen peroxide (H2O2) 10 mM for 30 min as a positive control. DCFH-DA (Invitrogen) was added to the cells for 1 hr. Each well was washed twice with PBS, and the cells were lysed in 0.1 N NaOH. DCF (green fluorescence) production was measured at Ex/Em = 485/535 nm using a LS50B fluorometer (Perkin-Elmer, Shelton, CT, USA). Additionally, a Zeiss LSM 700 Laser Confocal Microscope (Thornwood, NY, USA) was used for fluorescence observation.
Immunofluorescence analysis was performed to visualize ROS generation. Cells were seeded onto a coverslip in a 12-well plate at 3 × 104 cells per well and incubated for 24 hr at 37°C. After incubation, the cells were treated with PHMG-p (0-4 μg/mL) for 6 hr. Cells were treated with H2O2 10 mM for 30 min as a positive control. DCFH-DA (Invitrogen) was added to the cells for 1 hr. Each well was washed twice with PBS. The coverslip was separated from the 12-well plate and mounted onto a glass slide. Finally, a Zeiss LSM 700 Laser Confocal Microscope was used to detect the fluorescence of DCFH-DA.
Data were analyzed using Sigma Plot software (Jandel Science Software, San Rafael, CA, USA) and Excel (Microsoft, Redmond, WA, USA). Each in vitro assay was performed at least three times. The data from each assay are expressed as the mean ± standard deviation. Statistical analysis was performed using SPSS version 18.0 (SPSS Inc., Chicago, IL, USA). Differences between groups were assessed by one-way ANOVA followed by Duncan’s post hoc test. Statistical signiﬁcance was accepted at *p < 0.01 or **p < 0.05.
To evaluate the toxic effects of PHMG-p in epithelial cells, we analyzed whether PHMG-p reduces cell proliferation by measuring cell viability and wound healing ability. A549 epithelial cells were exposed to various concentrations of PHMG-p from 12 to 36 hr. Following exposure for 36 hr, PHMG-p significantly reduced cell viability (Fig. 1A). In the PHMG-p-treated group, the degree of wound healing decreased in a time- and dose-dependent manner, which means PHMG-p significantly inhibited cell proliferation (Fig. 1B). These both results indicate that PHMG-p influences cell fates.
Effects of PHMG-p on cell proliferation. A) A549 cells were treated with different concentrations (0-4 μg/mL) of PHMG-p from 12 to 36 hr. Cell viability indicated the percentage viable cells compared to untreated cells. Each value represents the mean ± standard deviation from three replicate wells and is representative of three separate experiments. Values significantly different compared to the control are indicated by **p < 0.01. B) Scratch assay in A549 cells was measured after exposed to PHMG-p for 12, 24, and 36 hr.
Flow cytometry analysis was performed to determine the effect of PHMG-p on the cell cycle and apoptosis which are related to cell proliferation. The percentage of cells in the G0/G1 phase increased while that of cells in the S phase decreased (Fig. 2A). Following flow cytometry analysis, the expression of G1/S checkpoint-related proteins was confirmed by western blotting. Cyclin D1 and CDK2/4 expression decreased in a dose-dependent manner (Fig. 2B). In the apoptosis assay using flow cytometry, cells in early and late apoptosis increased following treatment with 4 μg/mL of PHMG-p (Fig. 2C). The protein levels of cleaved caspase-3 and PARP were dose-dependently increased. Phosphorylation of p53 at serine 15 was measured to confirm the involvement of the p53 pathway. The protein expression of phosphorylated p53 was found to increase dose-dependently (Fig. 2D).
Effects of PHMG-p on cell cycle arrest and apoptosis. A) A549 cells were treated with or without PHMG-p for 36 hr. Cell cycle distribution was analyzed by PI staining using flow cytometry. The bars represent the percent of the cell cycle. B) Protein expression of cell cycle-related markers after 0-4 μg/mL exposure for 36 hr. Cyclin D1, CDK2, and CDK4 were measured by western blot analysis. C) After treatment for 36 hr, the cells were stained with annexin V and PI and analyzed by flow cytometry. D) Protein expression of cleaved caspase-3, PARP, and p-p53 was analyzed by western blot analysis. The relative abundance of each protein was normalized to that of GAPDH or β-actin.
A549 cells were transfected with p53 siRNA for 24 hr. Transfection efficiency was measured by determining the mRNA and protein levels (Fig. 3A and B). When p53 was silenced, cell cycle arrest and apoptosis were partially blocked (Fig. 3C). These results indicate that cell cycle arrest and apoptosis are mediated by the p53 pathway in epithelial cells exposed to PHMG-p.
Role of p53 in cell cycle arrest and apoptosis induced by PHMG-p. A-B) Cells were pretreated with p53 siRNA for 24 hr and exposed to PHMG-p for 36 hr. p53 siRNA transfection efficiency was identified at the mRNA (A) and protein (B) levels. The relative abundance of each protein was normalized to that of GAPDH. Values significantly different compared to the control are indicated by **p < 0.01. C) After treatment with p53 siRNA, cell cycle arrest-related proteins such as CDK2, CDK4, and cyclin D1 and apoptosis-related proteins such as cleaved caspase-3 and PARP were analyzed by western blotting.
