Triphenyl phosphate-induced cell injury through endoplasmic reticulum stress in human kidney cells

Abstract

Triphenyl phosphate (TPhP) is a typical organophosphorus flame retardant (OPFR). Due to its high production and widespread use, exposure to TPhP has been shown to induce nephrotoxicity in animal models. Endoplasmic reticulum (ER) stress is found to be correlated with kidney disease caused by exogenous environmental pollutants. Nevertheless, the connection between ER stress and the nephrotoxic effects caused by TPhP is limited. In this study, human renal tubular epithelial cells (HKC) were chosen to explore the effects of TPhP on cell viability, cell apoptosis, and ER stress. Our study indicated that cell viability was dramatically inhibited in a dose-dependent manner. The half lethal concentration (LC₅₀) value of TPhP after 48 hr exposure is 126.4 µM. A concentration-related Caspase-3 activation and apoptosis occurrence were observed in HKC cells following TPhP treatment. Additionally, the induction of ER stress was demonstrated by the up-regulated expression of ER stress-related genes. To elucidate the role of ER stress in cell damage, sodium 4-phenylbutyrate (4-PBA), an ER stress inhibitor, was used in the co-treatment with TPhP. Results revealed that 4-PBA treatment effectively alleviated TPhP-induced ER stress and cytotoxicity in HKC cells. Taken together, these results indicated that ER stress plays a primary role in TPhP-induced nephrocyte damage and 4-PBA could attenuate these effects.

INTRODUCTION

As a typical organophosphorus flame retardant (OPFR), triphenyl phosphate (TPhP) is widely used in commercial and industrial products to reduce fire risks, enhance material plasticity or inhibit foam production. (van der Veen and de Boer, 2012). Since TPhP is physically incorporated into polymers rather than chemically bonded, it can be easily released into the surrounding environment during usage. (van der Veen and de Boer, 2012). Nowadays, TPhP has already been detected in indoor air (Liu et al., 2023), surface water (Kim and Kannan, 2018), drinking water (Zhang et al., 2022), sediment (Li et al., 2019a; Li et al., 2023), house dust (Chen et al., 2020), as well as in foods, including fish, fruit, rice and vegetables (Gbadamosi et al., 2021). For instance, the mean concentration of TPhP was 610 ng/g in indoor dust from China (Chen et al., 2020), 4,260-1,710,000 ng/g for sediment from e-waste recycling areas of China (Li et al., 2019a), up to 14,000 ng/L in surface water from Norway (Green et al., 2008), and a mean level of 26.14 ng/g for rice from Australia (Li et al., 2019b). It is worth noting that TPhP has been detected in various human samples, including urine, serum, breast milk, nail and hair (Chen et al., 2021; Gao et al., 2020). In a survey of OPFRs in breast milk from China, TPhP was found to be detected 99% of breast milk, with the maximum estimated daily intake being 10,900 ng/(kg bw⁻¹·day⁻¹) (Chen et al., 2021). In another study, TPhP was identified as the primary contributor to daily ∑OPFRs intake from tap water, with exposure doses ranging from 0.63 to 11.33 ng/kg/d. (Zhang et al., 2022). Given that people are often inadvertently exposed to TPhP through daily life, its potential health hazards have attracted extensive concerns. In 2020, TPhP was recommended as a high-priority chemical for health risk assessment by the United States. Environmental Protection Agency (EPA) (OCSPP, 2019).

