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Cadmium (Cd) is an environmental contaminant that exhibits renal toxicity. The target transcription factors involved in Cd renal toxicity are still unknown. In this study, we demonstrated that Cd decreased the activity of the ARNT transcription factor, and knockdown of ARNT significantly decreased the viability of human proximal tubular HK-2 cells. Microarray analysis in ARNT knockdown cells revealed a decrease in the expression of a number of genes, including a known apoptosis inhibitor, BIRC3, whose gene and protein expression level was also decreased by Cd treatment. Although the BIRC family consists of 8 members, Cd suppressed only BIRC3 gene expression. BIRC3 is known to suppress apoptosis through the inhibition effect on caspase-3. Knockdown of BIRC3 by siRNA as well as Cd treatment increased the level of active caspase-3. Moreover, knockdown of BIRC3 not only triggered cell toxicity and apoptosis but also strengthened Cd toxicity in HK-2 cells. Meanwhile, the activation of caspase-3 by suppression of BIRC3 gene expression was mostly specific to Cd and to proximal tubular cells. These results suggest that Cd induces apoptosis through the inhibition of ARNT-regulated BIRC3 in human proximal tubular cells.
Cadmium (Cd) is an environmental contaminant that induces toxic effects in various tissues including the kidney1–3. Cd accumulates in many organs, particularly in the kidney, because of its long biological half-life (10–30 years)1. Chronic Cd exposure arises mainly from dietary source and cigarette smoking and can cause nephrotoxicity, osteomalacia, teratogenicity and reproductive dysfunction4,5. In chronic dietary Cd exposure, the kidney is the target organ. Proximal tubular cell damage is characterized as Cd-induced renal damage1. Renal tubular cells take up Cd as a detoxified form bound to metallothionein (MT), and Cd is released from the bound form after lysosomal metabolism2. Although unbound Cd stimulates MT production in the kidney and then binds to MT, the toxic effects occur when unbound Cd accumulates in renal tubular cells2,6. At the molecular level, Cd induces apoptosis and also has adverse effects on cellular proliferation, cell signaling, and DNA repair in various cell lines7–10. Cd induces apoptotic cell death through endoplasmic reticulum (ER)-mediated, mitochondrial-mediated and p53-mediated pathways8. Cd impairs cell survival and proliferation through an increase in the phosphorylation of c-Jun NH2-terminal kinase (JNK) and p38 MAP kinase10. Cd inhibits the initial steps of base excision repair system and suppresses the capacity of the nucleotide excision system through interference with various enzymes9. Although Cd has been reported to have adverse effect on cellular functions, the precise mechanism of Cd-induced proximal tubular cell toxicity remains unclear.
To elucidate the precise mechanism of Cd-induced renal toxicity, we previously used the DNA microarray method to screen genes whose expression is changed by Cd treatment in mouse kidney and cultured cells, including proximal tubular cells11–13. Among the genes whose expression is downregulated by Cd treatment, the genes coding for proteins involved in the ubiquitin-proteasome system (UPS) are associated with Cd-induced renal toxicity14,15. Previous studies demonstrated an association between increased gene expression of apoptotic factors and Cd-induced apoptosis8,14,15. Other research groups revealed that the disruption of gene expression is involved in Cd-induced renal toxicity16–18. These results strongly imply that Cd exerts cytotoxicity through the disruption of gene expression.
The transcriptional regulation of eukaryotic genes involves the organized assembly of multi-protein complexes on promoter regions19,20. However, the upstream pathways that regulate gene transcription are controlled by specific regulatory mechanisms; furthermore, not all genes are induced at the same time and with the same duration. Some genes, such as those responsible for correct protein folding, are immediately induced for transcription within minutes; whereas others, such as those involved in DNA damage repair and cell metabolism, are slowly responded to upstream inductions signaling, within hours21. Once activated, transcription factors bind to gene regulatory elements (cis-elements), and through interactions with co-factors of the transcription machinery, promote access to DNA and facilitate the recruitment of the RNA polymerase enzymes to the transcriptional start site19,22.
Previous studies have shown that Cd exposure enhances the activities of transcription factors that induce cellular protection pathways. For instance, metal response element (MRE)-binding transcription factor-1 (MTF-1) is activated by Cd and induces transcription of MTs23–25. The Nrf2 transcription factor is also reported to play a role in Cd exposure-mediated gene expression of MT26. Although several transcription factors involved in MT expression have been identified, the transcription factors that control the expression of genes associated with Cd renal toxicity are poorly understood.
