- Open Access
Inflammation contributes to NKX3.1 loss and augments DNA damage but does not alter the DNA damage response via increased SIRT1 expression
© Debelec-Butuner et al.; licensee BioMed Central. 2015
- Received: 4 April 2014
- Accepted: 29 January 2015
- Published: 15 February 2015
The oxidative stress response is a cellular defense mechanism that protects cells from oxidative damage and cancer development. The exact molecular mechanism by which reactive oxygen species (ROS) contribute to DNA damage and increase genome instability in prostate cancer merits further investigation. Here, we aimed to determine the effects of NKX3.1 loss on antioxidant defense in response to acute and chronic inflammation in an in vitro model. Oxidative stress-induced DNA damage resulted in increased H2AX(S139) phosphorylation (a hallmark of DNA damage), along with the degradation of the androgen receptor (AR), p53 and NKX3.1, upon treatment with conditioned medium (CM) obtained from activated macrophages or H2O2. Furthermore, the expression and stability of SIRT1 were increased by CM treatment but not by H2O2 treatment, although the level of ATM(S1981) phosphorylation was not changed compared with controls. Moreover, the deregulated antioxidant response resulted in upregulation of the pro-oxidant QSCN6 and the antioxidant GPX2 and downregulation of the antioxidant GPX3 after CM treatment. Consistently, the intracellular ROS level increased after chronic treatment, leading to a dose-dependent increase in the ability of LNCaP cells to tolerate oxidative damage. These data suggest that the inflammatory microenvironment is a major factor contributing to DNA damage and the deregulation of the oxidative stress response, which may be the underlying cause of the increased genetic heterogeneity during prostate tumor progression.
- Inflammatory microenvironment
- Prostate tumor
- DNA damage
Oxidative stress contributes to the initiation, promotion and progression of carcinogenesis. Excessive levels of reactive oxygen species (ROS) are generated by exposure to oxidative stress and cause sustained DNA damage. DNA damage occurs during the initiation step of carcinogenesis, and most likely results in abnormal gene expression. Because ROS accumulation results in a subsequent failure of signal transduction at the promotion step, the cells undergo a loss of genomic fidelity during the progression step . Notably, ROS are generated in excess amounts during chronic inflammation, and ROS-mediated DNA damage alters the genetic composition. This damage may promote oncogenic transformation, which occurs when genes encoding essential factors involved in DNA repair, apoptosis and cell cycle regulation are affected . Others have shown that ROS contribute to carcinogenesis by activating signaling pathways that regulate cellular proliferation, angiogenesis and metastasis [3-5]. ROS serve as secondary messengers for the activation of key transcription factors in response to pro-inflammatory cytokines, and they also regulate the transcription of genes involved in inflammatory responses [6,7]. Additionally, a number of protein kinase pathways, such as the MAPK pathway, are activated by oxidative signals during inflammation and inflammatory diseases. These pathways synergistically contribute to the activation of cytokine release, and combined with the loss of adhesion and the release of angiogenic factors, they may eventually contribute to cellular proliferation, differentiation and tumor progression . ROS play important roles in multiple signal transduction pathways, such as those mediated by TNFα  and p53. These two pathways can activate each other, as DNA damage caused by TNFα-induced ROS directly induces cell cycle and/or apoptosis regulation by p53 . In addition to the ability of cells to trigger proliferation in response to sustained ROS production at low levels, excess ROS generation or the accumulation of ROS may induce cell death depending on the ROS concentration and cell type . Although ROS can induce apoptosis or necrosis depending on the oxidative level , the appropriate cellular responses to ROS production are critical events and may protect the cell from sustained oxidative damage and support cell survival.
