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Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References

Extracellular ATP is an important signaling molecule mediating quite divergent specific biological effects. Even though recent studies suggest a potential role of ATP in cancer progress, its real impact in chemotherapeutic efficacy remains unclear. In the present study, we investigated the effect of ATP on the cytotoxicity of doxorubicin in various cancer cell types and found that ATP had no effect on doxorubicin cytotoxicity in colon, prostate, breast, and cervical cancers or in osteosarcoma. In contrast, ATP has divergent effects on lung cancer cells: it can protect against doxorubicin-induced cell death in non-metastatic lung cancer CL1.0 cells, but not in highly metastatic CL1.5 cells. Both apoptotic (characterized by sub-G1 peak, caspase 3 activation, poly(ADP-ribose) polymerase-1 cleavage) and necrotic (characterized by propidium iodide uptake and ROS production) features induced by doxorubicin in CL1.0 cells were reduced by ATP. In addition, ATP attenuated p53 accumulation, DNA damage (assessed by poly(ADP-ribose) formation and the comet assay) and topoisomerase II inhibition after doxorubicin treatment, and doxorubicin cytotoxicity was diminished by the p53 inhibitor pifithrin-α. Moreover, UTP, UDP, ADP, and pyrophosphate sodium pyrophosphate tetrabasic decahydrate diminished the antitumor effect of doxorubicin in CL1.0 cells, whereas purinergic P2 receptors antagonists did not abrogate the action of ATP. In summary, ATP fails to alter the antitumor efficacy of doxorubicin in most cancer cell types, except in CL1.0 cells, in which pyrophosphate mediates the cell protection afforded by ATP via attenuation of reactive oxygen species production, DNA damage, p53 accumulation, and caspase activation.

Doxorubicin (DXR) is the most prominent member of the anthracycline antibiotics and has been used to treat a wide variety of cancers for many years. The mechanisms underlying the cytotoxic effects of DXR include DNA double helix intercalation, inhibition of topoisomerase II, production of reactive oxygen species (ROS), mitochondrial dysfunction, induction of p53, and activation of caspase.[1-6] The development of resistance of tumor cells to DXR is a major problem in cancer therapy.

Adenosine 5′-triphosphate (ATP) and its breakdown product adenosine are widespread throughout body and both have been shown to regulate cell proliferation and differentiation via purinergic P2 and P1 receptors, respectively.[7] The expression of different P2 receptor subtypes and the increase in adenosine receptors in various cancer cells implicate a pathophysiological role in the regulation of cancer cell proliferation, differentiation, and apoptosis.[8, 9] Accumulating studies suggest a close connection between chronic inflammation in the tumor microenvironment and tumorigenesis.[10, 11] High concentrations of extracellular ATP released from necrotic cells during inflammation and chemotherapy are believed to be key players in the regulation of tumor biology, immune surveillance, and even chemotherapeutic efficacy.[12, 13] Accordingly, there are high concentrations of extracellular adenosine in tumor sites, and adenosine is considered to play a crucial role in immunosuppression.[14] The few current studies of extracellular ATP in the regulation of cancer cell growth and viability remain contentious. For example, ATP has been reported to enhance the cytotoxicity of etoposide in PC14 and A549 cells, as well as the antitumor effect of cisplatin in H460 lung carcinoma cells, due to increased uptake of the chemotherapeutic agents.[15, 16] In addition, ATP was demonstrated to inhibit the proliferation of LXF-289 cells,[17] but to stimulate the proliferation of A549 cells.[18]

Owing to the heterogeneity of cancer cell types and clinical developmental stages, it is worth investigating how extracellular ATP modulates the viability of cancer cells, especially in the case of chemotherapy. Because DXR is among the most widely used cancer chemotherapeutic agents, in the present study we sought to investigate not only the interaction between ATP and DXR on cytotoxicity in various cancer cell types, but also the molecular mechanisms underlying the action of ATP in regulating the death pathways induced by DXR.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References

Cells and reagents

The CL1.0, CL1.5, and PC9 cell lines were provided by Drs Pan-Chyr Yang and Zhixin Yang (National Taiwan University, Taipei, Taiwan). All other cell lines were purchased from American Type Culture Collection (Manassas, VA, USA). The CL1.0, CL1.5, MDAMB-231, MCF-7, MG63, and HeLa cells were cultured in DMEM containing 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. The PC3, HCT116, SW480, U2OS, HT-29, LNCaP, PC9, and A549 cells were cultured in RPMI 1640 medium supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. All cells were incubated at 37°C in a humidified atmosphere of 5% CO2 in air.