In this study, we screened transcription factors whose activities were changed by Cd treatment in HK-2 human proximal tubular cells. Among the transcription factors affected by Cd, we investigated the Cd-targeted transcription factors essential for Cd-induced renal toxicity. Furthermore, we examined the downstream factors of Cd-targeted transcription factors that are involved in Cd-induced renal toxicity.
To investigate Cd-induced cytotoxicity in HK-2 cells, we first examined cell viability of HK-2 cells treated with various concentrations of Cd using the MTT [3-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide] assay (Fig. 1a). HK-2 cells treated with 40µMCd for 6h showed 50% cell toxicity and exhibited less than 10% cell viability at 24h; however, at 3h, 40µMCd treatment did not induce cytotoxicity. Therefore, we selected these conditions, in which cells remain viable even with Cd treatment (40µM for 3h), to examine Cd activation-changed transcription factors using protein/DNA binding arrays in nuclear extracts27. Figure 1b shows confirmation and purity of our nuclear extraction process, in which the nuclear marker Lamin A/C is detected in nuclear fractions from both control and Cd-treated cells. Nuclear extracts from control and Cd-treated cells were then incubated with a mixture of 345 biotin-labeled DNA probes corresponding to transcription factor response elements (cis-elements). The quantity of cis-elements was determined using the protein/DNA binding array membrane on which elements that would hybridize with any cis-elements present were pre-spotted. Cd treatment affected the binding of several transcription factors to cis-elements compared with the control group (Fig. 1c). Overall, Cd increased the binding activities of 20 transcription factors by more than 2-fold (Table 1). Our results showed that binding activities of both p53 and MTF-1 transcription factors were increased in HK-2 cells in response to Cd treatment. We previously demonstrated that p53 protein level was dramatically increased by Cd treatment in vivo and in vitro 14,15. Moreover, another study demonstrated that the expression of apoptosis genes regulated by p53 is increased by Cd treatment in HK-2 cells14. A previous study showed that MTF-1 is induced to bind the MRE by Cd23–25. Our results are consistent with these findings. Cd also decreased the binding activities of 28 transcription factors, including PPAR (peroxisome proliferator activated receptor alpha) and Sp-1, by less than 0.5-fold (Table 2). Consistent with our results, several previous studies reported that PPAR and Sp-1 activities were decreased by Cd28,29. These studies help validate our findings and indicate our results may provide valuable information about the transcription factors involved in Cd toxicity or protection against Cd toxicity.
Our previous studies demonstrated that the suppression of gene expression is associated with Cd-induced renal toxicity14,15,30,31. These results imply that interruption in the regulation of gene expression is a key event in Cd toxicity. Therefore, we searched for a transcription factor that is involved in Cd toxicity, focusing on the transcription factors whose activities were decreased by Cd. In order to find the target transcription factor involved in Cd toxicity, it was determined whether the inhibition of expression of a transcription factor might affect the cell viability. As a result, each knockdown of six transcription factors triggered cell toxicity in HK-2 cells (Supplementary Table 1). Especially, the knockdown of ARNT [aryl hydrocarbon receptor nuclear translocator; known as hypoxia-inducible factor (HIF)-1β] by siRNA (Fig. 2a) conferred significant cell toxicity in HK-2 cells (Supplementary Table 1; Fig. 2b). EMSA assay showed that Cd treatment reduced the binding activity of ARNT (Fig. 2c). Knockdown of ARNT increased Cd toxicity in HK-2 cells (Fig. 2d), suggesting that decrease in the transcription activity of ARNT may strengthen Cd toxicity. In addition, although Cd did not affect the mRNA levels of ARNT (Fig. 2e), ARNT protein levels were decreased by Cd treatment in HK-2 cells (Fig. 2f). Together, this suggests that the ARNT transcription factor is a target in Cd-induced renal toxicity.