Toxicological studies have suggested that TPhP is an endocrine disruptor, inducing neurotoxicity, hepatotoxicity, immunotoxicity, as well as reproductive and developmental toxicity (An et al., 2023; Chen et al., 2015; Hong et al., 2022; Li et al., 2019c; Wang et al., 2019). In experimental studies with mouse or fish models, the exposure to TPhP could disturb placental tryptophan metabolism and cause abnormal neurobehavior in male offspring (Hong et al., 2022), induce ovary retardation and decrease egg production (Li et al., 2019c), and impair testicular histopathology (Chen et al., 2015). Liu et al. reported that TPhP exposure could affect plasma sex hormone levels, along with an increase of vitellogenin in zebrafish, which was found to be correlated with the alteration hypothalamic-pituitary-gonadal axis (Liu et al., 2013). Furthermore, studies have shown that TPhP exhibits hepatotoxicity, disrupts lipid metabolism, and results in the increased liver weight, fat accumulation, and hepatic steatosis (An et al., 2023; Wang et al., 2019). Besides these adverse outcomes, research has also shown that TPhP can negatively impact kidney function (Cui et al., 2020; Tsai et al., 2022). For example, in the study of Cui et al, they observed that TPhP could induce protein residue accumulation, lipid accumulation or inflammatory response in kidneys, and promote renal structure damage after exposing to mice with 1 mg/kg/d TPhP for 12 weeks, indicating potential kidney health hazards associated with TPhP contamination (Cui et al., 2020). Meanwhile, another typical OPFR, tris(1,3-dichloro-2-propyl) phosphate (TDCPP) has been found to enhance renal cancer cells proliferation or migration, and to promote the volume and weight of kidney tumors in vivo experiments (Zhou et al., 2022). Epidemiological studies have also suggested that some OPFRs are potential chemical determinants of chronic kidney disease (Tsai et al., 2022). However, thus far, studies regarding the toxicological assessment of OPFRs to the human kidney are still limited.

The endoplasmic reticulum (ER), a pivotal organelle in cells, plays important roles in protein folding and processing, lipid biosynthesis, and calcium homeostasis. Growing evidence indicates that ER stress plays a key role in cell damage, including cell apoptosis, autophagy and ferroptosis. For example, Tris (1,3-dichloro-2-propyl) phosphate (TDCIPP)-induced cell apoptosis was reported to have a relationship with ER stress-related apoptotic pathways in both human cornea epithelial cells and mouse spermatocyte cells (Feng et al., 2024; Xiang et al., 2017). It was also shown that ER stress is linked to autophagy and apoptosis in endothelial cells triggered by PM2.5 exposure. (Wang and Tang, 2020). Previous studies also revealed that ER stress was involved in renal diseases, such as diabetic nephropathy, acute kidney injury and renal fibrosis (Cybulsky, 2017). However, the relationship between ER stress and the nephrotoxic effects of TPhP is not well explained. The aim of this study was to explore whether ER stress was involved in the renal cell injury triggered by TPhP exposure, to identify the potential sensitive organelle, and to provide a scientific evidence for the mechanism of TPhP’s nephrotoxicity.

MATERIALS AND METHODS

Chemicals

TPhP (CAS No. 115-86-6) and 4-PBA (CAS No. 1716-12-7) were acquired from Sigma-Aldrich (St Louis, MO, USA). The following reagents and materials were used in this study: penicillin-streptomycin solution and trypsin-EDTA solution were sourced from Invitrogen (ThermoFisher Scientific, Waltham, MA, USA). The FITC Annexin V/Dead Cell Apoptosis Kit (catalog number: V13242), Hoechst 33342 (catalog number: H21492), and SYBR™ Green PCR Master Mix (catalog number: 4367659) were also purchased from the same supplier. For RNA extraction, Mini Kits from Qiagen Biotech Co. (Germany) were utilized, while cDNA synthesis was performed using kits provided by TaKaRa Biotech Co. (Japan). Cell culture dishes and plates were obtained from Corning Inc. (NY, USA).

Cell culture

HKC cells were acquired from the Gaining Biotechnology (Shanghai, China) and maintained in 75 cm² petri dishes containing 25 mL of DMEM/F12 medium. The culture medium was supplemented with 10% fetal bovine serum (Gibco, catalog number: 16000-044), 100 µg/mL streptomycin and 100 IU/mL penicillin. Cells were incubated at 37°C in a humidified atmosphere of 95% air and 5% CO₂. The medium was replaced, and cells were subcultured three to four times per week. All experiments were conducted using cells within 10 passages to ensure consistency and reliability.

TPhP exposure

TPhP was dissolved in dimethyl sulfoxide (DMSO) as a stock solution and then diluted with culture medium to appropriate concentrations for subsequent experiments. To investigate the impact of TPhP on cell viability, HKC cells were treated with TPhP at concentrations ranging from 0 to 200 µM for 48 hr. According to the result of half lethal concentration (LC₅₀) calculation, HKC cells were exposed to TPhP at three concentrations (LC₅₀ = 125 µM, 3/5 LC₅₀ = 75 µM, 1/5 LC₅₀ = 25 µM) and incubated at 37°C for 48 hr in the following experiments. Additionally, to further analyze the role of ER stress in TPhP-triggered toxic effects, the cells were treated with TPhP (125 μM), 4-PBA (0.5 mM), or a combination (125 μM TPhP + 0.5 mM 4-BPA) for 48 hr.

