Cellular Signalling 27 (2015) 1713–1719
Contents lists available at ScienceDirect
Cellular Signalling journal homepage: www.elsevier.com/locate/cellsig
Diarachidonoylphosphoethanolamine induces necrosis/necroptosis of malignant pleural mesothelioma cells Yoshiko Kaku a,1, Ayako Tsuchiya a,1, Takeshi Kanno a, Takashi Nakano b, Tomoyuki Nishizaki a,⁎ a b
Division of Bioinformation, Department of Physiology, Hyogo College of Medicine, 1–1 Mukogawa-cho, Nishinomiya 663–8501, Japan Division of Respiratory Medicine, Department of Internal Medicine, Hyogo College of Medicine, Nishinomiya 663–8501, Japan
a r t i c l e
i n f o
Article history: Received 6 January 2015 Received in revised form 24 April 2015 Accepted 5 May 2015 Available online 22 May 2015 Keywords: Diarachidonoylphosphoethanolamine Malignant pleural mesothelioma cell Necrosis Necroptosis Mitochondria
a b s t r a c t The present study investigated 1,2-diarachidonoyl-sn-glycero-3-phosphoethanolamine (DAPE)-induced cell death in malignant pleural mesothelioma (MPM) cells. DAPE reduced cell viability in NCIH28, NCI-H2052, NCI-H2452, and MSTO-211H MPM cell lines in a concentration (1–100 μM)-dependent manner. In the flow cytometry using propidium iodide (PI) and annexin V (AV), DAPE significantly increased the population of PI-positive and AV-negative cells, corresponding to primary necrosis, and that of PI-positive and AV-positive cells, corresponding to late apoptosis/secondary necrosis, in NCIH28 cells. DAPE-induced reduction of NCI-H28 cell viability was partially inhibited by necrostatin-1, an inhibitor of RIP1 kinase to induce necroptosis, or knocking-down RIP1. DAPE generated reactive oxygen species (ROS) followed by disruption of mitochondrial membrane potentials in NCI-H28 cells. DAPEinduced mitochondrial damage was attenuated by cyclosporin A, an inhibitor of cyclophilin D (CypD). DAPE did not affect expression and mitochondrial localization of p53 protein in NCI-H28 cells. DAPE significantly decreased intracellular ATP concentrations in NCI-H28 cells. Overall, the results of the present study indicate that DAPE induces necroptosis and necrosis of MPM cells; the former is mediated by RIP1 kinase and the latter is caused by generating ROS and opening CypD-dependent mitochondrial permeability transition pore, to reduce intracellular ATP concentrations. © 2015 Elsevier Inc. All rights reserved.
1. Introduction Cell death is classified into three types: apoptosis, necrosis, and necroptosis. A recent highlight has focused upon necroptosis, an alternative form of programmed cell death [1]. Death receptors such as tumor necrosis factor α (TNF-α) receptor and Fas or proapoptotic Bcl-2 family members related to mitochondrial damage
Abbreviations: DAPE, 1,2-diarachidonoyl-sn-glycero-3-phosphoethanolamine; MPM, malignant pleural mesothelioma; PI, propidium iodide; AV, annexin V; Nec-1, necrostatin-1; ROS, reactive oxygen species; MPT, mitochondrial permeability transition; CypD, cyclophilin D; CsA, cyclosporine A; DPPE, 1,2dipalmitoleoyl-sn-glycero-3-phosphoethanolamine; PP2A, protein phosphatase 2A; PTP1B, protein tyrosine phosphatase 1B; DLPE, 1,2-dilinoleoyl-sn-glycero-3phosphoethanolamine; DOPE, 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine; TNF-α, tumor necrosis factor α; RIP1, receptor interacting protein 1; FADD, Fas-associated death domain; tBid, truncated Bid; FBS, fetal bovine serum; MTT, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide; PBS, phosphate buffered saline; carboxy-H2DCFDA, 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate; RT-PCR, reverse transcription-polymerase chain reaction; SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis; HRP, horseradish peroxidase; NAC, N-acetylL-cysteine. ⁎ Corresponding author. Tel.: +81 798 45 6397; fax: +81 798 45 6649. E-mail address: [email protected] (T. Nishizaki). 1 These two authors contributed equally to this work.
http://dx.doi.org/10.1016/j.cellsig.2015.05.007 0898-6568/© 2015 Elsevier Inc. All rights reserved.
mediate both in apoptosis and necroptosis. Receptor interacting protein 1 (RIP1) associated with death receptors forms complex IIa including Fas-associated death domain (FADD) and caspase-8, causing activation of caspase-8 and the effector caspase-3, to induce apoptosis [2]. Caspase-8, alternatively, proteolyzes Bid into truncated Bid (tBid), which makes pores in the mitochondria, allowing release of apoptosis-related factors such as cytochrome c, thereby activating caspase-9 and the effector caspase-3, to induce mitochondrial apoptosis [3]. RIP1 is phosphorylated by RIP1 kinase, thereby forming complex IIb together with RIP3, to induce necroptosis [2]. Apoptotic stimuli such as oxidative stress cause pro-apoptotic Bcl-2 family member-mediated damages of mitochondria, to release cytochrome c followed by activation of caspase-9 and caspase-3, to induce mitochondrial apoptosis [4]. Mitochondrial damage, alternatively, activates ATPase and reduces intracellular ATP concentrations, to induce necrosis [5]. Accumulating evidence has pointed to the role of reactive oxygen species (ROS) in cell death. ROS disrupts integrity of the mitochondrial inner membrane, to open the mitochondrial permeability transition (MPT) pore, resulting in reduced ATP production, responsible for necrosis [6]. Cyclophilin D (CypD) is a component of the MPT pore, and cyclosporine A (CsA) inhibits CypD.