To reduce the oxidative damage caused by ROS, the antioxidant response system is activated through several constitutive and inducible detoxification mechanisms, including the expression of enzymes such as glutathione peroxidases (GPx), catalase, superoxide dismutases (SODs), peroxiredoxins (PRDXs), glutathione-S-transferases (GSTs), NADP (H) quinone oxidoreductase (NQO1), epoxide hydrolase, heme oxygenase (HO-1), UDP-glucuronosyl transferases (UGTs), and gamma-glutamylcysteine synthetase. The upregulated expression of these enzymes exhibits a distinct cellular defense to protect cells from oxidative damage and cancer development [7,10].
In addition to the mechanisms that regulate ROS, cell proliferation is controlled by stress-sensing molecules such as Sirtuin 1 (SIRT1), an NAD-dependent deacetylase that responds to the levels of redox pairs NAD+/NADH and NADP+/NADPH. Under oxidative conditions, SIRT1 deacetylates a number of transcription factors, including p53, NBS1 and FOXO, which subsequently contribute to cellular metabolic responses, such as cell cycle regulation and DNA damage [7,11,12]. Therefore, SIRT1 has been linked to tumor cell survival by deregulating apoptosis and promoting senescence. Particularly, in prostate cancer, the deacetylation of AR by SIRT1 represses androgen-induced AR transcription and contributes to AR-induced tumorigenesis .
NKX3.1 is an androgen-regulated gene that encodes a homeobox protein with a tumor suppressor function in prostate cells [14,15]. The AR response is ubiquitous in prostate tumors, and NKX3.1 is upregulated by androgens; in contrast, NKX3.1 loss has been reported in prostate tumors . Furthermore, the functional loss of NKX3.1 expression upon cytokine exposure has been reported in previous studies of the inflammatory microenvironment [17,18], strengthening its tumor suppressor role in prostate carcinogenesis. The pro-inflammatory cytokines TNFα and IL-1β induce the C-terminal phosphorylation of NKX3.1 by casein kinase 2 (CK2), resulting in a shortened half-life [17,18]. Additionally, loss of NKX3.1 expression in pathogen E. coli infected prostate lobes in mice has been shown to be correlated with reduced AR expression . It was previously reported that the loss of NKX3.1 expression was related not only to the loss of AR transactivating function [17,18] but also to high ROS level upon cytokine exposure, particularly TNFα . Concurrently, the loss of p53 expression was also observed in the inflammatory microenvironment, promoted the progression of prostate cancer, perhaps correlating with increased oxidative stress. This effect was partially restored by suppressing AKT and MDM2 phosphorylations, leading to p53 degradation .
In this study, we aimed to identify the role of cytokine-induced NKX3.1 loss in the deregulation of the antioxidant defense during acute and chronic exposure to both cytokines and ROS. Therefore, the effect of antioxidant treatment on the inflammation- and/or oxidative stress-induced degradation of NKX3.1, AR and p53 was analyzed. Cultures of the prostate cancer cell line LNCaP were exposed to conditioned medium (CM) with adjusted amounts of pro-inflammatory cytokines (TNFα) for 24 h for acute treatment and for 2 weeks for chronic treatment. Cells were also chronically treated with H2O2 for 2 weeks to compare the effects of pro-inflammatory cytokines and ROS exposure.
Macrophage differentiation and conditioned media (CM) collection
The U937 monocyte cell line was cultured in RPMI 1640 medium including 10% FBS (fetal bovine serum) at 37°C with 5% CO2. To achieve macrophage differentiation and cytokine production, cells (8×105) were seeded into 75-cm2 culture flasks 2 h prior to treatment. Next, PMA was added at a final concentration of 16 nM for 16 h, and the adherent clusters (differentiated monocytes) were maintained. The cells were washed twice before the addition of 20 ml of fresh medium, and the cells were then allowed to rest for 3 h. Then, lipopolysaccharide (LPS) was added at a final concentration of 10 ng/ml to induce cytokine secretion. The cells were incubated for an additional 24 h, and the supernatant (conditioned medium - CM) was collected and filtered (using a 0.22-μm filter) for further use. To ensure that the CM was cell-free, diluted CM was cultured in an empty flask (25 cm2) for one week and analyzed.