Assay of cell viability

Cell viability was analyzed using a cell counting kit-8 (CCK-8). HKC cells were seeded in a 96-well microplate with a density of 1×10⁴ cells per well. After 48 hr of exposure to TPhP (0-200 μM) or DMSO (0.05%), 15 μL CCK-8 solution was added and incubated at 37°C for 2 hr. Absorbance was measured at 450 nm using a microplate reader (Bio-rad, California, USA).

Apoptosis assay

Cell apoptosis of HKC cells was detected using FITC Annexin V-FITC/PI Kit. Briefly, HKC cells were seeded in a 96-well blank plate with a density of 1×10⁴ cells per well. After 48 hr of exposure to TPhP (25, 75 and 125 µM) or DMSO (0.05%), cells were washed twice with phosphate-buffered saline (PBS). Subsequently, HKC cells were stained with Annexin V-FITC/PI staining solution for 15 min. Then, cell nuclei were probed with Hoechst 33342 for 5-10 min. Finally, cell apoptosis was imaged and analyzed using an Operetta™ High Content Screening (HCS) instrument.

Analysis of Caspase-3 activity

Caspase-3 activity was analyzed using commercial assay kits (Beyotime Institute of Biotechnology, China). In brief, cells were plated in 6-well plates with an initial density of 5×10⁵ cells per well, after adherent incubation overnight, cells were treated with TPhP (25, 75 and 125 µM) or DMSO (0.05%) for 48 hr. HKC cells were lysed with ice-cold lysis buffer. After centrifugation, the supernatant was collected and incubated with Ac-DEVD-pNA overnight at 37°C. Subsequently, absorbance was measured at 405 nm using a microplate reader. A standard curve was generated and Caspase-3 activity was normalized to protein content and presented as a percentage relative to the control group.

Real-time RT-PCR

HKC cells were plated in 6-well plates with a density of 5×10⁵ cells/well, after adherent incubation overnight, cells were treated with TPhP (25, 75 and 125 µM) or DMSO (0.05%) for 48 hr. Total RNA of HKC cells was extracted using the RNeasy mini kit (Qiagen, Hilden, Germany). The concentration and purity of total RNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Fisher, MA, USA). We synthesized the first strand cDNA from 1 µg of total RNA with a reverse transcription kit (Takara, Dalian, China). PCR primers (Table 1) of ER-stress related genes (Bip, CHOP, XBP1, eIF2α, ATF4, ATF6 and Caspase-12) were designed using Primer 6.0 software. Primer specificity was verified by observing melting curves, and its efficiencies were determined using a standard curve approach. Only primers with a single peak in their melting curves and amplification efficiency between 90% and 110% were selected for further analysis. The mRNA expression levels were quantified using the QuantStudio Q7 PCR system with SYBR Green PCR Master Mix reagent kits (Thermo Fisher, MA, USA). A portion of 0.5 μg cDNA was amplified in a 20 μL PCR reaction system including 2×SYBR Green PCR Master Mix, 1 μM forward and reverse primers and RNase free water. The PCR procedure included an initial denaturation step at 95°C for 10 min, followed by 40 cycles of denaturation at 95°C for 15 sec and annealing/extension at 60°C for 1 min. Every sample was analyzed in triplicate. The relative gene expression levels were calculated via the comparative Ct method (2^-ΔΔCt)), normalized to the endogenous control β-actin, and expressed as fold changes relative to the vehicle control.

Table 1. Sequences of primers used for real-time RT-PCR amplification.