1714
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Phosphatidylethanolamine is a membrane phospholipid and includes huge kinds, depending upon a combination of free fatty acids on the α and β position. Accumulating evidence has pointed to a wide variety of roles for phosphatidylethanolamine such as membrane fusion, cell cycle, autophagy, apoptosis, and cognitive function [7–11]. In our earlier study, 1,2-dipalmitoleoyl-sn-glycero-3-phosphoethanolamine (DPPE), which contains palmitoleic acid on the α and β position, induced apoptosis of malignant pleural mesothelioma (MPM) cells by enhancing activities of protein phosphatase 2A (PP2A) and protein tyrosine phosphatase 1B (PTP1B) [12]. Another phosphatidylethanolamine 1,2dilinoleoyl-sn-glycero-3-phosphoethanolamine (DLPE), which contains linoleic acid on the α and β position, also enhanced PP2A and PTP1B activities, but such effect was not obtained with 1,2-diarachidonoylsn-glycero-3-phosphoethanolamine (DAPE), which contains arachidonic acid on the α and β position, or 1,2-dioleoyl-sn-glycero-3phosphoethanolamine (DOPE), which contains oleic acid on the α and β position [13]. The present study was conducted to understand whether and how DAPE induces MPM cell death. We show here that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells and that DAPE also induces necrosis by producing ROS, to open CypD-dependent MPT pore and disrupt mitochondrial membrane potentials, thereby reducing intracellular ATP concentrations. 2. Materials and methods 2.1. Cell culture NCI-H28, NCI-H2052, NCI-H2452, and MSTO-211H cell lines were purchased from American Type Culture Collection (Manassas, VA, USA). Cells were grown in RPMI-1640 medium supplemented with 10% (v/v) heat-inactivated fetal bovine serum (FBS), 0.003% (w/v) L-glutamine, penicillin (final concentration, 100 U/ml), and streptomycin (final concentration, 0.1 mg/ml), in a humidified atmosphere of 5% CO2 and 95% air at 37 °C. 2.2. Assay of cell viability Cell viability was evaluated by the method using 3-(4,5-dimethyl-2thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) as previously described [11]. 2.3. Construction and transfection of siRNA The siRNA silencing the RIP1-targeted gene (RIP1 siRNA) and the negative control siRNA (NC siRNA) were obtained from Ambion (Carlsbad, CA, USA). The sequences of RIP1 siRNA used here were CCACUAGU CUGACGGAUAAtt and UUAUCCGUCAGACUAGUGGta. The NC siRNA contained the scrambled sequence with the same GC content and nucleic acid composition. RIP1 siRNA and NC siRNA were reversetransfected into cells using a Lipofectamine reagent (Invitrogen, Carlsbad, CA, USA). Cells were used for experiments 48 h after transfection. Fig. 1. DAPE-induced MPM cell death. (A) Cell viability. MPM cells were treated with DAPE at concentrations as indicated for 24 h, and cell viability was quantified with an MTT assay. Data represent the mean (± SEM) percentage of basal levels (MTT intensities in untreated cells)(n = 4 independent experiments for cells except for NCI-H2452 cell line; n = 8 independent experiments for NCI-H2452 cell line). (B) Flow cytometry using PI and annexin V-FITC (AV). NCI-H28 cells were untreated (Control) and treated with DAPE (30 μM) for 48 h. Typical profiles are shown in the upper panel. In the graphs, each column represents the mean (± SEM) percentage of cells in 4 fractions against total cells (n = 4 independent experiments). P values, unpaired t-test. NS, not significant. (C) Cell cycle analysis. NCI-H28 cells were untreated (Control) and treated with DAPE (30 μM) for 24 h. Typical profiles are shown in the upper panel. In the graphs, each column represents the mean (± SEM) percentage for each phase of cell cycling (n = 4 independent experiments). P values, unpaired t-test. NS, not significant.
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
1715
with a flow cytometer (FACSCalibur) at an excitation of 488 nm and an emission of 585 nm, and analyzed using a BD CellQuest Pro software (Becton Dickinson). 2.6. Monitoring of ROS production Cells were incubated in the culture medium containing 10 μM 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate (carboxyH 2 DCFDA) (Life Technologies, Tokyo, Japan) at 37 °C for 30 min. ROS generation was monitored at a 488 nm argon laser with a confocal scanning laser microscope (LSM 510, Carl Zeiss Co., Ltd., Oberkochen, Germany). 2.7. Monitoring of mitochondrial membrane potentials
Fig. 2. DAPE-induced necroptosis of NCI-H28 cells. Cells were treated with DAPE (100 μM) in the presence and absence of Nec-1 (10 μM) for 24 h, and cell viability was quantified with an MTT assay. Data represent the mean (± SEM) percentage of basal levels (MTT intensities in untreated cells)(n = 4 independent experiments). P value, Dunnett's test.
Cells were incubated in a DePsipherTM solution at 37 °C for 20 min, and the fluorescent signals were observed with a confocal scanning laser microscope (LSM 510, Carl Zeiss Co., Ltd.) at 543 nm helium–neon laser through a long-pass 560 nm filter for red aggregations and at 488 nm argon laser through a band-pass 505–530 nm filter for green monomeric form. 2.8. Real-time reverse transcription-polymerase chain reaction (RT-PCR)
2.4. Flow cytometry for apoptosis analysis Cells were suspended in a binding buffer and stained with both propidium iodide (PI) and annexin V (AV)-FITC and loaded on a flow cytometer (FACSCalibur, Becton Dickinson, Franklin Lakes, USA) available for FL1 (AV) and FL2 (PI) bivariate analyisis. Data from 20,000 cells/sample were collected, and the quadrants were set according to the population of viable, unstained cells in untreated samples. CellQuest analysis of the data was used to calculate the percentage of the cells in the respective quadrants.
2.5. Flow cytometry for cell cycle analysis Cells were harvested by a trypsinization, fixed with 70% (v/v) ethanol at 4 °C overnight. Fixed cells were incubated in phosphate buffered saline (PBS) containing 1.5 μg/ml RNase A for 1 h at 37 °C, followed by staining with 5 μg/ml of PI for 20 min on ice. Then, cells were collected on a nylon mesh filter (pore size, 40 μm), and cell cycles were assayed
Total RNAs from cells were purified by an acid/guanidine/thiocyanate/chloroform extraction method using the Sepasol-RNA I Super kit (Nacalai, Kyoto, Japan). After purification, total RNAs were treated with RNase-free DNase I (2 units) at 37 °C for 30 min to remove genomic DNAs, and 10 μg of RNAs was resuspended in water. Then, random primers, dNTP, 10 × RT buffer, and Multiscribe Reverse Transcriptase were added to an RNA solution and incubated at 25 °C for 10 min followed by 37 °C for 120 min to synthesize the first-strand cDNA. Real-time RT-PCR was performed using a SYBR Green Realtime PCR Master Mix (Takara Bio, Otsu, Japan) and the Applied Biosystems 7900 real-time PCR detection system (ABI, Foster City, CA). Thermal cycling conditions were as follows: first step, 94 °C for 4 min; the ensuing 40 cycles, 94 °C for 1 s, 65 °C for 15 s, and 72 °C for 30 s. The primers used here were as follows: sense, 5′-GCCATCTACAAGCAGTCACAGCAC AT-3′ and anti-sense, 5′-GGCACAAACACGCACCTCAAAGC-3′ for p53; and sense, 5′-GACTTCAACAGCGACACCCACTCC-3′ and anti-sense, 5′-AGGTCC ACCACCCTGTTGCTGTAG-3′ for GAPDH. Expression of the p53 mRNA was normalized by that of GAPDH mRNA.
Fig. 3. DAPE-induced NCI-H28 cell death is partially suppressed by knockingdown RIP1. (A) NCI-H28 cells were transfected with NC siRNA or RIP1 siRNA, and 48 h later, Western blotting was carried out. The signal intensity for RIP1 was normalized by that for β-actin. In the graphs, each column represents the mean (± SEM) normalized intensity (n = 4 independent experiments). P value, unpaired t-test. (B) Cells, transfected with NC siRNA or RIP1 siRNA, were treated with DAPE at concentrations as indicated for 24 h, followed by MTT assay. In the graph, each column represents the mean (± SEM) percentage of basal levels (MTT intensities in NC siRNA-transfected cells without DAPE treatment)(n = 4 independent experiments). P value, unpaired t-test.
1716
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 4. DAPE-induced ROS production in NCI-H28 cells. Cells, treated with DAPE (100 μM) in the presence and absence of NAC (2 mM) for periods of time as indicated, were reacted with carboxy-H2DCFDA, and the fluorescent intensity was measured. Typical fluorescent images (FIs) are shown in the left panel. DIC, differential interference contrast. Bars, 50 μm. We chose a visual field with almost equivalent cell numbers in the DIC images for analysis. In the graphs, each point represents the mean (± SEM) signal intensity in total cells (n = 4 independent experiments).
2.9. Separation into mitochondrial and cytosolic components Cells were homogenized with a sonicator in a Buffer A solution (20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, and 250 mM sucrose, pH 7.5). Lysates were centrifuged at 1000 g for 10 min, and the supernatant was further centrifuged at 10,000 g for 1 h. The pellet and the supernatant were used as mitochondrial and cytosolic components, respectively.