Doxorubicin, adenosine, ADP, ATP, UDP, UTP, propidium iodide (PI), MTT, oxidized ATP (oxATP), reactive blue-2 (RB-2), pyridoxalphosphate-6-azophenyl-1-2′,4′-disulphonic acid (PPADS), dichlorodihydrofluorecein diacetate (DCFH2-DA), and sodium pyrophosphate tetrabasic decahydrate (PPi) were purchased from Sigma Aldrich (St Louis, MO, USA). MitoSOX was purchased from Life Technologies (Carlsbad, CA, USA). Antibodies specific to p53, phosphorylated (p-) p53 (Ser15,Ser20), caspase 3, and poly(ADP-ribose) polymerase-1 (PARP-1) were purchased from Cell Signaling Technology (Beverly, MA, USA). Antibodies specific to murine double minute (MDM) 2 and MDM4 were purchased from GeneTex (Irvine, CA, USA). The antibody specific to poly(ADP-ribose) (PAR) was obtained from BD Pharmingen (San Diego, CA, USA).

Cell viability assay

After treatment with ATP (0.1–5 mM), zVAD (20 μM) and/or DXR (0.01–10 μM), cells were incubated with MTT solution (5 mg/mL in PBS) for 1 h, as described previously.[19]

Apoptosis assay

After drug treatment cells were trypsinized, washed by PBS then fixed overnight at −20°C in PBS/70% ethanol (v/v). After centrifugation (400 g, 2 min, 4°C), cells were incubated in 0.1 M phosphate-citric acid buffer (0.2 mM NaHPO4, 0.1 M citric acid [pH 7.8]) for 30 min at room temperature. Then, the cells were centrifuged and re-suspended in solution containing Triton X-100 (0.1%), RNase (100 μg/mL), and PI (80 μg/mL). The PI fluorescence was measured using flow cytometry.

Necrosis assay

The integrity of the cell membrane was assessed by determining the ability of cells to take up PI, as described previously.[19]

Detection of cytosolic and mitochondrial ROS

Cytosolic and mitochondrial ROS levels were measured by DCFH2-DA (5 μM) and MitoSOX (5 μM), respectively, as described previously.[19]

Western blot analysis and real-time PCR

Cells were lysed and immunoblotting was performed as described previously.[19] To determine p53 mRNA expression, total RNA was extracted, reverse transcribed to cDNA, and subjected to PCR fluorescence detection as described previously.[19] The human p53 forward and reverse primers used in the present study were 5′-GAT CCG TGG GCG TGA GCG-3′ and 5′-CTT CAG GTG GCT GGA GTG-3′, respectively. Results are expressed as fold induction of p53 mRNA normalized against β-actin, which was used as a loading control.

Stability of p53 protein

Cells were transfected with a p53 plasmid using Lipofectamine 2000 Reagent (Life Technologies). After 24 h, cells were treated with the agents indicated for different periods of time. Total cell lysates were prepared for western blot analysis.

Comet assay

After treatment, cells were detached, centrifuged at 400g for 2 min at 4°C, and resuspended in PBS before the Comet assay was performed according to the previously published protocol.[20]

Topoisomerase activity assay

The reaction was performed using a commercially available kit (TG1001-1; TopoGEN, Prot Orange, FL, USA). After drug treatment, a nuclear extract was obtained by manual preparation and 1 μg nuclear protein was added to the assay mixture containing 0.2 μg kDNA. After incubation for 30 min at 37°C, 4 μL stop loading dye was added to samples, followed by the addition of proteinase K (50 μg/mL) for 10–30 min at 37°C to digest the proteins in the sample. Samples were subjected to electrophoresis on a 0.8% agarose gel with 1 × Tris-acetate-EDTA (TAE) buffer. The DNA was stained with ethidium bromide then destained in distilled water before being photographed under UV light.

Statistical analysis

Data are presented as the mean ± SEM of at least three independent experiments. Analysis of variance was used to assess the statistical significance of differences, with P < 0.05 considered significant.

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References

Combined effects of ATP and DXR on various cancer cells

In the present study, we used colon cancer (SW480, HT-29, HCT116), prostate cancer (LNCaP, PC3), breast cancer (MCF-7, MDAMB-231), cervical cancer (HeLa), lung cancer (A549, PC9, CL1.5, CL1.0), and osteosarcoma (U2OS, MG63) cells to determine the combined effects of ATP (0.3 and 1 mM) and DXR (1, 3 and 10 μM) on cell viability. As shown in Figures 1 and 2(a,b,d), these cell lines exhibited different susceptibilities to DXR and ATP. Specifically, CL1.0 cells were the most susceptible to DXR-induced cytotoxicity (IC50 0.1 μM), with SW480, HT-29, HCT116, LNCaP, PC3, HeLa, MCF-7, and CL1.5 exhibiting less susceptibility (IC50 ~1–3 μM) and MDAMB-231, A549, PC9, U2OS, and MG63 being resistant (IC50 ~10 μM). Most cell types, except MCF7, A549, CL1.0, and CL1.5 cells, were sensitive to ATP (1, 5 mM)-induced cytotoxicity, albeit to different extents. In the combination study, we found that 0.3, 1 or 5 mM ATP had no effect on the concentration-dependent death response of most cell lines (except CL1.0 cells) to 0.3–10 μM DXR (Fig. 1).