Target gene	GenBank Accession No	primer sequences	Product length(bp)
β- actin	NM_001101.3	FW: ATTGGCAATGAGCGGTTCC	144
		RW: TGTGTTGGCGTACAGGTCTT
Bip	X87949.1	FW: GGAGGACAAGAAGGAGGA	148
		RW: AGTGAAGGCGACATAGGA
eIF2α	NM_032025.5	FW: GGCATTATACTGGCTCTATCT	132
		RW: CACTTGGAACTGCTTGGT
ATF4	NM_001675.4	FW: ACCTTCTTACAACCTCTTCC	114
		RW: TGGCTTCCTATCTCCTTCA
CHOP	NM_001195056.1	FW: CTGGAAGCCTGGTATGAG	118
		RW: GGTCAAGAGTGGTGAAGAT
XBP1	NM_001079539.2	FW: GCATTCTGGACAACTTGGA	116
		RW: GGAGGCTGGTAAGGAACT
ATF6	AB015856.1	FW: GCACAGGACACATCAGAT	97
		RW: GTTCTTCAGTTAGCACCATC
Caspase-12	NM_001191016.3	FW: CCTCAGACAGCACATTCC	135
		RW: GGCAGTTACGGTTGTTGA

Protein expression of p-eIF2α, t-eIF2α, CHOP, XBP1s, and Caspase-12

Immunofluorescence staining was performed to detect the protein expression of ER stress markers, including phosphorylated eIF2α (p-eIF2α), total eIF2α (t-eIF2α), CHOP, XBP1s, and Caspase-12. Briefly, cells were plated in black-wall 96-well microplates and treated with different compounds, fixed with 4% formaldehyde, and washed twice with PBS. Cell membranes were permeabilized with 0.1% Triton X-100 for 10 min, followed by blocking in 5% bovine serum albumin (BSA) prepared in PBS for 30 min at room temperature. Primary antibody incubation was carried out overnight at 4°C using the following specific antibodies and dilutions in 5% BSA-PBS: rabbit anti-p-eIF2α (Cell Signaling, 3398; a dilution of 1:150), rabbit anti-eIF2α (Abcam, 169528; a dilution of 1:400), mouse anti-CHOP (Cell Signaling, 2895; a dilution of 1:500), rabbit anti-XBP1s (Cell Signaling, 40435; a dilution of 1:200), and rabbit anti-Caspase-12 (ThermoFisher, PA5-19963; a dilution of 1:300). After washing with PBS, cells were incubated with species-appropriate secondary antibodies for 1 hr at room temperature protected from light: Alexa Fluor Plus 488-conjugated anti-mouse (ThermoFisher, A32723) or Alexa Fluor Plus 555-conjugated anti-rabbit (ThermoFisher, A32732). Cell nuclei were labeled with Hoechst 33342 and subjected to image acquisition on HCS platform.

Statistical analysis

All experiments were conducted at least in triplicate. Statistical analysis was performed using SPSS 26.0 software and data were expressed as the means ± standard deviation (SD). Differences between the control and treatment groups were evaluated by one-way analysis of variance (ANOVA) followed by post-hoc LSD tests with A p < 0.05 or p < 0.01 was considered statistically significant. Concentration-response curves were plotted, and LC₅₀ values were calculated using GraphPad Prism 5 software.

RESULTS

TPhP inhibited cell viability

The viability of HKC cells after exposure to 0, 6.25, 12.5, 25, 50, 75, 100, 150 and 200 µM TPhP for 24 hr was evaluated by CCK-8 assay (Fig. 1). The results indicated that cytotoxicity was enhanced as TPhP concentration increased. As shown in Fig. 1 A, cell viability of HKC cells was significantly reduced by 23.2%, 43.8%, 62.4% and 67.8%, respectively after 48 hr exposure to TPhP at dosage of 75, 100, 150 and 200 µM. Furthermore, according to nonlinear regression analysis, the 48 hr LC₅₀ value of TPhP was determined to be 126.4 µM (Fig. 1B).

Fig. 1

Cell viability in HKC cells. (A) Effect of TPhP on cell viability following treatment with TPhP (0-200 μM) for 48 hr. (B) Logarithmic transformation of TPhP concentrations and cell viability data was fit to a nonlinear regression curve to determine LC₅₀. Results are means ± SD of three replicate experiments (n = 6 wells per exposure dose). Compared with the control group, * p < 0.05 and ** p < 0.01.