2.10. Western blotting Samples were loaded on 10% (v/v) sodium dodecyl sulfate (SDS)–polyacrylamide gel electrophoresis (PAGE) and transferred to polyvinylidene difluoride membrane. After blocking with TBST [20 mM Tris, 150 mM NaCl, 0.1 % (v/v) Tween-20, pH 7.5] containing 5% (w/v) of bovine serum albumin, blotting membrane was reacted with antibodies against p53 (Cell Signaling Technology, Inc., Danvers, MA, USA) or VDAC1 (Santa Cruz Biotechnology, Inc., Dallas, Texas, USA), followed by a horseradish peroxidase (HRP)conjugated anti-mouse IgG antibody. For β-actin detection, blotting membrane was reacted with an anti-β-actin antibody (Sigma, Missouri, SL, USA) followed by an HRP-conjugated antimouse IgG antibody. Immunoreactivity was detected with an ECL kit (Invitrogen) and visualized using a chemiluminescence detection system (GE Healthcare, Piscataway, NJ, USA). Protein
concentrations for each sample were determined with a BCA protein assay kit (Thermo Fisher Scientific, Rockford, IL, USA). 2.11. Monitoring of intracellular ATP levels Intracellular ATP levels were assayed using a luminescent ATP detection assay kit containing luciferase and D-luciferin (Wako Pure Chemical Industries, Osaka, Japan). ATP can be relatively quantified by detecting the fluorescence signal emitted in luciferase-mediated reaction of D-luciferin with ATP into D-oxyluciferin. Before and after treatment with DAPE, cells were incubated in the ATP detection kit for 1 min by shaking, and 10 min later under the dark conditions, the emitted light was detected with a microplate reader (ARVO X4; PerkinElmer, Inc., Waltham, MA, USA). 2.12. Statistical analysis The data presented were mean ± standard error of the mean (SEM). Statistical differences were analyzed by Dunnett's test and unpaired t-test. Values of p b 0.05 were considered statistically significant. 3. Results 3.1. DAPE induces MPM cell death Our initial attempt was to assess the effect of DAPE on cell viability for MPM cell lines. DAPE reduced cell viability for all the investigated
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
1717
Fig. 5. DAPE-induced disruption of mitochondrial membrane potentials in NCI-H28 cells. (A) Mitochondrial membrane potentials were monitored in cells treated with DAPE (100 μM) for periods of time as indicated. (B) Cells were preincubated in the presence and absence of CsA (10 μM) for 1 h, and mitochondrial membrane potentials were monitored 20 min after treatment with and without DAPE (100 μM) for 20 min. Typical red and green fluorescent images were shown in each upper panel. DIC, differential interference contrast. Bars, 50 μm. We chose a visual field with almost equivalent cell numbers in the DIC images for analysis. In the graphs, each value represents the mean (± SEM) red or green signal intensity in total cells (n = 4 independent experiments). MP, membrane potential. P values, Dunnett's test.
MPM cell lines NCI-H28, NCI-H2052, NCI-H2452, and MSTO-211H in a concentration (1–100 μM)-dependent manner, reaching nearly 0% of basal levels at 100 μM (Fig. 1A). In the flow cytometry analysis using PI and AV, PI is a marker of dead cells and AV, which detects externalized phosphatidylserine residues, is a marker of apoptotic cells [14]. In this assay, the populations of PI-positive/AV-negative, PI-negative/AV-positive, and PIpositive/AV-positive cells correspond to primary necrosis, early apoptosis, and late apoptosis/secondary necrosis, respectively [15]. DAPE significantly increased the populations of PI-positive/AV-negative and PI-positive/AV-positive cells for NCI-H28 cell line (Fig. 1B). This indicates that DAPE induces both necrosis and apoptosis of NCI-H28 cells. In the cell cycle analysis, DAPE significantly increased the proportions at the S and G2/M phase of cell cycling in parallel with decreased population at the G1 phase in NCI-H28 cells (Fig. 1C). This implies that DAPE induces cell cycle arrest at the S and G2/M phase in NCI-H28 cells. DAPE-induced reduction of NCI-H28 cell viability was partially prevented by necrostatin-1 (Nec-1), an inhibitor of RIP1 kinase to induce necroptosis (Fig. 2). This suggests that DAPE induces necroptosis. To obtain further evidence for this, RIP1 was knocked down using RIP1 siRNA (Fig. 3A). DAPE (100 μM)-induced NCI-H28 cell death was
significantly suppressed by knocking down RIP1, although cell death induced by lower concentrations (10–70 μM) of DAPE was not affected (Fig. 3B). Collectively, these results suggest that DAPE induces MPM cell death at least in part due to necroptosis. 3.2. DAPE generates ROS and damages mitochondrial membrane potentials ROS are recognized to trigger necrosis and apoptosis. DAPE increased intracellular ROS levels in a treatment time (1–20 min)-dependent manner in NCI-H28 cells, and the increase was found from at 1-min treatment, reaching the maximum at 10-min treatment (Fig. 4). DAPEinduced ROS rise was abolished by N-acetyl-L-cysteine (NAC), a ROS scavenger (Fig. 4). These results indicate that DAPE generates ROS in NCI-H28 cells. We subsequently monitored mitochondrial membrane potentials in NCI-H28 cells using DePsipherTM. DePsipherTM, a mitochondrial activity marker, has the properties of aggregating upon membrane polarization forming an orange-red fluorescent compound. If the potential is disturbed, the dye has no access to the transmembrane space and remains in or reverts to its green monomeric form. DAPE increased green fluorescent signals in a treatment time (1–20 min)-dependent manner in parallel with decreased orange-red fluorescent signals (Fig. 5A),
1718
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 6. Effects of DAPE on p53 expression and intracellular localization in NCI-H28 cells. (A) Real-time RT-PCR was carried out in cells treated with DAPE (100 μM) for periods of time as indicated. Expression of the p53 mRNA was normalized by that of the GAPDH mRNA and the value before treatment with DAPE (0 h) was regarded as 1. In the graph, each point represents the mean (± SEM) relative expression of the p53 mRNA (n = 4 independent experiments). (B) Western blotting was performed in cells treated with DAPE (100 μM) for periods of time as indicated using an anti-p53 antibody. Expression of p53 protein was normalized by that of β-actin. In the graph, each point represents the mean (± SEM) normalized expression of p53 protein (n = 4 independent experiments). (C) Cells treated with DAPE (100 μM) for periods of time as indicated were separated into the cytosolic and mitochondrial fractions, and Western blotting was carried out in each fraction using antibodies against p53 and VDAC1. In the graph, each column represents the mean (± SEM) relative mitochondrial localization of p53 (the signal intensity for p53 in the mitochondrial fraction/that in the cytosolic and mitochondrial fractions)(n = 4 independent experiments).