Measurement of cytokines in CM
Before feeding the LNCaP cells with the collected CM (cell-free), the TNFα (Invitrogen, USA) levels were analyzed using an ELISA according to the manufacturer’s instructions. Because cytokine exposure is a major component of the inflammatory microenvironment, the times (0, 2, 4, 6, 12 and 24 h) and doses (62.5, 125, 250 or 500 pg/ml TNFα-containing conditioned medium) for the courses of CM treatments were optimized as reported in our previous study . TNFα was chosen as a measure of the CM concentration, which was adjusted by diluting the CM with normal medium before application to the LNCaP cells. As a result, the concentrations of macrophage-secreted cytokines were adjusted and maintained at picogram levels. In our studies, the effective concentration of TNFα was 400 times less than the concentration of recombinant TNFα (rTNFα) (sigma, UK) reported in other studies [17-19].
Cell culture and treatments
LNCaP cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA) and were propagated as recommended using RPMI 1640 medium supplemented with 10% FBS, L-glutamine (2 mM), penicillin (100 U/ml) and streptomycin (100 μg/ml) at 37°C with 5% CO2. For the acute exposures, the CM (62, 125, and 250 pg/ml of TNFα) treatments were performed for 24 h; for the chronic exposures, the CM treatments continued for 2 weeks, and lower doses (50 and 100 pg/ml of TNFα) were used. TNFα concentrations were adjusted by diluting the CM using RPMI 1640 medium as described previously . A chronic oxidative condition was also induced by treating the cells with 50, 100 or 200 μM H2O2 for 2 weeks for comparison of the effects of cytokine exposure and oxidative stress.
The NKX3.1 open reading frame was amplified (using the primers F: GGATCCATGCTCAGGGTTCCGGAGCCG and R: GAATTCGGTTGTCACCTGAGCTGGCATTA) and cloned into the pcDNA4/HisMax-TOPO vector (Invitrogen, USA) according to the manufacturer’s instructions to obtain HM-NKX3.1 and the HM-vector constructs. Then, transfections were performed using the Fugene HD reagent (Roche, Germany) for 24 h. The cells were incubated for an additional 18 to 42 h, as appropriate.
The siNKX3.1 and siAR transfections were performed as recommended by the supplier (Dharmacon). Briefly, 4×105 cells were seeded into 6-cm plates, and the medium was changed (w/o antibiotics). A transfection mix was prepared by adding 6 μl of Dharmafect II (tube 1) and 200 pmol of siNKX3.1, siAR or scrambled siRNA (tube 2) into 94 μl of transfection medium (w/o antibiotics and serum). After incubation for 5 min at RT, the tubes were mixed and incubated for 15 min at RT and then added onto the cells dropwise. The transfected cells were incubated for an additional 24 h before harvesting.
The following antibodies were purchased from the manufacturers: AR (Millipore, USA), p53, pH2AX(S139) and pATM(S1981) (Abcam, UK), GAPDH (Ambion, UK), β-actin and SIRT1 (Sigma, UK), Caspase-3 (R&D, UK), and β-tubulin (ABM, UK). The NKX3.1 custom antibody was a gift from Prof. Dr. F. Saatcioglu (University of Oslo). The HRP-conjugated anti-mouse and anti-rabbit (Amersham, UK) and the AlexaFluor 488- and 594-conjugated secondary antibodies (Invitrogen, USA) were purchased and used as recommended by the manufacturers.
LNCaP cells (8×103) were seeded into 96-well plates, and the transfections were carried out on the following day. Two days later, the cells were incubated with DCFH-DA (2′ 7′- dichlorodihydrofluorescein diacetate, Molecular Probes, 10 μM) for 30 min at 37°C. Next, the treatments were performed following gentle washes using phenol red-free medium. Finally, the fluorescence intensity was measured every 20 min for up to 3 h using a Fluoroscan fluorometer (Thermo Science, USA).