image

Figure 1. Effects of ATP on doxorubicin (DXR)-induced cytotoxicity in different cancer cell lines. Cancer cells were treated with different concentrations of DXR and/or ATP, as indicated. After 24 h, cell viability as assessed using the MTT assay. Data are the mean ± SEM.

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image

Figure 2. Inhibition by ATP of doxorubicin (DXR)-induced cytotoxicity in CL1.0 cells. (a) Effects of different concentrations of DXR on the cell viability of CL1.0 and CL1.5 cells after 24 h incubation. (b,d) Cell viability of CL1.0 (b) and CL1.5 (d) cells treated with different concentrations of DXR and/or ATP for 24 h. (c) Effects of ATP (5 mM) treatment on DXR (1 μM)-induced cytotoxicity in CL1.0 cells. Pre, cells were pretreated with ATP 30 min prior to the addition of DXR; Co, cells were treated with ATP and DXR simultaneously; Post, cells were treated with ATP 3, 6 or 9 h after exposure to DXR. Cell viability was determined 24 h after the addition of DXR using the MTT assay. Data are the mean ± SEM. *P < 0.05 compared with DXR alone in the absence of ATP.

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When cells were treated with the combination of ATP with DXR, we found that 1 μM DXR-induced cell death in CL1.0 cells was inhibited by ATP (0.1–5 mM) in a concentration-dependent manner. In contrast, ATP (2 and 5 mM) had no effect on the antitumor activity of 1 μM DXR in CL1.5 cells (Fig. 2b,d). Moreover, the cytoprotective effect of ATP in CL1.0 cells was still observed when cells were treated with ATP 3 h after exposure to DXR. However, delaying ATP treatment to 6 h after exposure of CL1.0 cells to DXR resulted in a failure of ATP-induced cytoprotection (Fig. 2c). These results suggest that ATP has a cytoprotective effect in CL1.0 lung cancer cells and that this action can be achieved 3 h after DXR treatment.

Inhibition by ATP of DXR-induced cell death in CL1.0 lung cancer cells

After observing the selective cytoprotection of ATP in CL1.0 cells, we undertook experiments to elucidate the mechanisms underlying this effect. The 1 μM DXR-induced increase in the sub-G1 peak cell population was reduced following combined treatment with 5 mM ATP or 20 μM zVAD (Fig. 3a,b). Moreover, ATP inhibited DXR-induced formation of cleaved (mature) caspase 3 and the cleavage of PARP-1 (an index of caspase 3 activation; Fig. 3c). To determine whether cell necrosis also contributes to cell death in DXR-treated CL1.0 cells, we investigated cell membrane integrity and intracellular ROS levels. The results of the PI uptake assay in non-fixed cells revealed that ATP attenuates DXR-induced cell necrosis (Fig. 3d), as well as cytosolic and mitochondrial ROS production (Fig. 3e,f). In addition, 10 mM N-acetylcysteine (NAC) and zVAD had an additive effect on the protection against DXR-induced cell death (Fig. 3g). These results suggest that both caspase-dependent apoptosis and ROS-dependent necrosis are involved in DXR-induced cell death, and that attenuation of both death pathways contributes to the cytoprotective effects of ATP.

image

Figure 3. Attenuation by ATP of doxorubicin (DXR)-induced cell apoptosis, necrosis, and reactive oxygen species (ROS) production in CL1.0 cells. Cells were treated with 1 μM DXR, 5 mM ATP, 20 μM zVAD and/or 10 mM N-acetylcysteine (NAC) for 24 h (a,g) or for the times indicated (b–f). (a,b) Cells were harvested, fixed and stained with PI, and the DNA content was analyzed by flow cytometry. (b) Proportion of cells in the sub-G1 phase in the presence or absence of ATP and/or DXR after 18 or 24 h. (c) Representative immunoblotting results for caspase 3 and poly(ADP-ribose) polymerase-1 (PARP-1). (d) Cells were incubated with PI and the percentage of necrotic cells (PI positive) was determined using a FACScan. (e,f) Intracellular ROS production, as determined by dichlorodihydrofluorecein diacetate (DCFH2-DA) (e) or MitoSOX (f), in the absence or presence of ATP and/or DXR. (g) Cell viability as determined by the MTT assay. Where appropriate, data are given as the mean ± SEM. *P < 0.05 compared with DXR alone. †A significant (P < 0.05) additive effect was seen for the combination of NAC + zVAD in inhibiting DXR-induced cell death.