indicating that DAPE perturbs mitochondrial membrane potentials in NCI-H28 cells. An increase in green fluorescent signals was found from at 3-min treatment with DAPE, reaching the maximum at 10-min treatment (Fig. 5A). This suggests that ROS production precedes mitochondrial damage. DAPE-induced mitochondrial damage was prevented by CsA, an inhibitor of CypD (Fig. 5B). This suggests that DAPE may initiate CypDdependent MPT, to disrupt mitochondrial membrane potentials. 3.3. DAPE reduces intracellular ATP levels, regardless of p53-mediated mitochondrial damage Evidence has shown that in response to oxidative stress, p53 accumulates in the mitochondria and opens the MPT pore, to induce necrosis [16]. In the real-time RT-PCR analysis, DAPE did not affect expression of the p53 mRNA in NCI-H28 cells, except for a slight increase at 1-h treatment (Fig. 6A). DAPE did not also affect basal expression levels of p53 protein (Fig. 6B). To examine subcellular distribution of p53 protein,
cells were separated into the mitochondrial and cytosolic components. The immunoreactive signals for VDAC1, a mitochondrial marker, were detected in the mitochondrial components, but no signal was found in the cytosolic components (Fig. 6C). This confirms that mitochondrial and cytosolic components are successfully separated. DAPE did not increase mitochondrial localization of p53 protein as compared with that before treatment; conversely, DAPE did not significantly decrease the mitochondrial localization (Fig. 6C). This indicates that no accumulation of p53 protein in the mitochondria is induced by DAPE. This also suggests no participation of p53 in DAPE-induced MPM cell death. Intracellular ATP decrease in association with mitochondrial damage is a key factor for necrosis [5]. We finally assayed intracellular ATP concentrations in NCI-H28 cells. DAPE significantly reduced intracellular ATP concentrations as compared with those before treatment (Fig. 7). Overall, these results indicate that DAPE reduces intracellular ATP levels through a p53-independent mitochondrial pathway.
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 7. DAPE-induced intracellular ATP reduction in NCI-H28 cells. Intracellular ATP was assayed in cells treated with DAPE (100 μM) for periods of time as indicated. The fluorescence signal for ATP before treatment with DAPE (0 h) was regarded as 1. In the graph, each point represents the mean (± SEM) relative ATP level (n = 4 independent experiments).
4. Discussion In the present study, DAPE reduced cell viability in all the investigated MPM cell lines in a concentration (1–100 μM)-dependent manner. DAPE significantly increased the populations of PI-positive/AV-negative and PI-positive/AV-positive NCI-H28 cells, which correspond to primary necrosis and late apoptosis/secondary necrosis, respectively. In the cell cycle analysis, DAPE increased the proportions at the S and G2/M phase of cell cycling in parallel with decreased the population at the G1 phase in NCI-H28 cells, indicating that DAPE arrests cell cycle at the S and G2/M phase in NCI-H28 cells. We have obtained the data that DAPE does not significantly activate caspase-3, 4, 8, and 9 in NCI-H28 cells (unpublished data). Taken together, these results indicate that DAPE induces both caspase-independent apoptosis and necrosis of MPM cells. The present study focused upon the mechanism underlying nonapoptotic MPM cell death induced by DAPE. DAPE-induced reduction of NCI-H28 cell viability was partially inhibited by the RIP1 kinase inhibitor Nec-1 or knocking-down RIP1. This suggests that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells. DAPE, on the other hand, increased intracellular ROS levels in NCIH28 cells, and the effect was abolished by the ROS scavenger NAC. This accounts for DAPE-induced ROS production. DAPE also perturbed mitochondrial membrane potentials in NCI-H28 cells. DAPE-induced ROS production and mitochondrial damage were found from at 1-min and 3-min treatment, respectively. This implies that DAPE produces ROS, which triggers to disrupt mitochondrial membrane potentials. In response to death stimuli, apoptosis and necrosis occur per continuum and coexist in the same cell [5]. Production of ROS, channelmediated calcium uptake, activation of non-apoptotic proteases, and/
1719
or enzymatic destruction of cofactors required for ATP production induce both apoptosis and necrosis in dying cells. ROS damages integrity of the mitochondrial inner membrane, allowing release of apoptosisrelated factors from the mitochondria, responsible for apoptosis, or causing loss of the ability to generate ATP, responsible for necrosis [6]. In the present study, CsA, an inhibitor of the MPT component CypD, prevented DAPE-induced perturbation of mitochondrial membrane potentials. This indicates that DAPE produces ROS, to open CypDdependent MPT pore and disrupt mitochondrial membrane potentials. Expectedly, DAPE apparently decreased intracellular ATP levels in NCIH28 cells, supporting the notion for DAPE-induced necrosis in association with mitochondrial damage. Oxidative stress is shown to cause accumulation of p53 in the mitochondria, to open the MPT pore [16]. DAPE, however, affected neither basal expression levels nor mitochondrial localization of p53 protein in NCI-H28 cells. This rules out the participation of p53 in DAPEinduced mitochondrial damage and cell death. Overall, these results indicate that DAPE produces ROS, which opens CypD-dependent and p53-independnet MPT pore, to disrupt mitochondrial membrane potentials followed by intracellular ATP reduction, and then leading to necrosis of NCI-H28 cells. In conclusion, the results of the present study show that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells and alternatively, necrosis by stimulating ROS production and initiating CypD-dependent MPT, to impair mitochondrial membrane potentials followed by intracellular ATP reduction. This may provide fresh insight into DAPE signaling relevant to necrosis/necroptosis. Statement of conflicts of interest The authors have no conflicts of interest. References [1] A. Degterev, Z. Huang, M. Boyce, Y. Li, P. Jagtap, N. Mizushima, G.D. Cuny, T.J. Mitchison, M.A. Moskowitz, J. Yuan, Nat. Chem. Biol. 1 (2005) 112–119. [2] D.E. Christofferson, J. Yuan, Curr. Opin. Cell Biol. 22 (2010) 263–268. [3] H. Li, H. Zhu, C.J. Xu, J. Yuan, Cell 94 (1998) 491–501. [4] M. Kvansakul, M.G. Hinds, Cell Death Dis. 4 (2013) (e909). [5] W.X. Zong, C.B. Thompson, Genes Dev. 20 (2006) 1–15. [6] E.J. Griffiths, A.P. Halestrap, Biochem. J. 307 (1995) 93–98. [7] F. Deeba, H.N. Tahseen, K.S. Sharad, N. Ahmad, S. Akhtar, M. Saleemuddin, O. Mohammad, Biochim. Biophys. Acta 1669 (2005) 170–181. [8] K. Emoto, T. Kobayashi, A. Yamaji, H. Aizawa, I. Yahara, K. Inoue, M. Umeda, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 12867–12872. [9] K. Emoto, N. Toyama-Sorimachi, H. Karasuyama, K. Inoue, M. Umeda, Exp. Cell Res. 232 (1997) 430–434. [10] Y. Ichimura, T. Kirisako, T. Takao, Y. Satomi, Y. Shimonishi, N. Ishihara, N. Mizushima, I. Tanida, E. Kominami, M. Ohsumi, T. Noda, Y. Ohsumi, Nature 408 (2000) 488–492. [11] T. Yaguchi, T. Nagata, T. Nishizaki, Behav. Brain Funct. 6 (2010) 52. [12] Y. Kaku, A. Tsuchiya, T. Kanno, T. Nakano, T. Nishizaki, Anticancer Res. 34 (2014) 1759–1764. [13] A. Tsuchiya, T. Kanno, T. Nishizaki, Cell. Physiol. Biochem. 34 (2014) 617–627. [14] D.M. Vanags, M.I. Pörn-Ares, S. Coppola, D.H. Burgess, S. Orrenius, J. Biol. Chem. 271 (1996) 31075–31085. [15] G. Pietra, R. Mortarini, G. Parmiani, A. Anichini, Cancer Res. 61 (2001) 8218–8226. [16] A.V. Vaseva, N.D. Marchenko, K. Ji, S.E. Tsirka, S. Holzmann, U.M. Moll, Cell 149 (2012) 1536–1548.