Protein extraction and western blotting
For protein extraction, LNCaP cells were lysed using a modified RIPA buffer (10 mM Tris-Cl (pH: 8.0), 1% Triton X-100, 0.1% SDS, 0.1% Na deoxycholate, 1 mM EDTA, 1 mM EGTA, 140 mM NaCl) containing protease and phosphatase inhibitors. Then, the concentrations were determined using the BCA assay (Sigma, UK). SDS-PAGE and western blots were performed under standard conditions with 50 μg of protein lysate per lane. The proteins were separated on 10-12% gels and transferred to PVDF membranes (Amersham, UK) using a wet transfer blotter. The PVDF membrane was blocked with 5% dry milk in TBS-T (Tris-Buffered-Saline solution containing 0.1% Tween 20). The primary and secondary antibody incubations were performed in TBS-T containing 0.5% dry milk or 5% BSA at RT for 1 h or at 4°C o/n. The membranes were developed using the ECL prime reagent (Amersham, UK) for 5 min and were photographed using Kodak X-Ray films in a dark room.
Real-time cell proliferation assay
The Xcelligence proliferation assay platform was used for real-time measurements. Briefly, the LNCaP cells (8×103) were transfected with an HM vector and HM-NKX3.1 (24 h), seeded into 96-well plates (E-plates, Roche GmbH, Germany) and cultured for 24 h. The treatments were performed as described, and the proliferation rate and morphological changes were monitored. Impedance values were collected every 10 min for 48 h.
Total RNA was isolated from the LNCaP cells using the RNeasy kit (Qiagen, CA, USA), and the yield was calculated using absorbance readings at 260/280 nm. Then, cDNA synthesis was performed using a cDNA synthesis kit (Invitrogen, USA) as recommended by the manufacturer.
Real time PCR
To study the expression of specific genes, quantitative RT-PCR was performed using a SYBR Green PCR kit and the LC480 PCR system (Roche, Germany). The relative abundance of each transcript was calculated using the comparative cycle threshold (CT) method with GAPDH as an invariant control. The following primers were used: GPX2_F: CAGTCTCAAGTATGTCCGT, GPX2_R: AGGCTCAATGTTGATGGT; GPX3_F: CTTGCACCATTCGGTCT, GPX3_R: CGGACATACTTGAGGGTAG; PRDX6_F: TAGTGTGATGGTCCTTCCAAC, PRDX6_R: AGCGGAGGTATTTCTTGC; QSCN6_F: GAGGCTACGTGCACTACT, QSCN6_R: CTGCAAGGCGAGCATTGA; ENOX2_F: CTGAACGTGAAGCACTG, ENOX2_R: ATCAAGACGGTGCAAGTAG; SOD1_F: TGTACCAGTGCAGGTCC, SOD1_R: GCCAATGATGCAATGGTC; SOD2_F: TGTCCAAGGCTCAGGTT, SOD2_R: CTGAAGGTAGTAAGCGTGC; NKX3.1_F: TCTATCAGCATCTGACAGGTGAA, NKX3.1_R: AGCAGGGTTTGTTATGCATGTAG; SIRT1_F: TGCGGGAATCCAAAGGATAATTCAGTGTC, SIRT1_R: CTTCATCTTTGTCATACTTCATGGCTCTATG; and GAPDH_F: CATTGCCCTCAACGACCACTTT, GAPDH_R: GGTGGTCCAGGGGTCTTACTCC.
Student’s t test was applied to determine the statistical significance between pairs where necessary.