The phosphorylation of ATM and H2AX were measured to evaluate the effect of PHMG-p on DNA damage. After PHMG-p treatment for 12 hr, the phosphorylation of ATM (serine 1981) and H2AX (serine 139) increased dose-dependently (Fig. 4A). The ATM inhibitor caffeine was used to confirm the effect of DNA damage on the phosphorylation of p53. The inhibition of ATM activated by PHMG-p reduced the phosphorylation of p53 (Fig. 4B).
Effects of PHMG-p on DNA damage. A) A549 cells were treated with PHMG-p for 12 hr. Ataxia-telangiectasia mutated (ATM) and H2AX were used as DNA damage markers. Protein expression of p-ATM and p-H2AX was evaluated by western blot analysis. B) A549 cells were pre-treated with caffeine (5 mM) and then treated with 4 μg/mL PHMG-p for 12 hr. To identify the effects of the ATM inhibitor (caffeine) on the ATM pathway, the expression of related proteins such as p-ATM and p-p53 was evaluated by western blotting. The relative abundance of each protein was normalized to that of GAPDH.
ROS is associated with inducing DNA damage. To visualize ROS generation, immunofluorescence was conducted. Cells treated with PHMG-p produced ROS dose-dependently (Fig. 5A and B). The antioxidant N-acetylcystein (NAC), a glutathione precursor, decreased the protein levels of p-ATM and p-H2AX induced by PHMG-p (Fig. 5C).
Role of reactive oxygen species generated by PHMG-p. A) A549 cells were treated with PHMG-p for 6 hr and 10 mM H2O2 for 30 min. ROS generation was observed by confocal microscopy and B) fluorescence measurement of the reporter DCF; the result is shown as a percent relative to the control. C) Cells were pre-incubated with N-acetylcysteine (NAC) for 1 hr and then treated with PHMG-p for 12 hr to determine the effects of DNA damage induced by ROS. ATM and H2AX were used as DNA damage markers and evaluated by western blot analysis. The relative abundance of each protein was normalized to that of GAPDH.
Many studies have shown that recurrent epithelial cell injury leads to cell cycle arrest and apoptosis, which results in aberrant repair and inappropriate re-epithelialization. While damage to epithelial cells takes place, the epithelial barrier collapses by gradually losing integrity and causes cell cycle arrest, which may be a protective mechanism to allow for the repair of injured cells. If epithelial cells are not repaired or fail to proliferate, this may lead to apoptosis (O’Reilly, 2001). Excessive damage to epithelial cells eventually stimulates the release of pro-fibrotic cytokines, which activate inflammation and fibroblasts proliferation to create a fibrotic environment (Wilson and Wynn, 2009; Camelo et al., 2014). Therefore, epithelial injury, including cell cycle arrest and apoptosis, is considered the main initiator event in the early stages of pulmonary fibrosis pathogenesis (Uhal, 2003; Uhal, 2008; Yang et al., 2010). In this study, we investigated the effects of PHMG-p on the regulation of cell cycle arrest and apoptosis as an initial response to fibrosis. Our results demonstrated that PHMG-p increased ROS-mediated DNA damage, which activates the p53 pathway and eventually induces G1/S cell cycle arrest and apoptosis.
In the balance between cell proliferation and apoptosis, the regulation of the cell cycle and apoptosis are essential for growth and homeostasis in normal cells. PHMG-p inhibited cell proliferation (Fig. 1); G1/S cell cycle arrest and apoptosis were observed in cells exposed to PHMG-p (Fig. 2). CDKs and cyclins are key factors which regulate switching between cycle phases. Cyclin D1 and CDK2/4 are essential proteins during the G1/S phase of the cell cycle (Lim and Kaldis, 2013). The cyclin D1-CDK4 complex phosphorylates Rb, releases E2F, and increases cyclin E-CDK2, which allows cell cycle progression to the S phase (Lim and Kaldis, 2013; Barr et al., 2017). Cells exposed to PHMG-p showed G1/S cell cycle arrest with suppressed levels of cyclin D1 and CDK2/4 proteins (Fig. 2B), suggesting that PHMG-p induced G1/S cell cycle arrest by blocking the cyclin D1–CDK4/6 and cyclin E–CDK2 complexes which are important for the G1/S transition phase. Interestingly, epithelial cells at G1/S phase can be associated with epithelial-mesenchymal transition (EMT). EMT is a dynamic cellular process in which polarized epithelial cells lose their epithelial phenotype and gain mesenchymal characteristics. Certain chemicals that induce pulmonary fibrosis, including PHMG-p, as well as bleomycin and paraquat, promote EMT (Kim et al., 2018; Shin et al., 2018). Song’s (2007) study on EMT and apoptosis regulated by TGF-β demonstrated that apoptosis was induced mostly in cells at the G2/M phase, whereas EMT was only induced in cells at the G1/S phase. Taken together, this suggests that the G1/S cell cycle arrest induced by PHMG-p may precondition cells undergoing EMT, contributing to the expansion of fibroblasts. Epithelial cells at other phases may result in apoptosis.