Contents lists available at ScienceDirect
Cellular Signalling journal homepage: www.elsevier.com/locate/cellsig
Diarachidonoylphosphoethanolamine induces necrosis/necroptosis of malignant pleural mesothelioma cells Yoshiko Kaku a,1, Ayako Tsuchiya a,1, Takeshi Kanno a, Takashi Nakano b, Tomoyuki Nishizaki a,⁎ a b
Division of Bioinformation, Department of Physiology, Hyogo College of Medicine, 1–1 Mukogawa-cho, Nishinomiya 663–8501, Japan Division of Respiratory Medicine, Department of Internal Medicine, Hyogo College of Medicine, Nishinomiya 663–8501, Japan
a r t i c l e
i n f o
Article history: Received 6 January 2015 Received in revised form 24 April 2015 Accepted 5 May 2015 Available online 22 May 2015 Keywords: Diarachidonoylphosphoethanolamine Malignant pleural mesothelioma cell Necrosis Necroptosis Mitochondria
a b s t r a c t The present study investigated 1,2-diarachidonoyl-sn-glycero-3-phosphoethanolamine (DAPE)-induced cell death in malignant pleural mesothelioma (MPM) cells. DAPE reduced cell viability in NCIH28, NCI-H2052, NCI-H2452, and MSTO-211H MPM cell lines in a concentration (1–100 μM)-dependent manner. In the flow cytometry using propidium iodide (PI) and annexin V (AV), DAPE significantly increased the population of PI-positive and AV-negative cells, corresponding to primary necrosis, and that of PI-positive and AV-positive cells, corresponding to late apoptosis/secondary necrosis, in NCIH28 cells. DAPE-induced reduction of NCI-H28 cell viability was partially inhibited by necrostatin-1, an inhibitor of RIP1 kinase to induce necroptosis, or knocking-down RIP1. DAPE generated reactive oxygen species (ROS) followed by disruption of mitochondrial membrane potentials in NCI-H28 cells. DAPEinduced mitochondrial damage was attenuated by cyclosporin A, an inhibitor of cyclophilin D (CypD). DAPE did not affect expression and mitochondrial localization of p53 protein in NCI-H28 cells. DAPE significantly decreased intracellular ATP concentrations in NCI-H28 cells. Overall, the results of the present study indicate that DAPE induces necroptosis and necrosis of MPM cells; the former is mediated by RIP1 kinase and the latter is caused by generating ROS and opening CypD-dependent mitochondrial permeability transition pore, to reduce intracellular ATP concentrations. © 2015 Elsevier Inc. All rights reserved.
1. Introduction Cell death is classified into three types: apoptosis, necrosis, and necroptosis. A recent highlight has focused upon necroptosis, an alternative form of programmed cell death [1]. Death receptors such as tumor necrosis factor α (TNF-α) receptor and Fas or proapoptotic Bcl-2 family members related to mitochondrial damage
Abbreviations: DAPE, 1,2-diarachidonoyl-sn-glycero-3-phosphoethanolamine; MPM, malignant pleural mesothelioma; PI, propidium iodide; AV, annexin V; Nec-1, necrostatin-1; ROS, reactive oxygen species; MPT, mitochondrial permeability transition; CypD, cyclophilin D; CsA, cyclosporine A; DPPE, 1,2dipalmitoleoyl-sn-glycero-3-phosphoethanolamine; PP2A, protein phosphatase 2A; PTP1B, protein tyrosine phosphatase 1B; DLPE, 1,2-dilinoleoyl-sn-glycero-3phosphoethanolamine; DOPE, 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine; TNF-α, tumor necrosis factor α; RIP1, receptor interacting protein 1; FADD, Fas-associated death domain; tBid, truncated Bid; FBS, fetal bovine serum; MTT, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide; PBS, phosphate buffered saline; carboxy-H2DCFDA, 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate; RT-PCR, reverse transcription-polymerase chain reaction; SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis; HRP, horseradish peroxidase; NAC, N-acetylL-cysteine. ⁎ Corresponding author. Tel.: +81 798 45 6397; fax: +81 798 45 6649. E-mail address: [email protected] (T. Nishizaki). 1 These two authors contributed equally to this work.
http://dx.doi.org/10.1016/j.cellsig.2015.05.007 0898-6568/© 2015 Elsevier Inc. All rights reserved.
mediate both in apoptosis and necroptosis. Receptor interacting protein 1 (RIP1) associated with death receptors forms complex IIa including Fas-associated death domain (FADD) and caspase-8, causing activation of caspase-8 and the effector caspase-3, to induce apoptosis [2]. Caspase-8, alternatively, proteolyzes Bid into truncated Bid (tBid), which makes pores in the mitochondria, allowing release of apoptosis-related factors such as cytochrome c, thereby activating caspase-9 and the effector caspase-3, to induce mitochondrial apoptosis [3]. RIP1 is phosphorylated by RIP1 kinase, thereby forming complex IIb together with RIP3, to induce necroptosis [2]. Apoptotic stimuli such as oxidative stress cause pro-apoptotic Bcl-2 family member-mediated damages of mitochondria, to release cytochrome c followed by activation of caspase-9 and caspase-3, to induce mitochondrial apoptosis [4]. Mitochondrial damage, alternatively, activates ATPase and reduces intracellular ATP concentrations, to induce necrosis [5]. Accumulating evidence has pointed to the role of reactive oxygen species (ROS) in cell death. ROS disrupts integrity of the mitochondrial inner membrane, to open the mitochondrial permeability transition (MPT) pore, resulting in reduced ATP production, responsible for necrosis [6]. Cyclophilin D (CypD) is a component of the MPT pore, and cyclosporine A (CsA) inhibits CypD.