We next examined which downstream factor of ARNT may be involved in Cd toxicity. Because ARNT binding activity was decreased by Cd and the knockdown of ARNT by siRNA decreased cell viability, we performed DNA microarray analysis to determine the gene expressions that were decreased in ARNT knockdown HK-2 cells. The results showed that the expressions of 27 genes were decreased by less than 0.5 fold in ARNT knockdown cells (Table 3). Interestingly, real-time RT-PCR showed that the gene expression of BIRC3 [baculoviral IAP (inhibitor of apoptosis protein) repeat containing 3; known as cIAP2], an apoptosis inhibitor, was decreased by both ARNT siRNA as well as Cd treatment (Fig. 3a,b). Moreover, the protein level of BIRC3 was drastically decreased by Cd treatment in HK-cells (Fig. 3c). To investigate whether the Cd-induced decrease in BIRC3 gene expression was mediated via ARNT, BIRC3 mRNA levels were examined in ARNT knockdown HK-2 cells treated with Cd (Fig. 3d). BIRC3 mRNA levels were decreased by Cd treatment in both control and ARNT knockdown cells. However, the significant decrease in BIRC3 gene expression by ARNT knockdown without Cd was vanished upon Cd treatment. To determine the significance, we performed statistical analysis (Supplementary Table 2). Even with the F(0.95) of siRNA*Cd as 3.89, the F value of 6.92 is within the rejection region. Moreover, the P value of siRNA*Cd is below 0.05. Therefore, the siRNA effect and Cd treatment effect is significantly dependent. These results suggest that Cd treatment reduces cellular BIRC3 level through the suppression of ARNT transcription activity.
We next examined the impact of BIRC3 knockdown on cell viability using siRNA transfection. The mRNA and protein levels of BIRC3 were decreased by siRNA in a dose-dependent manner (Fig. 4a,b). Moreover, BIRC3 siRNA transfection significantly decreased HK-2 cell viability compared to control siRNA transfection cells (Fig. 4c). BIRC3 is a member of the BIRC family that is also known as the IAP family32–34. Eight family members have been identified in human, including BIRC1/NAIP, BIRC2/cIAP1, BIRC3/cIAP2, BIRC4/XIAP, BIRC5/Survivin, BIRC6/Apollon, BIRC7/ML-IAP and BIRC8/ILP32–34. To determine whether BIRC3 is specifically involved in Cd renal toxicity, we next examined the effect of Cd treatment on the mRNA levels of all BIRC family members (Fig. 5). Although the mRNA level of BIRC4 was increased by Cd treatment, those of the other BIRC family members were unaffected by Cd treatment in HK-2 cells. These results suggest that specific reduction of BIRC3 levels by Cd-induced suppression of ARNT activity is involved in Cd renal toxicity.
Previous studies showed that Cd induces apoptosis in proximal tubular cells8,14,15, and that BIRC3 inhibits apoptosis through the activity of caspases35. Therefore, we next examined whether BIRC3 is involved in Cd-induced apoptosis. Consistent with published studies, Cd treatment increased the level of cleaved caspase-3, the active form of casapse-3, in HK-2 cells (Fig. 6a). Moreover, knockdown of BIRC3 also increased the level of cleaved caspase-3 in HK-2 cells (Fig. 6b). These results suggest that knockdown of BIRC3 may induce apoptosis. Therefore, we performed TUNEL assays and found that knockdown of BIRC3 also induced apoptosis in HK-2 cells (Fig. 6c). Furthermore, knockdown of BIRC3 significantly increased Cd toxicity in HK-2 cells (Fig. 6d). Together, this demonstrates that a decrease in BIRC3 is involved in Cd-induced apoptosis in HK-2 cells.
In addition to Cd, other various metal(loid) toxicants also induce apoptosis in cultured cells. Methylmercury and inorganic mercury induce apoptosis in neuroblastoma and kidney cells, respectively36–38. The arsenic compound also causes apoptosis in hepatic cells39,40. To determine the specificity of BIRC3 function, we next examined the changes in BIRC3 gene expression and cleaved caspase-3 levels in several cell lines treated with metal(loid)s. Under the condition that methylmercury decreased the viability of human neuroblastoma (IMR-32 cells) by 20%, methylmercury increased cleaved caspase-3 levels; on the other hand, mRNA level of BIRC3 was not changed by methylmercury treatment (Fig. 7a–c). In HK-2 cells, methylmercury decreased cell viability by 10% as well as increased cleaved caspase-3 levels, but had no effect on BIRC3 mRNA levels (Fig. 7d–f). Inorganic mercury decreased BIRC3 mRNA levels in HK-2 cells even when the viability was almost the same as the control group; on the other hand, although inorganic mercury treatment reduced viability of HK-2 cells to 60% compared to control, cleaved caspase-3 levels were unchanged (Fig. 7g–i). In mouse hepatic cells (AML-12 cells), arsenic decreased the cell viability by 30%, with a significant increase in the mRNA level of Birc3 (Fig. 7j,l). Western blot analysis showed the protein levels of caspase-3 and cleaved caspase-3 were slightly increased by arsenic treatment (Fig. 7k). Although 6h of Cd treatment (10 to 30µM) decreased AML-12 cell viability to approximately 30%, only 10µMCd treatment decreased Birc3 mRNA levels (Fig. 7m,o). No cleaved caspase-3 levels were detected in AML-12 cells treated with Cd (Fig. 7n). As the western blot analysis did not clearly show the band of cleaved caspase-3 protein, we confirmed the analysis using staurosporin, an apoptosis inducer. Staurosporin drastically increased the protein level of cleaved caspase-3 in AML-12 cells (Supplementary Fig. 1). These data suggest that the activation of caspase-3 by suppression of BIRC3 gene expression by Cd treatment may mainly occur in proximal tubular cells.