TPhP enhanced cell apoptosis

Cell apoptosis, a major form of programmed cell death, was assessed using an Annexin V-FITC/PI kit to investigate the potential cytotoxic effects of TPhP. Following staining, early-stage apoptotic cells were labeled exclusively with Annexin V-FITC (green), while late-stage apoptotic cells were double-stained with both Annexin V-FITC (green) and propidium iodide (red). As illustrated in Fig. 2A, 75 and 125 µM TPhP-treated groups exhibited a significant increase in both green and red fluorescence intensities. In contrast, minimal green and red fluorescence was observed in the control and 25 µM TPhP-treated groups. Based on quantitative analysis, it was shown that TPhP treatment triggered apoptosis in a dose-dependent manner, with apoptotic ratio of 29.7% and 57.5% in the 75 and 125 µM TPhP-treated groups, respectively. However, no significant differences were observed for 25 µM TPhP treatment when compared with the control group (Fig. 2B).

Fig. 2

Cell apoptosis and Caspase-3 activity in HKC cells. Effect of TPhP on cell apoptosis and Caspase-3 activity in HKC cells following treatment with TPhP (0, 25, 75 and 125 μM) for 48 hr. (A) Fluorescence images of cell apoptosis. Apoptotic cells stained with Annexin V-FITC/PI probe. Viable cells show negative signals of Annexin V-FITC and PI. Early apoptotic cells are Annexin V-FITC positive and PI negative. Late apoptotic cells are both Annexin V-FITC and PI positive. Scale bar = 100 µm (same in every image and shown in the image of nuclei for example). (B) quantitative analysis of cell apoptosis using HCS platform. (C) Caspase-3 activity. Results are means ± SD of three replicate experiments. Compared with the control group, ** p < 0.01.

TPhP increased Caspase-3 activity

To further evaluate the induction of apoptosis, we determined the enzymatic activity of caspase-3, a key executioner protease in the process of apoptosis. Caspase-3 activity exhibited a concentration-dependent increase, rising by 14.8%, 57.9%, and 118.3% following exposure to 25, 75, and 125 µM TPhP, respectively. Although Caspase-3 activity was elevated in the 25 µM TPhP group, no statistical difference was observed compared to the control group (Fig. 2C).

TPhP induced ER stress

Accumulating evidence demonstrates the pivotal involvement of the endoplasmic reticulum (ER) in mediating toxicant-induced apoptotic cell death. To examine the potential induction of ER stress by TPhP in HKC cells, the mRNA levels of ER stress-related markers were determined using quantitative real-time PCR. As illustrated in Fig. 3, the transcriptional levels of Bip, CHOP, ATF6, eIF2α, ATF4, XBP1, and Caspase-12 were significantly upregulated at TPhP concentrations of 75 and 125 µM compared to the control group. However, eIF2α mRNA expression was only elevated following exposure to 125 µM TPhP. These findings suggested that TPhP treatment triggers ER stress in HKC cells.

Fig. 3

Levels of ER stress-related genes in HKC cells. Effect of TPhP on ER stress-related genes in HKC cells following treatment with TPhP (0, 25, 75 and 125 μM) for 48 hr. Results are means ± SD of three replicate samples. Compared with the control group, * p < 0.05 and ** p < 0.01.

4-BPA alleviated the adverse effects of TPhP

To explore the role of ER stress in TPhP-induced toxicity in HKC cells, the cells were exposed to 125 µM TPhP for 48 hr, either with or without the ER stress inhibitor 4-PBA. Then, cell viability, cell apoptosis, Caspase-3 activity and ER-stress-related genes and protein were investigated. As shown in Fig. 4A, cell viability was increased upon co-treatment with 4-PBA compared to that group exposed to 125 µM TPhP alone (72.6% compared with 59.7%), however, no statistical difference was observed between these two groups. The apoptotic percentage was reduced to 34.1%, compared to 57.5% in cells treated solely with 125 µM TPhP (Fig. 4B). Moreover, the Caspase-3 activity was decreased to 166.5% compared with 218.3% in cells treated with 125 µM TPhP alone (Fig. 4C). Real-time PCR analysis revealed that co-treatment with 4-PBA significantly reduced the mRNA expression levels of ER stress-related genes compared to cells exposed to 125 µM TPhP alone (Fig. 5). Similarly, immunofluorescence assay further confirmed the above findings, showing that co-treatment with 4-PBA reduced the protein levels of ER stress-related markers (p/t-eIF2α, CHOP, XBP1, and Caspase-12) relative to treatment with 125 µM TPhP alone (Fig. 6.).