NKX3.1, AR, and p53 degradation is restored by LNAC treatment
NKX3.1 is required for antioxidant gene expression to limit oxidative damage
SIRT1 leads to oxidative stress resistance in the inflammatory microenvironment
The accumulation of damage to DNA, proteins and lipids is characterized by an increase in intracellular oxidative stress levels due to a progressive decrease of ROS scavenging [18,20]. Several lines of evidence indicate that the loss of oxidative tolerance is age dependent and associated with DNA damage as well as metabolic deregulation . The SIRT1-mediated increase in oxidative stress tolerance and the concurrent activation of the p53-mediated DNA damage response were correlated with an extended lifespan in mouse models. A decrease in p53 activity reduces the essential role of p53 in tumor prevention in older animals . Thus, a dramatic increase in the frequency of cancer provides a likely explanation for the correlation between tumorigenesis and the accumulation of DNA mutations [23,24] that might be related to a decreased stress tolerance and increased genetic heterogeneity during multiple inflammatory exposures over a lifetime.
Furthermore, because the SIRT1 level was increased in the cells treated with CM, we elucidated the DNA damage response activation by examining the p-ATM(S1981) level. Interestingly, we found that ATM(S1981) phosphorylation was not significantly increased upon CM treatment. LNAC only induced p-ATM(S1981) in control cells and not after CM exposure, suggesting that the reduced oxidative stress levels might trigger a cell cycle progression and DNA damage response, presumably via SIRT1-mediated p53 and NBS1 activations respectively. However, this hypothesis requires further investigation. We also examined whether apoptosis was induced in response to DNA damage, and found that the ATM-mediated response upon CM exposure did not generate caspase-3 cleavage in LNCaP cells, either with or without LNAC treatment (Figure 4B). These data demonstrate that apoptosis was not activated, whereas the cells continued to proliferate with damaged DNA, resulting in genetic heterogeneity. However, this proliferation was suppressed by the metabolic activity sensor and NAD+-dependent deacetylase SIRT1, which slows down the cell cycle through the deacetylation of functional p53 to allow time for DNA damage repair. This process requires the presence of functional NKX3.1 in prostate cells [28,29]. This mechanism might result in oxidative stress tolerance and is commonly observed in cancer progression.
Loss of NKX3.1 under conditions of chronic inflammation leads to an abrogated antioxidant response
Chronic inflammation with sustained oxidative stress is well known to promote carcinogenesis . To mimic chronic inflammation in vitro, we fed LNCaP cells with CM (50 and 100 pg/ml of TNFα) or normal media or H2O2 (50, 100 and 200 μM) for a 2-week period. First, we analyzed the proteasomal degradation of NKX3.1, which correlated with exposure to increasing concentrations of CM (Figure 4C) but not with H2O2 treatments, confirming that cytokines are the major cause of NKX3.1 degradation. Secondly, we investigated the expression of antioxidant genes and detected the upregulation of the pro-oxidant QSCN6 (3.8-fold), marginal upregulation of the antioxidant GPX2 (increase 1.3-fold) and downregulation of GPX3 (2.5-fold) after CM (100 pg/ml of TNFα) treatment (Figure 4D). These alterations may be due to the chronic exposure to inflammatory cytokines, which might be sufficient to maintain cell survival. Surprisingly, treatment of cells with H2O2 did not result in significant changes in the expression of antioxidant response factors (data not shown). We also measured the intracellular ROS levels after the chronic treatments to gain insight into the changes in the oxidative conditions within the cells. Chronic CM exposure dose dependently correlated with an increase in the ROS level, but chronic H2O2 exposure maintained the ROS level close to the basal concentration (Figure 4E). These data suggest that inflammatory cytokine release is the major factor underlying the deregulated antioxidant response and sustained oxidative damage in prostate cells. Additionally, we found that NKX3.1 loss caused by CM exposure inversely correlated with an increased ROS level. When the NKX3.1 level remained stable after H2O2 exposure, there was no change in the ROS level. Thus, these data suggest that the increased ROS concentration after chronic CM exposure might be a consequence of NKX3.1 loss.
NKX3.1 suppresses the proliferation enhanced by the inflammatory microenvironment
This research was supported by a grant for project 113S044 (COST action EU-ROS) from TUBITAK to KSK.
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