Here we have found that Cd induces apoptosis through the quantitative alleviation of apoptosis inhibitor, BIRC3, through the suppression of its transcription in human proximal tubular cells. Recent studies have reported that chronic Cd exposure can induce apoptosis in renal cells8,41–46, and Cd induces via an ER-mediated pathway and mitochondrial-mediated pathway. In porcine renal proximal tubular epithelial LLC-PK1 cells, Cd induces ER stress and causes the activation of the unfolded protein response (UPR)-dependent apoptotic pathway, such as the IRE1-XBP1-JNK pathway45,46. In rat proximal tubule WKPT-0293 Cl.2 cells and mouse renal mesangial cells, Cd stimulates the release of pro-apoptotic factors from mitochondria41,44. Cd induces the swelling of mitochondria and subsequent cytochrome c release42,43. Our recent studies demonstrated that Cd induces apoptosis through p53 overaccumulation in human and rat proximal tubular cells13–15. In p53-mediated apoptosis, Cd inhibits the degradation of p53 by suppression of the gene expression of the UBE2D family, which is an E2 family enzyme in the UPS13–15. In the present study, we suggest a new apoptotic pathway involved in Cd toxicity. To the best of our knowledge, the ARNT-BIRC3 pathway is the first elucidated mechanism involved in Cd-induced apoptosis in proximal tubular cells (Fig. 8).
Our findings demonstrate that Cd suppresses the binding activity of the ARNT transcription factor in regulating BIRC3 expression. Our previous reports revealed that transcription factors FOXF1 and YY1 are involved in the pathway by which Cd decreases the expression of genes coding for UBE2D proteins14,27. Therefore, together these results suggest that Cd induces apoptosis through the inhibition of anti-apoptosis proteins as well as the promotion of pro-apoptosis proteins. This also suggests that the initiation pathway responsible for Cd-induced apoptosis is the inhibition of transcription activity. Many studies have examined Cd toxicity, however only a few studies have reported the transcript pathway involved in Cd toxicity. A recent study reported that phosphorylation of the transcription factor FOXO3 promotes cell survival upon Cd treatment47. The upregulation of the transcription factor Snail by the activation of Notch1 signaling is reported to be involved in the Cd-induced decrease in cell-cell adhesion48. In this study, we propose a novel transcription factor ARNT in driving HK-2 cells to apoptotic cell death.
Our protein/DNA binding array analysis in this study revealed that the binding activities of 48 transcription factors were affected by Cd treatment. Our previous study determined that Cd decreased the activity of the FOXF1 transcription factor, which suppressed the downstream factor UBE2D4, leading to apoptosis through p53 overaccumulation14. Our results identified FOXF1 as one of the transcription factors whose activities were decreased by Cd treatment (Table 2). In Supplementary table 1, the list of transcription factors whose gene knockdown affects cell viability is shown. In addition to ARNT and FOXF1, such transcription factors as GATAs and MEF2A may play essential roles in the pathway of Cd toxicity. Moreover, an increase in DNA binding activity of the apoptotic-related transcription factor p53, which was accumulated in proximal tubular cells by Cd, was detected in our protein/DNA binding array upon Cd treatment (Table 1). Thus, the results from our protein/DNA binding array can propose useful and valid information to help elucidate transcription factors involved in Cd toxicity.