1714
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Phosphatidylethanolamine is a membrane phospholipid and includes huge kinds, depending upon a combination of free fatty acids on the α and β position. Accumulating evidence has pointed to a wide variety of roles for phosphatidylethanolamine such as membrane fusion, cell cycle, autophagy, apoptosis, and cognitive function [7–11]. In our earlier study, 1,2-dipalmitoleoyl-sn-glycero-3-phosphoethanolamine (DPPE), which contains palmitoleic acid on the α and β position, induced apoptosis of malignant pleural mesothelioma (MPM) cells by enhancing activities of protein phosphatase 2A (PP2A) and protein tyrosine phosphatase 1B (PTP1B) [12]. Another phosphatidylethanolamine 1,2dilinoleoyl-sn-glycero-3-phosphoethanolamine (DLPE), which contains linoleic acid on the α and β position, also enhanced PP2A and PTP1B activities, but such effect was not obtained with 1,2-diarachidonoylsn-glycero-3-phosphoethanolamine (DAPE), which contains arachidonic acid on the α and β position, or 1,2-dioleoyl-sn-glycero-3phosphoethanolamine (DOPE), which contains oleic acid on the α and β position [13]. The present study was conducted to understand whether and how DAPE induces MPM cell death. We show here that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells and that DAPE also induces necrosis by producing ROS, to open CypD-dependent MPT pore and disrupt mitochondrial membrane potentials, thereby reducing intracellular ATP concentrations. 2. Materials and methods 2.1. Cell culture NCI-H28, NCI-H2052, NCI-H2452, and MSTO-211H cell lines were purchased from American Type Culture Collection (Manassas, VA, USA). Cells were grown in RPMI-1640 medium supplemented with 10% (v/v) heat-inactivated fetal bovine serum (FBS), 0.003% (w/v) L-glutamine, penicillin (final concentration, 100 U/ml), and streptomycin (final concentration, 0.1 mg/ml), in a humidified atmosphere of 5% CO2 and 95% air at 37 °C. 2.2. Assay of cell viability Cell viability was evaluated by the method using 3-(4,5-dimethyl-2thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) as previously described [11]. 2.3. Construction and transfection of siRNA The siRNA silencing the RIP1-targeted gene (RIP1 siRNA) and the negative control siRNA (NC siRNA) were obtained from Ambion (Carlsbad, CA, USA). The sequences of RIP1 siRNA used here were CCACUAGU CUGACGGAUAAtt and UUAUCCGUCAGACUAGUGGta. The NC siRNA contained the scrambled sequence with the same GC content and nucleic acid composition. RIP1 siRNA and NC siRNA were reversetransfected into cells using a Lipofectamine reagent (Invitrogen, Carlsbad, CA, USA). Cells were used for experiments 48 h after transfection. Fig. 1. DAPE-induced MPM cell death. (A) Cell viability. MPM cells were treated with DAPE at concentrations as indicated for 24 h, and cell viability was quantified with an MTT assay. Data represent the mean (± SEM) percentage of basal levels (MTT intensities in untreated cells)(n = 4 independent experiments for cells except for NCI-H2452 cell line; n = 8 independent experiments for NCI-H2452 cell line). (B) Flow cytometry using PI and annexin V-FITC (AV). NCI-H28 cells were untreated (Control) and treated with DAPE (30 μM) for 48 h. Typical profiles are shown in the upper panel. In the graphs, each column represents the mean (± SEM) percentage of cells in 4 fractions against total cells (n = 4 independent experiments). P values, unpaired t-test. NS, not significant. (C) Cell cycle analysis. NCI-H28 cells were untreated (Control) and treated with DAPE (30 μM) for 24 h. Typical profiles are shown in the upper panel. In the graphs, each column represents the mean (± SEM) percentage for each phase of cell cycling (n = 4 independent experiments). P values, unpaired t-test. NS, not significant.
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
1715
with a flow cytometer (FACSCalibur) at an excitation of 488 nm and an emission of 585 nm, and analyzed using a BD CellQuest Pro software (Becton Dickinson). 2.6. Monitoring of ROS production Cells were incubated in the culture medium containing 10 μM 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate (carboxyH 2 DCFDA) (Life Technologies, Tokyo, Japan) at 37 °C for 30 min. ROS generation was monitored at a 488 nm argon laser with a confocal scanning laser microscope (LSM 510, Carl Zeiss Co., Ltd., Oberkochen, Germany). 2.7. Monitoring of mitochondrial membrane potentials
Fig. 2. DAPE-induced necroptosis of NCI-H28 cells. Cells were treated with DAPE (100 μM) in the presence and absence of Nec-1 (10 μM) for 24 h, and cell viability was quantified with an MTT assay. Data represent the mean (± SEM) percentage of basal levels (MTT intensities in untreated cells)(n = 4 independent experiments). P value, Dunnett's test.
Cells were incubated in a DePsipherTM solution at 37 °C for 20 min, and the fluorescent signals were observed with a confocal scanning laser microscope (LSM 510, Carl Zeiss Co., Ltd.) at 543 nm helium–neon laser through a long-pass 560 nm filter for red aggregations and at 488 nm argon laser through a band-pass 505–530 nm filter for green monomeric form. 2.8. Real-time reverse transcription-polymerase chain reaction (RT-PCR)
2.4. Flow cytometry for apoptosis analysis Cells were suspended in a binding buffer and stained with both propidium iodide (PI) and annexin V (AV)-FITC and loaded on a flow cytometer (FACSCalibur, Becton Dickinson, Franklin Lakes, USA) available for FL1 (AV) and FL2 (PI) bivariate analyisis. Data from 20,000 cells/sample were collected, and the quadrants were set according to the population of viable, unstained cells in untreated samples. CellQuest analysis of the data was used to calculate the percentage of the cells in the respective quadrants.
2.5. Flow cytometry for cell cycle analysis Cells were harvested by a trypsinization, fixed with 70% (v/v) ethanol at 4 °C overnight. Fixed cells were incubated in phosphate buffered saline (PBS) containing 1.5 μg/ml RNase A for 1 h at 37 °C, followed by staining with 5 μg/ml of PI for 20 min on ice. Then, cells were collected on a nylon mesh filter (pore size, 40 μm), and cell cycles were assayed
Total RNAs from cells were purified by an acid/guanidine/thiocyanate/chloroform extraction method using the Sepasol-RNA I Super kit (Nacalai, Kyoto, Japan). After purification, total RNAs were treated with RNase-free DNase I (2 units) at 37 °C for 30 min to remove genomic DNAs, and 10 μg of RNAs was resuspended in water. Then, random primers, dNTP, 10 × RT buffer, and Multiscribe Reverse Transcriptase were added to an RNA solution and incubated at 25 °C for 10 min followed by 37 °C for 120 min to synthesize the first-strand cDNA. Real-time RT-PCR was performed using a SYBR Green Realtime PCR Master Mix (Takara Bio, Otsu, Japan) and the Applied Biosystems 7900 real-time PCR detection system (ABI, Foster City, CA). Thermal cycling conditions were as follows: first step, 94 °C for 4 min; the ensuing 40 cycles, 94 °C for 1 s, 65 °C for 15 s, and 72 °C for 30 s. The primers used here were as follows: sense, 5′-GCCATCTACAAGCAGTCACAGCAC AT-3′ and anti-sense, 5′-GGCACAAACACGCACCTCAAAGC-3′ for p53; and sense, 5′-GACTTCAACAGCGACACCCACTCC-3′ and anti-sense, 5′-AGGTCC ACCACCCTGTTGCTGTAG-3′ for GAPDH. Expression of the p53 mRNA was normalized by that of GAPDH mRNA.
Fig. 3. DAPE-induced NCI-H28 cell death is partially suppressed by knockingdown RIP1. (A) NCI-H28 cells were transfected with NC siRNA or RIP1 siRNA, and 48 h later, Western blotting was carried out. The signal intensity for RIP1 was normalized by that for β-actin. In the graphs, each column represents the mean (± SEM) normalized intensity (n = 4 independent experiments). P value, unpaired t-test. (B) Cells, transfected with NC siRNA or RIP1 siRNA, were treated with DAPE at concentrations as indicated for 24 h, followed by MTT assay. In the graph, each column represents the mean (± SEM) percentage of basal levels (MTT intensities in NC siRNA-transfected cells without DAPE treatment)(n = 4 independent experiments). P value, unpaired t-test.
1716
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 4. DAPE-induced ROS production in NCI-H28 cells. Cells, treated with DAPE (100 μM) in the presence and absence of NAC (2 mM) for periods of time as indicated, were reacted with carboxy-H2DCFDA, and the fluorescent intensity was measured. Typical fluorescent images (FIs) are shown in the left panel. DIC, differential interference contrast. Bars, 50 μm. We chose a visual field with almost equivalent cell numbers in the DIC images for analysis. In the graphs, each point represents the mean (± SEM) signal intensity in total cells (n = 4 independent experiments).
2.9. Separation into mitochondrial and cytosolic components Cells were homogenized with a sonicator in a Buffer A solution (20 mM HEPES, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, and 250 mM sucrose, pH 7.5). Lysates were centrifuged at 1000 g for 10 min, and the supernatant was further centrifuged at 10,000 g for 1 h. The pellet and the supernatant were used as mitochondrial and cytosolic components, respectively.