Fig. 4

4-BPA alleviated the cytotoxicity of TPhP in HKC cells. Cell viability (A), Cell apoptosis (B) or Caspase-3 activity (C) of HKC cells following treatment with TPhP (125 μM) in the absence or presence of 4-PBA (0.5 mM) for 48 hr. Results are means ± SD of three replicate samples. Compared with the control group, ** p < 0.01. Compared with the TPhP (125 μM)-treated group, ^# p < 0.05 and ^## p < 0.01.

Fig. 5

4-PBA partially alleviated ER stress in HKC cells. (A) Expression levels of ER stress-related genes in HKC cells following treatment with TPhP (125 μM) in the absence or presence of 4-PBA (0.5 mM) for 48 hr. (Results are means ± SD of three replicate samples. Compared with the control group, * p < 0.05 and ** p < 0.01. Compared with the TPhP (125 μM)-treated group, ^# p < 0.05 and ^## p < 0.01.

Fig. 6

4-PBA partially alleviated ER stress in HKC cells. (A) Fluorescence images of p-eIF2α, t-eIF2α, CHOP, XBP1s and Caspase-12 in HKC cells following treatment with TPhP (125 μM) in the absence or presence of 4-PBA (0.5 mM) for 48 hr. Blue fluorescence represents nuclei labeled by Hoechst 33342, green or red fluorescence represents the targeted protein labeled by green (Alexa Fluor Plus 488) or red (Alexa Fluor Plus 555) fluorescent secondary antibodies. Scale bar = 100 µm (same in every image and shown in the image of nuclei for example). (B) Quantitative analysis of protein level using HCS platform. Results are means ± SD of three replicate samples. Compared with the control group, * p < 0.05 and ** p < 0.01. Compared with the TPhP (125 μM)-treated group, ^# p < 0.05 and ^## p < 0.01.

DISCUSSION

Nowadays, TPhP has attracted worldwide attention because of its potential environmental and health risks (Wang et al., 2021). For example, besides neurotoxicity, hepatotoxicity, immunotoxicity and reproductive toxicity, TPhP has been also observed to result in nephrotoxicity in mice (Cui et al., 2020). Studies have indicated that ER stress is correlated with environmental pollutant-induced chronic kidney diseases (Gallazzini and Pallet, 2018), which means that ER stress might be involved in TPhP-induced nephrotoxicity. However, the specifics of the link between nephrotoxic effects and ER stress remain to be clarified. Therefore, we selected HKC cells as an in vitro model to evaluate whether TPhP exposure can induce renal dysfunction and explore the potential role of ER stress in this pathological process.

Presently, TPhP decreased cell viability at concentration ≥75 µM, with LC₅₀ value being 126.4 µM. In contrast, TPhP showed relatively stronger cytotoxicity in GC-2 cells compared with HKC cells (48 hr LC₅₀ values of TPhP for GC-2 cells was 61.61 µM) (Feng et al., 2023). Whereas, in A549 cells, TPhP resulted in a 59.2% reduction in cell viability at a concentration of up to 300 µM after 48 hr of exposure, indicating relatively lower cytotoxicity compared to HKC cells, which showed a 56.2% decrease in cell viability at a concentration of 100 µM TPhP (Yu et al., 2019). These differences may result from variations in experimental models or cell sources, which exhibit different susceptibilities to TPhP.

Apoptosis is recognized as a key pathological feature of chronic kidney diseases (Nowak and Edelstein, 2020). As a biomarker of cell apoptosis, Capsase-3 is responsible for executing apoptosis in cells responding to endogenous or exogenous inducers (Wang and Tang, 2020). For example, activation of Caspase-3 and cell apoptosis induced by OPFRs have been observed in various cell models, including human lung cells, human corneal cells and mouse spermatocyte cells, etc. (Feng et al., 2024; Nowak and Edelstein, 2020; Xiang et al., 2017; Yu et al., 2019). It was also reported that cell apoptosis and Caspase-3 activation were correlated with renal diseases (Davis and Ryan, 1998; Erdemli et al., 2024). In current study, increased Caspase-3 activity was accompanied by a dramatic elevation in cell apoptosis and inhibition of cell viability in HKC cells after exposing to 75 and 125 µM TPhP, suggesting that cell apoptosis induced by TPhP treatment might ultimately lead to renal cytotoxicity.