ARNT is a member of the basic helix-loop-helix/Per-ARNT-Sim family. Previous studies showed that ARNT heterodimerizes with the aryl hydrocarbon receptor (AhR) or oxygen sensitive alpha subunit (HIF-1α or HIF-2α)49–51. AhR is a ligand-binding protein that functions as the response factor to environmental pollutant exposure, including ligands such as benzo[a]pyrene and 2,3,7,8-tetrachlorodibenzo-p-dioxin52,53. The non-ligand bound AhR binds instead to two heat shock proteins (XAP2 and HSP90) and the cochaperone p23 in the cytosol52. Ligand binding disrupts the complex and causes AhR to translocate into the nucleus. After heterodimerization with ARNT, AhR/ARNT binds to xenobiotic response elements (XREs) or dioxin responsive elements (DREs) in the promoter sequences of target genes52. In normoxia, HIF-1α and HIF-2α are hydroxylated and rapidly degraded by the UPS50,54. In hypoxia, HIF-1α and HIF-2α are stabilized and bind to ARNT to regulate the expression of downstream target genes50,54. HIF-mediated pathways are essential cellular responses to hypoxia such as angiogenesis, erythropoiesis, cell growth, and cell differentiation54–57. Nonetheless, previous studies suggested that independent of its role in AhR and HIF signaling, ARNT has new regulatory functions in the expression of cyclooxygenase-2, 12(S)-lipoxygenase, and p21 genes under normoxia conditions58–60. In addition, ARNT is associated with the proliferation and survival of tumor cell lines by regulating cellular processes49,58. These observations, including our findings, indicate that ARNT activity is not only induced by such signals as AhR and HIF pathway but is also suppressed by signals apart from the AhR or HIF pathway. In this study, HIF-1 activity was decreased by Cd treatment (Table 2); therefore, Cd-mediated suppression of ARNT activity may involve the AhR and HIF pathway. Further studies are required to elucidate whether the AhR or HIF pathway may be associated with Cd-suppressed ARNT activity.
Apoptosis is a highly programmed pathway of cell death that is typically achieved through the activation of caspases35. During death receptor- or mitochondrial-mediated apoptosis, initiator caspases, such as caspase-8/9 and -10, are recruited via binding proteins FADD, cytochrome c and Apaf-161,62. After recruitment of initiator caspases, effector caspases such as caspase-3 and -7 are activated by initiator caspases following cleavage into large and small subunits63,64. Cells have evolved important mechanisms to regulate caspase activity, for example, using the BIRC family. BIRC family members possess one or more baculorvirus IAP repeat (BIR) domains that selectively inhibits the activity of caspase-9, -3, or -732–34. BIRC4, also known as XIAP (X-linked IAP), is the most characterized member of the family35. BIRC4 blocks apoptosis by inhibiting caspases, using the interaction its BIR2 and BIR3 regions with active-site pocket of caspases65. Based on the mechanism of BIRC4, most BIRC family members were suggested to neutralize caspase activities in the same manner. However, recent studies have reported that BIRC family members are functionally non-equivalent and regulate caspase activities via distinct mechanisms66–68. BIRC2, BIRC3 and BIRC4 contain a C-terminal RING zinc finger domain with E3 ubiquitin ligase activity that mediates proteasomal degradation of cellular targets as well as themselves69. BIRC2 was demonstrated to interact with caspase-7 independently on the active site pocket66. BIRC2 ubiquitinates active caspase-3 and -7 and mediates their proteasomal degradation, thereby suppressing apoptosis67. In this study, the decrease in BIRC3 protein level increased the level of the active form of caspase-3. This suggests that BIRC3 may be directly involved in degradation of the active form of caspase-3 rather than interrupting the active site of caspase-3 in HK-2 cells. Further studies are required for elucidating the precise mechanism underlying BIRC3-mediated caspase-3 deactivation upon Cd treatment.
Our most striking observation in this study is our novel finding, to the best of our knowledge, that the ARNT-BIRC3 pathway is involved in Cd-induced apoptosis in proximal tubular cells. The regulation of BIRC3 gene expression has been previously unknown; however, our study suggests that ARNT may be a key transcription factor in BIRC3 regulation. Finally, we provide valuable information about the critical transcription factors for elucidating the toxic and defense mechanism pathways in response to Cd.
Human proximal tubular cells (HK-2 cells) were purchased from ATCC (Manassas, MA, USA), and cultured in Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F-12 Ham (DMEM/F-12) (Sigma-Aldrich, St. Louis, MO, USA), supplemented with 10% fetal bovine serum (FBS) (Gibco, Grand Island, NY, USA), 25U/mL penicillin (DS Pharm, Osaka, Japan), 25µg/mL streptomycin (DS Pharm), 1% Insulin-Transferrin-Selenium-X (Gibco), 10 ng/mL EGF (epidermal growth factor; Sigma-Aldrich), and 5 ng/mL hydrocortisone at 37°C in a humidified incubator containing 5% CO2.