2.10. Western blotting Samples were loaded on 10% (v/v) sodium dodecyl sulfate (SDS)–polyacrylamide gel electrophoresis (PAGE) and transferred to polyvinylidene difluoride membrane. After blocking with TBST [20 mM Tris, 150 mM NaCl, 0.1 % (v/v) Tween-20, pH 7.5] containing 5% (w/v) of bovine serum albumin, blotting membrane was reacted with antibodies against p53 (Cell Signaling Technology, Inc., Danvers, MA, USA) or VDAC1 (Santa Cruz Biotechnology, Inc., Dallas, Texas, USA), followed by a horseradish peroxidase (HRP)conjugated anti-mouse IgG antibody. For β-actin detection, blotting membrane was reacted with an anti-β-actin antibody (Sigma, Missouri, SL, USA) followed by an HRP-conjugated antimouse IgG antibody. Immunoreactivity was detected with an ECL kit (Invitrogen) and visualized using a chemiluminescence detection system (GE Healthcare, Piscataway, NJ, USA). Protein
concentrations for each sample were determined with a BCA protein assay kit (Thermo Fisher Scientific, Rockford, IL, USA). 2.11. Monitoring of intracellular ATP levels Intracellular ATP levels were assayed using a luminescent ATP detection assay kit containing luciferase and D-luciferin (Wako Pure Chemical Industries, Osaka, Japan). ATP can be relatively quantified by detecting the fluorescence signal emitted in luciferase-mediated reaction of D-luciferin with ATP into D-oxyluciferin. Before and after treatment with DAPE, cells were incubated in the ATP detection kit for 1 min by shaking, and 10 min later under the dark conditions, the emitted light was detected with a microplate reader (ARVO X4; PerkinElmer, Inc., Waltham, MA, USA). 2.12. Statistical analysis The data presented were mean ± standard error of the mean (SEM). Statistical differences were analyzed by Dunnett's test and unpaired t-test. Values of p b 0.05 were considered statistically significant. 3. Results 3.1. DAPE induces MPM cell death Our initial attempt was to assess the effect of DAPE on cell viability for MPM cell lines. DAPE reduced cell viability for all the investigated
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
1717
Fig. 5. DAPE-induced disruption of mitochondrial membrane potentials in NCI-H28 cells. (A) Mitochondrial membrane potentials were monitored in cells treated with DAPE (100 μM) for periods of time as indicated. (B) Cells were preincubated in the presence and absence of CsA (10 μM) for 1 h, and mitochondrial membrane potentials were monitored 20 min after treatment with and without DAPE (100 μM) for 20 min. Typical red and green fluorescent images were shown in each upper panel. DIC, differential interference contrast. Bars, 50 μm. We chose a visual field with almost equivalent cell numbers in the DIC images for analysis. In the graphs, each value represents the mean (± SEM) red or green signal intensity in total cells (n = 4 independent experiments). MP, membrane potential. P values, Dunnett's test.
MPM cell lines NCI-H28, NCI-H2052, NCI-H2452, and MSTO-211H in a concentration (1–100 μM)-dependent manner, reaching nearly 0% of basal levels at 100 μM (Fig. 1A). In the flow cytometry analysis using PI and AV, PI is a marker of dead cells and AV, which detects externalized phosphatidylserine residues, is a marker of apoptotic cells [14]. In this assay, the populations of PI-positive/AV-negative, PI-negative/AV-positive, and PIpositive/AV-positive cells correspond to primary necrosis, early apoptosis, and late apoptosis/secondary necrosis, respectively [15]. DAPE significantly increased the populations of PI-positive/AV-negative and PI-positive/AV-positive cells for NCI-H28 cell line (Fig. 1B). This indicates that DAPE induces both necrosis and apoptosis of NCI-H28 cells. In the cell cycle analysis, DAPE significantly increased the proportions at the S and G2/M phase of cell cycling in parallel with decreased population at the G1 phase in NCI-H28 cells (Fig. 1C). This implies that DAPE induces cell cycle arrest at the S and G2/M phase in NCI-H28 cells. DAPE-induced reduction of NCI-H28 cell viability was partially prevented by necrostatin-1 (Nec-1), an inhibitor of RIP1 kinase to induce necroptosis (Fig. 2). This suggests that DAPE induces necroptosis. To obtain further evidence for this, RIP1 was knocked down using RIP1 siRNA (Fig. 3A). DAPE (100 μM)-induced NCI-H28 cell death was
significantly suppressed by knocking down RIP1, although cell death induced by lower concentrations (10–70 μM) of DAPE was not affected (Fig. 3B). Collectively, these results suggest that DAPE induces MPM cell death at least in part due to necroptosis. 3.2. DAPE generates ROS and damages mitochondrial membrane potentials ROS are recognized to trigger necrosis and apoptosis. DAPE increased intracellular ROS levels in a treatment time (1–20 min)-dependent manner in NCI-H28 cells, and the increase was found from at 1-min treatment, reaching the maximum at 10-min treatment (Fig. 4). DAPEinduced ROS rise was abolished by N-acetyl-L-cysteine (NAC), a ROS scavenger (Fig. 4). These results indicate that DAPE generates ROS in NCI-H28 cells. We subsequently monitored mitochondrial membrane potentials in NCI-H28 cells using DePsipherTM. DePsipherTM, a mitochondrial activity marker, has the properties of aggregating upon membrane polarization forming an orange-red fluorescent compound. If the potential is disturbed, the dye has no access to the transmembrane space and remains in or reverts to its green monomeric form. DAPE increased green fluorescent signals in a treatment time (1–20 min)-dependent manner in parallel with decreased orange-red fluorescent signals (Fig. 5A),
1718
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 6. Effects of DAPE on p53 expression and intracellular localization in NCI-H28 cells. (A) Real-time RT-PCR was carried out in cells treated with DAPE (100 μM) for periods of time as indicated. Expression of the p53 mRNA was normalized by that of the GAPDH mRNA and the value before treatment with DAPE (0 h) was regarded as 1. In the graph, each point represents the mean (± SEM) relative expression of the p53 mRNA (n = 4 independent experiments). (B) Western blotting was performed in cells treated with DAPE (100 μM) for periods of time as indicated using an anti-p53 antibody. Expression of p53 protein was normalized by that of β-actin. In the graph, each point represents the mean (± SEM) normalized expression of p53 protein (n = 4 independent experiments). (C) Cells treated with DAPE (100 μM) for periods of time as indicated were separated into the cytosolic and mitochondrial fractions, and Western blotting was carried out in each fraction using antibodies against p53 and VDAC1. In the graph, each column represents the mean (± SEM) relative mitochondrial localization of p53 (the signal intensity for p53 in the mitochondrial fraction/that in the cytosolic and mitochondrial fractions)(n = 4 independent experiments).