PHMG-p induced apoptosis with increasing the cleavage of PARP and caspase-3. The apoptotic cells in the lungs are cleared by macrophages, which lead to inflammatory and fibrotic responses (Wang et al., 2006). Recently, Park et al. (2018) reported that epithelial cells exposed to PHMG-p were attached to lung fibroblasts and then taken up, which induced the release of cytokines. Sisson et al. (2010) showed that apoptosis of alveolar epithelial cells after injury resulted in fibroblast proliferation and collagen deposition in the mouse model. Therefore, in contrast with growth arrest, apoptotic bodies induced by PHMG-p may contribute to a pro-fibrotic microenvironment by stimulating other cells.
The phosphorylation of p53 is an important factor for the regulation of cell cycle arrest and apoptosis (Amundson et al., 1998). Phosphorylated p53 induces p21, a cyclin-dependent kinase inhibitor that induces cell cycle arrest by binding to the cyclin/CDK complex (Chen, 2016). Additionally, the activation of p53 triggers mitochondria insults, which activate pro-apoptotic proteins, resulting in apoptosis (Chen, 2016). We observed that cell cycle arrest and apoptosis induced by PHMG-p were attenuated by p53 silencing (Fig. 3C), indicating that they are dependent on the p53 pathway. Phosphorylation of p53 is caused by cellular stress/DNA damage, which leads to activation of the ATM pathway. The proteins that detect DNA damage, including ATM and H2AX, were phosphorylated by treatment with PHMG-p (Fig. 4A). In addition, the expression of p-p53 was reduced by treatment with caffeine, suggesting that p53 activation is caused by DNA damage (Fig. 4B). The main cause of cellular damage is ROS, which generates free radicals and react with DNA to alter DNA structure and function. ROS plays an important role in pulmonary fibrosis by stimulating apoptosis and inflammatory pathways (Todd et al., 2012). If high concentrations of ROS are present, signal transduction, DNA damage, and p53 levels are increased. In addition, ROS causes G1/S arrest and ultimately leads to apoptosis, which removes the cells damaged by ROS (Boonstra and Post, 2004). PHMG-p induced ROS generation, which was associated with DNA damage (Fig. 5). The level of p-ATM protein induced by PHMG-p was decreased by treatment with antioxidant (Fig. 5C). This suggests that the activation of p53 was induced to remove the DNA damage caused by ROS. The activation of p53 may be linked to cell cycle arrest and apoptosis. Taken together, this suggests that PHMG-p triggers G1/S arrest and apoptosis through the ROS/ATM/p53 pathway in lung epithelial cells (Fig. 6).
Proposed scheme of cell cycle arrest and apoptosis pathway induced by PHMG-p. PHMG-p induces ROS generation and leads to DNA damage. PHMG-p phosphorylates H2AX and ATM, which increases the expression of phosphorylated p53. Activated p53 induces G1/S arrest and apoptosis.
Bleomycin and paraquat are the most widely studied chemicals known to induce pulmonary fibrosis. Similar to PHMG-p, cell cycle arrest and apoptosis in lung epithelial cells were observed in the bleomycin and paraquat treatment model (Takeyama et al., 2004; Jang et al., 2018). Exposing bleomycin to epithelial cells led to ROS generation, which catalyzes the formation of DNA lesions (Wallach-Dayan et al., 2006). This is followed by the activation of H2AX and ATM and induces p53 signaling. This leads to G2/M cell cycle arrest (Chen and Stubbe, 2005; He et al., 2016). In addition, paraquat leads to pulmonary fibrosis by damaging epithelial cells through oxidation (Takeyama et al., 2004; Blanco-Ayala et al., 2014). Therefore, we suggest that p53-mediated apoptosis and cell cycle arrest of lung epithelial cells caused by ROS/DNA damage is a key process in pulmonary fibrosis induced by PHMG-p.
In conclusion, we demonstrated that epithelial cell injury occurs following PHMG-p exposure. PHMG-p induced G1/S cell cycle arrest and apoptosis through a p53-dependent pathway. Upstream, DNA damage by ROS was confirmed by the inhibition of ROS generation and DNA damage. These findings indicated that PHMG-p triggered cell cycle arrest and apoptosis through the ROS/ATM/p53 pathway in lung epithelial cells. Our study contributes to the understanding of the early stages of the epithelial injury mechanism of fibrosis caused by PHMG-p.
This work was supported by the National Research Foundation of Korea Grant funded by the Korean Government (NRF-2017R1C1B5018375).
The authors declare that there is no conflict of interest.