Human neuroblastoma cells (IMR-32 cells) were kindly provided from Dr. Katsuhiko Muraki (Laboratory of Cellular Pharmacology, School of Pharmacy, Aichi Gakuin University, Japan). IMR-32 cells were cultured in DMEM supplemented with 10% FBS, 25U/mL penicillin, 25µg/mL streptomycin, and 1% MEM Non-essential Amino Acid Solution (Sigma-Aldrich) at 37°C in a humidified incubator containing 5% CO2.
Mouse normal hepatocytes (AML12 cells) were purchased ATCC, and cultured in DMEM/F-12, supplemented with 10% FBS, 1mM sodium pyruvate (Gibco), 25U/mL penicillin, 25µg/mL streptomycin, 1% Insulin-Transferrin-Selenium-X, and 10 ng/mL EGF at 37°C in a humidified incubator containing 5% CO2.
HK-2 cells and AML-12 cells were grown in plates at a density of 250 cells/mm2 and cultured for 48h. IMR-32 cells were grown in plates at a density of 500 cells/mm2 and cultured for 48h. The culture medium was discarded and the cells were treated with Cd (CdCl2; Wako Pure Chemical Industries, Osaka, Japan), methylmercury (CH3HgCl; GL Sciences Inc., Tokyo, Japan), inorganic mercury (HgCl2; Wako Pure Chemical Industries), or arsenic (NaAsO2; Wako Pure Chemical Industries) in serum-free culture medium for various times.
HK-2 and AML-12 cells were grown in 96-well plates and cultured for 48h. After treatment, culture medium was replaced with fresh 10% FBS-DMEM/F-12 containing MTT [3-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide; DOJINDO Laboratories, Kumamoto, Japan] and incubated for another 4h at 37°C. After removing the medium, 100μL dimethyl sulfoxide (Wako Pure Chemical Industries) was added to MTT formazan. Absorbance at 570nm was measured by the DTX880 multimode detector (Beckman Coulter Inc., Brea, CA, USA).
IMR-32 cells were grown in 96-well plates and cultured for 48h. After treatment, 10% (w/v) Alamar Blue (Invitrogen, Grand Island, NY, USA) was added and incubated for 4h at 37°C. Fluorescence was measured at excitation wavelength of 540nm and an emission wavelength of 595nm by the DTX880 multimode detector.
After treatment, cells were washed twice with ice-cold phosphate-buffered saline (PBS) (Nissui Pharmaceutical, Tokyo, Japan) and harvested in RIPA buffer [25mM Tris (pH 7.6), 150mM NaCl, 1% NP-40, 1% sodium deoxycholate 0.1% SDS; Thermo Fisher Scientific, Waltham, MA, USA]. Protein concentrations were measured using the BCA protein assay kit (Thermo Fisher Scientific). Protein samples were separated on SDS-polyacrylamide gels and transferred to a polyvinylidene fluoride membrane. The membrane was probed with primary antibodies and subsequently probed with horseradish peroxidase-conjugating (HRP) secondary antibodies (1:10000; GE Healthcare, Little Chalfont, UK). Proteins were detected by enhanced chemiluminescence using ImmunoStar® Zeta (Wako Pure Chemical Industries). The chemiluminescence images were taken using the LAS-500 or LAS-4000 (GE Healthcare) device. Primary antibodies were purchased as follows: GAPDH (1:2000) from American Research Products (Waltham, MA, USA); Lamin A/C (1:1000), ARNT (1:1000), BIRC3 (1:1000), caspase-3 (1:1000) and cleaved caspase-3 (1:1000) from Cell Signaling Technology (Danvers, MA, USA); and β-actin (1:1000) from Sigma-Aldrich.
Nuclei were extracted with the Nuclear Extraction Kit (Panomics; Affymetrix, Santa Clara, CA, USA). After Cd treatment, HK-2 cells were washed twice with ice-cold PBS and lysed on ice for 10min in extraction buffer A including a protease inhibitor, phosphatase inhibitor, and dithiothreitol (DTT). The cells were harvested from the assay plates by scraping and pipetting up and down several times to disrupt the cell clumps. Nuclei were collected by centrifugation at 14,000×g for 3min at 4°C. The pellet was resuspended in extraction buffer B including a protease inhibitor, phosphatase inhibitor, and DTT and incubated at 4°C for 1h. The mixture was then centrifuged at 14,000×g for 5min at 4°C, and the supernatant was collected. Protein concentration was measured using the BCA protein assay kit.