indicating that DAPE perturbs mitochondrial membrane potentials in NCI-H28 cells. An increase in green fluorescent signals was found from at 3-min treatment with DAPE, reaching the maximum at 10-min treatment (Fig. 5A). This suggests that ROS production precedes mitochondrial damage. DAPE-induced mitochondrial damage was prevented by CsA, an inhibitor of CypD (Fig. 5B). This suggests that DAPE may initiate CypDdependent MPT, to disrupt mitochondrial membrane potentials. 3.3. DAPE reduces intracellular ATP levels, regardless of p53-mediated mitochondrial damage Evidence has shown that in response to oxidative stress, p53 accumulates in the mitochondria and opens the MPT pore, to induce necrosis [16]. In the real-time RT-PCR analysis, DAPE did not affect expression of the p53 mRNA in NCI-H28 cells, except for a slight increase at 1-h treatment (Fig. 6A). DAPE did not also affect basal expression levels of p53 protein (Fig. 6B). To examine subcellular distribution of p53 protein,
cells were separated into the mitochondrial and cytosolic components. The immunoreactive signals for VDAC1, a mitochondrial marker, were detected in the mitochondrial components, but no signal was found in the cytosolic components (Fig. 6C). This confirms that mitochondrial and cytosolic components are successfully separated. DAPE did not increase mitochondrial localization of p53 protein as compared with that before treatment; conversely, DAPE did not significantly decrease the mitochondrial localization (Fig. 6C). This indicates that no accumulation of p53 protein in the mitochondria is induced by DAPE. This also suggests no participation of p53 in DAPE-induced MPM cell death. Intracellular ATP decrease in association with mitochondrial damage is a key factor for necrosis [5]. We finally assayed intracellular ATP concentrations in NCI-H28 cells. DAPE significantly reduced intracellular ATP concentrations as compared with those before treatment (Fig. 7). Overall, these results indicate that DAPE reduces intracellular ATP levels through a p53-independent mitochondrial pathway.
Y. Kaku et al. / Cellular Signalling 27 (2015) 1713–1719
Fig. 7. DAPE-induced intracellular ATP reduction in NCI-H28 cells. Intracellular ATP was assayed in cells treated with DAPE (100 μM) for periods of time as indicated. The fluorescence signal for ATP before treatment with DAPE (0 h) was regarded as 1. In the graph, each point represents the mean (± SEM) relative ATP level (n = 4 independent experiments).
4. Discussion In the present study, DAPE reduced cell viability in all the investigated MPM cell lines in a concentration (1–100 μM)-dependent manner. DAPE significantly increased the populations of PI-positive/AV-negative and PI-positive/AV-positive NCI-H28 cells, which correspond to primary necrosis and late apoptosis/secondary necrosis, respectively. In the cell cycle analysis, DAPE increased the proportions at the S and G2/M phase of cell cycling in parallel with decreased the population at the G1 phase in NCI-H28 cells, indicating that DAPE arrests cell cycle at the S and G2/M phase in NCI-H28 cells. We have obtained the data that DAPE does not significantly activate caspase-3, 4, 8, and 9 in NCI-H28 cells (unpublished data). Taken together, these results indicate that DAPE induces both caspase-independent apoptosis and necrosis of MPM cells. The present study focused upon the mechanism underlying nonapoptotic MPM cell death induced by DAPE. DAPE-induced reduction of NCI-H28 cell viability was partially inhibited by the RIP1 kinase inhibitor Nec-1 or knocking-down RIP1. This suggests that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells. DAPE, on the other hand, increased intracellular ROS levels in NCIH28 cells, and the effect was abolished by the ROS scavenger NAC. This accounts for DAPE-induced ROS production. DAPE also perturbed mitochondrial membrane potentials in NCI-H28 cells. DAPE-induced ROS production and mitochondrial damage were found from at 1-min and 3-min treatment, respectively. This implies that DAPE produces ROS, which triggers to disrupt mitochondrial membrane potentials. In response to death stimuli, apoptosis and necrosis occur per continuum and coexist in the same cell [5]. Production of ROS, channelmediated calcium uptake, activation of non-apoptotic proteases, and/
1719
or enzymatic destruction of cofactors required for ATP production induce both apoptosis and necrosis in dying cells. ROS damages integrity of the mitochondrial inner membrane, allowing release of apoptosisrelated factors from the mitochondria, responsible for apoptosis, or causing loss of the ability to generate ATP, responsible for necrosis [6]. In the present study, CsA, an inhibitor of the MPT component CypD, prevented DAPE-induced perturbation of mitochondrial membrane potentials. This indicates that DAPE produces ROS, to open CypDdependent MPT pore and disrupt mitochondrial membrane potentials. Expectedly, DAPE apparently decreased intracellular ATP levels in NCIH28 cells, supporting the notion for DAPE-induced necrosis in association with mitochondrial damage. Oxidative stress is shown to cause accumulation of p53 in the mitochondria, to open the MPT pore [16]. DAPE, however, affected neither basal expression levels nor mitochondrial localization of p53 protein in NCI-H28 cells. This rules out the participation of p53 in DAPEinduced mitochondrial damage and cell death. Overall, these results indicate that DAPE produces ROS, which opens CypD-dependent and p53-independnet MPT pore, to disrupt mitochondrial membrane potentials followed by intracellular ATP reduction, and then leading to necrosis of NCI-H28 cells. In conclusion, the results of the present study show that DAPE induces RIP1 kinase-mediated necroptosis of MPM cells and alternatively, necrosis by stimulating ROS production and initiating CypD-dependent MPT, to impair mitochondrial membrane potentials followed by intracellular ATP reduction. This may provide fresh insight into DAPE signaling relevant to necrosis/necroptosis. Statement of conflicts of interest The authors have no conflicts of interest. References [1] A. Degterev, Z. Huang, M. Boyce, Y. Li, P. Jagtap, N. Mizushima, G.D. Cuny, T.J. Mitchison, M.A. Moskowitz, J. Yuan, Nat. Chem. Biol. 1 (2005) 112–119. [2] D.E. Christofferson, J. Yuan, Curr. Opin. Cell Biol. 22 (2010) 263–268. [3] H. Li, H. Zhu, C.J. Xu, J. Yuan, Cell 94 (1998) 491–501. [4] M. Kvansakul, M.G. Hinds, Cell Death Dis. 4 (2013) (e909). [5] W.X. Zong, C.B. Thompson, Genes Dev. 20 (2006) 1–15. [6] E.J. Griffiths, A.P. Halestrap, Biochem. J. 307 (1995) 93–98. [7] F. Deeba, H.N. Tahseen, K.S. Sharad, N. Ahmad, S. Akhtar, M. Saleemuddin, O. Mohammad, Biochim. Biophys. Acta 1669 (2005) 170–181. [8] K. Emoto, T. Kobayashi, A. Yamaji, H. Aizawa, I. Yahara, K. Inoue, M. Umeda, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 12867–12872. [9] K. Emoto, N. Toyama-Sorimachi, H. Karasuyama, K. Inoue, M. Umeda, Exp. Cell Res. 232 (1997) 430–434. [10] Y. Ichimura, T. Kirisako, T. Takao, Y. Satomi, Y. Shimonishi, N. Ishihara, N. Mizushima, I. Tanida, E. Kominami, M. Ohsumi, T. Noda, Y. Ohsumi, Nature 408 (2000) 488–492. [11] T. Yaguchi, T. Nagata, T. Nishizaki, Behav. Brain Funct. 6 (2010) 52. [12] Y. Kaku, A. Tsuchiya, T. Kanno, T. Nakano, T. Nishizaki, Anticancer Res. 34 (2014) 1759–1764. [13] A. Tsuchiya, T. Kanno, T. Nishizaki, Cell. Physiol. Biochem. 34 (2014) 617–627. [14] D.M. Vanags, M.I. Pörn-Ares, S. Coppola, D.H. Burgess, S. Orrenius, J. Biol. Chem. 271 (1996) 31075–31085. [15] G. Pietra, R. Mortarini, G. Parmiani, A. Anichini, Cancer Res. 61 (2001) 8218–8226. [16] A.V. Vaseva, N.D. Marchenko, K. Ji, S.E. Tsirka, S. Holzmann, U.M. Moll, Cell 149 (2012) 1536–1548.