The endoplasmic reticulum, a critical organelle, plays a vital role in regulating protein folding, protein maturation and calcium homeostasis, which are crucial for sustaining normal biological functions (Li et al., 2020). Disturbances in the ER homeostasis can lead to the accumulation of unfolded or misfolded proteins in the ER lumen, resulting in a condition known as ER stress. (Giamogante et al., 2021). Numerous studies have shown that ER stress plays a key role in triggering cell apoptosis (Beilankouhi et al., 2023; Ko et al., 2022). Bip acts as a crucial regulator of ER stress modulates the activation of three transmembrane ER stress sensors: inositol-requiring enzyme 1 (IRE1), protein kinase RNA-like endoplasmic reticulum kinase (PERK), and activating transcription factor 6 (ATF6) (Giamogante et al., 2021). Under physiological conditions, these three proteins remain bound to Bip, maintaining an inactive state. However, under stress conditions, Bip dissociates from these proteins, leading to their activation and initiating the unfolded protein response (UPR) (Zhang et al., 2015). In the PERK pathway, activation of PERK leads to the phosphorylation of eIF2α, which in turn promotes the transcriptional expression of ATF4 and the upregulation of CHOP, a key inducer of apoptosis (Giamogante et al., 2021; Ibrahim et al., 2019). In the IRE1 pathway, activated IRE1 could promote the maturation of XBP1, which then initiates CHOP expression. Simultaneously, the enzymatic activity of Caspase-12 is also increased by IRE1 activation. As the apoptosis-related factors, Caspase-12 can trigger apoptosis via activating Caspase-3 activity, whereas, CHOP could induce apoptosis by affecting the ratio of Bcl/Bax (McCullough et al., 2001). In Wu’s study, ER stress was shown to mediate pancreatic cell apoptosis, as evidenced by elevated expression of CHOP and Caspase-12 (Wu et al., 2021). Similarly, Wang and Tang reported that PM2.5-induced cell apoptosis is mediated through ER stress, in which the induction of Caspase-12 and CHOP expression are key events (Wang and Tang, 2020). Currently, the observed upregulated transcriptional levels of Bip, CHOP and Caspase-12, along with elevated mRNA expression of eIF2α, ATF4, XBP1 and ATF6, strongly suggested that ER stress was triggered by TPhP exposure.

4-PBA, a chemical chaperone, inhibits ER stress by facilitating the proper folding of misfolded proteins or inhibiting their aggregation within the ER (Kolb et al., 2015). Various studies have documented that 4-PBA can alleviate negative effects by inhibiting ER stress (Xia et al., 2022; Zeng et al., 2017). In our work, 4-PBA was used to explore the relationship between ER stress and nephrotoxic effects in HKC cells induced by TPhP. The results implied that co-treatment with 4-PBA reduced the expression of ER stress-related genes and protein. Additionally, following co-treatment with 4-PBA, reduced apoptotic ratio, increased cell viability, and decreased Caspase-3 activity were observed in TPhP-treated HKC cells. These findings indicated that 4-PBA can suppress ER stress activation and mitigate ER stress-related cell damage induced by TPhP exposure.

In this study, we observed that exposure to TPhP for 48 hr resulted in a dose-dependent reduction in HKC cell viability, with an LC₅₀ value of 126.4 µM. Furthermore, treatment with TPhP induced renal cell apoptosis, activated Caspase-3 activity, and upregulated the mRNA expression of ER stress-related genes. Notably, these adverse effects could be alleviated by 4-PBA, an ER stress inhibitor. Therefore, these results suggest that TPhP-induced renal cytotoxicity is mediated via ER stress-regulated apoptotic pathway, the induction of ER stress-related genes and the subsequent increase in Caspase-3 activity are critical events. The collective findings suggest that the endoplasmic reticulum (ER) is a sensitive target organelle in HKC cells, offering scientific evidence to support further investigation into the mechanisms of TPhP's nephrotoxicity.

Funding

This study was supported by the Talent Development Plan for High-level Public Health Technical Personnel Project (Gugan- 03-41).

Conflict of interest

The authors declare no competing interests that could influence the work described in this study.

Data availability

The data in this study are included in the article/supplementary materials. Contact the corresponding author(s) directly to request the underlying data.

Author contributions

Hejun Duan: sampling, paper writing. Liyu Huang: Software, Data analysis, Editing. Yixing Feng: Organization, Investigation, Experiment, Funding acquisition.

Ethical approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

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