HK-2 cells were grown in 60-mm dishes at a density of 250 cells/mm2 and cultured for 48h. Culture medium was discarded and the cells were treated with 40µMCd in serum-free culture medium for 3h. HK-2 cells were separated into nuclear and post-nuclear fractions. The protein/DNA binding array was performed using the Combo Protein/DNA Array (Affymetrix)27. In brief, 20μg of nuclear extracts were mixed with a biotin-labeled probe mix, and the mixture was incubated at 15°C for 30min. The protein-bound probes in the mixture were isolated from the non-bound probes using a spin column. The protein-bound probes were eluted with column elution buffer and denatured at 95°C for 3min. The eluted probes were then added to the hybridization buffer and hybridized to the array membrane spotted with 345 consensus sequences complementary to the probes at 42°C overnight. The membrane was washed twice in 2×saline sodium citrate (SSC)/0.5% sodium dodecyl sulfate (SDS) at 42°C for 20min and then twice in 0.1×SSC/0.5% SDS at 42°C for 20min. The membrane was then blocked with 1×blocking buffer. The biotin-labeled probes were detected with streptavidin-horseradish peroxidase diluted 1:10000. The image was acquired using an LAS-4000 device. Spot density was evaluated using ImageQuantTL software (GE Healthcare).
The gel shift assay was performed using the EMSA kit purchased from Panomics (Affymetrix). HK-2 cells were grown in 60-mm dishes at a density of 250 cells/mm2 and cultured for 48h. After treatment, HK-2 cells were separated into nuclear and post-nuclear fractions. Nuclear protein (3µg) was incubated with 10 ng DNA probe (biotin-labeled binding sequence to transcription factor) and 1µg poly d(I-C) with binding buffer for 30min at 15°C in a thermal cycler (Takara Bio, Shiga, Japan). For the competition assay, 1,320 ng cold DNA probe was added. The protein-bound probe was electrophoresed on a 5.0% (w/v) TBE (Tris borate EDTA)-polyacrylamide gel in 0.5×TBE buffer at 4°C and then transferred to a Biodyne® B nylon membrane (Pall Corporation, Port Washington, NY, USA) in 0.5×TBE buffer. The membrane was fixed by UV crosslinking (CL-1000 Ultraviolet Crosslinker; UVP, Upland, CA, USA) with 120 mJ/cm2. The membrane was blocked and probed with Streptavidin-HRP. The chemiluminescence images were taken using a LAS-3000 device.
Silencer Select Pre-designed siRNAs were purchased from Ambion (Grand Island, NY, USA) as follows: s1613 and s1615 (Silencer® Select Pre-designed siRNA) for human ARNT; and s1451, s1452 and s1453 (Silencer® Select Pre-designed siRNA) for human BIRC3. Control siRNA (Silencer® Select Negative Control #1 siRNA) was also purchased from Ambion. siRNA transfection was performed using Lipofectamine RNAiMAX (Invitrogen). After the siRNA mixture was incubated for 15min with Lipofectamine RNAiMAX and Opti-MEM® I Reduced Serum Medium (Opti-MEM; Gibco), HK-2 cells were transfected with the siRNA mixture (1nM siRNA/sequence, 0.2% Lipofectamine RNAiMAX, 10% Opti-MEM).
Cells were washed twice with ice-cold PBS and total RNA was extracted with the PureLinkTM RNA Mini Kit (Ambion, Grand Island, NY, USA) and QuickGene-810 (Fujifilm). RNA quantification and purity was measured using BioSpec-nano (Shimadzu, Kyoto, Japan).
DNA microarray analysis was performed using previously described methods11. In brief, purified total RNA (5μg) was applied to an OpArray Human V4.0 slide with 35,035 genes (Operon Technologies, Alameda, CA, USA). The Low RNA Input Fluorescent Linear Amplification Kit (Agilent Technologies, Santa Clara, CA, USA) was used to synthesize complementary RNA (cRNA) using double-stranded cDNA as a template. A primer containing a poly dT and a T7 polymerase promoter was annealed to poly A+RNA. Reverse transcriptase was then added to the reaction mix and double-stranded cDNA was transcribed from the untreated or siRNA-treated cells in the presence of cyanine (Cy) 3-CTP or Cy5-CTP (PerkinElmer, Waltham, MA, USA), respectively. These two sets of fluorescence-labeled cRNA were mixed and hybridized to an Operon microarray slide for 16h at 42°C using a Lucidea SlidePro Hybridizer (Amersham Bioscience, Little Chalfont, UK). Fluorescent images were recorded with CRBIO (Hitachi Software Engineering, Tokyo, Japan). Digitized image data were processed with DNASIS Array software (Hitachi Software Engineering). Following global normalization, the data were filtered to exclude genes with low-level expression. The intensity of the Cy5 (siRNA-treated) to Cy3 (control) ratio was calculated. Information on each gene was obtained from the National Center for Biotechnology Information (NCBI) database.
Total RNA was incubated with the PrimeScript reverse transcription (RT) Reagent Kit (Perfect Real Time) (Takara Bio) to generate cDNA. Real-time PCR was performed with SYBR Premix Ex TaqII (Perfect Real Time) (Takara Bio) and the Thermal Cycler Dice Real Time System (Takara Bio). PCR conditions were as follows: 10s hot-start at 95°C followed by 40 cycles of 5s at 95°C and 30s at 60°C. Gene expression was normalized to GAPDH or ß-actin mRNA levels. The oligonucleotide sequences of the primers were as follows: sense, 5′-CTAGTGGCCATTGGCAGATT-3′, and antisense, 5′-CAATGTTGTGTCGGGAGATG-3′, for the human ARNT gene; sense,5′-CATGTGTGTGGAGGGTGAAG-3′, and antisense, 5′-TTTAACAGGGGACAGCATCC-3′, for the human BIRC1 gene; sense, 5′-GCATTTTCCCAACTGTCCAT-3′, and antisense, 5′-ATTCGAGCTGCATGTGTCTG-3′, for the human BIRC2 gene; sense, 5′-CAACAGATCTGGCAAAAGCA-3′, and antisense, 5′-TTGCTCAATTTTCCACCACA-3′, for the human BIRC3 gene; sense, 5′-TGGGGTTCAGTTTCAAGGAC-3′, and antisense, 5′-TGCAACCAGAACCTCAAGTG-3′, for the human BIRC4 gene; sense, 5′-GTTGCGCTTTCCTTTCTGTC-3′, and antisense, 5′-TCTCCGCAGTTTCCTCAAAT-3′, for the human BIRC5 gene; sense, 5′-TGACGCTTTCAACCTCACTG-3′, and antisense, 5′-GTGTCCGCTGGACCAGTTAT-3′, for the human BIRC6 gene; sense, 5′-TGGCCTCCTTCTATGACTGG-3′, and antisense, 5′-ACCTCACCTTGTCCTGATGG-3′, for the human BIRC7 gene; sense, 5′-AAGCCCGGCTCATTACTTTT-3′, and antisense, 5′-ATCTTCCTTGGGCTTCCAGT-3′, for the human BIRC8 gene; sense, 5′- GCACCGTCAAGGCTGAGAAC-3′, and antisense, 5′-TGGTGAAGACGCCAGTGGA-3′, for the human GAPDH gene; sense, 5′-TGACGTGTGTGACACCAATG-3′, and antisense, 5′-TGCTGCAGTGTTTCCTTTTG-3′, for the mouse Birc3 gene; sense, 5′-CCTAAGGCCAACCGTGAAAA-3′, and antisense, 5′-AGCCATACAGGGACAGCACA-3′, for the mouse ß-actin gene.
HK-2 cells were transferred to a Millicell EZ SLIDE (Millipore, Billerica, MA, USA) at a density of 250 cells/mm2 with the siRNA mixture (1nM siRNA/sequence, 0.2% Lipofectamine RNAiMAX, 10% Opti-MEM) and incubated for 48h. Apoptotic cells were detected using the In Situ Cell Death Detection Kit, Fluorescein (Roche, Mannheim, Germany) according to the manufacturer’s protocol. Fluorescence images were taken using a BIOREVO BZ-9000 microscope (Keyence, Osaka, Japan).
Statistical analyses were performed using one or two-way analysis of variance (ANOVA). When the F value was significant (P<0.05), Bonferroni’s multiple t-test was performed for post-hoc comparison (P<0.05).
We thank sincerely Mrs. H. Hasegawa, K. Asai, T. Ieda, H. Shinkai, and Y. Noda for their excellent experimental support. This work was supported by The Study of the Health Effects of Heavy Metals, Organized by the Ministry of the Environment, Japan.
J.Y.L. designed and conducted experiments, analyzed results, and wrote the manuscript. M.T. conducted Protein/DNA binding array experiment, analyzed results and revised the manuscript. G.W.H. and M.Y.L. revised the manuscript. M.S. designed this project, analyzed results, and revised the manuscript.
The authors declare that they have no competing interests.
Electronic supplementary material
Supplementary information accompanies this paper at 10.1038/s41598-017-17494-9.
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