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Reduction by ATP of the DXR-induced p53-dependent death pathway

Because CL1.0 cells express p53,[21] we investigated whether p53 is involved in the cytoprotective action of ATP. To this end, cells were pretreated with pifithrin-α (PFTα; 3 or 10 μM), a p53 inhibitor that has been shown to inhibit DXR-induced apoptosis.[22] Alone, PFTα treatment of cells resulted in a moderate reduction in cell viability; furthermore, PFTα pretreatment partially protected CL1.0 cells against the effects of 1 μM DXR. However, after PFTα treatment, the cytoprotective effects of ATP (5 mM) were no longer evident (Fig. 4a), suggesting the involvement of p53 in ATP-induced cytoprotection.

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Figure 4. Effects of ATP on p53-dependent cell death caused by doxorubicin (DXR) in CL1.0 cells. Cells were treated with 1 μM DXR, 5 mM ATP, and different concentrations of pifithrin-α (PFTα) for 24 h (a) or for different times, as indicated (b–d). (a) Cell viability was determined by the MTT assay. (b) Representative immunoblotting results for p53 and murine double minute (MDM) 2. (c) Effects of ATP and DXR, alone and in combination, on p53 mRNA expression in CL1.0 cells (normalized against β-actin gene expression). (d) Cells were transfected with a pcDNA-p53 plasmid and treated with 10 μg/mL cycloheximide (CHX) or 5 mM ATP for 0–12 h, as indicated. After drug treatment, total cell lysates were prepared for immunoblotting of p53. β-actin was used as a loading control. Where appropriate, data are given as the mean ± SEM. *P < 0.05 compared with DXR alone. Pifithrin-α exhibited significant (P < 0.05) cytotoxicity.

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Because p53 is a protein with a short half-life that undergoes proteasomal degradation following MDM2-mediated ubiquitination, we investigated its expression following DXR and/or ATP treatment. As shown in Figure 4(b), DXR-induced upregulation of p53 expression and phosphorylation at both the Ser15 and Ser20 sites were reduced in the presence of ATP. Furthermore, following DXR treatment, there was a transient downregulation of MDM2 (a negative regulator and a gene target of p53) at 1 h, with expression gradually increasing thereafter in parallel with increases in p53 over the 9–24 h period. This biphasic effect on MDM2 expression was abrogated in the presence of ATP (Fig. 4b). In contrast, levels of MDM4, an MDM2 homolog that hetero-oligomerizes with MDM2 to stimulate its stability and E3 ubiquitin ligase activity,[23] were not changed by DXR treatment, either alone or in combination with ATP (data not shown).

Next, we examined whether ATP attenuation of p53 expression occurs at the transcriptional level. The moderate increase in p53 mRNA expression induced by DXR at 6 h was not altered by ATP (Fig. 4c). We wondered whether the protein stability of p53 in the resting state was regulated by ATP. To overcome limited p53 expression in the resting state, which hinders assessment of p53 stability, we overexpressed p53 and treated cells with 10 μg/mL cycloheximide, an inhibitor of protein synthesis. Under these conditions, without new protein synthesis, ATP decreased the half-life of p53 from 6 to 4.5 h (Fig. 4d), suggesting the involvement of p53 destabilization in the cytoprotective effect of ATP.

Effects of ATP on DXR-induced DNA damage

Because p53 induction is an event associated with DNA damage, we investigated the effect of ATP on DXR-induced DNA damage, using PARylated PARP-1 expression as an index of DNA damage. We found that DXR-induced PAR modification of PARP-1 is inhibited by ATP (Fig. 5a). The Comet assay revealed that ATP + DXR cotreatment resulted in less DNA damage than seen following treatment of cells with DXR alone (Fig. 5c).

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Figure 5. Inhibition by ATP of doxorubicin (DXR)-induced DNA damage and topoisomerase II inhibition. CL1.0 cells were treated with 1 μM DXR, 5 mM ATP, or 10 mM NAC for 1–24 h as indicated (a,b) or for 1 (c) or 3 h (d). (a,b) Representative immunoblotting results using antibodies against poly(ADP-ribose) (PAR) and p53. (c) The Comet assay was used to assess the extent of DNA damage. (d) Topoisomerase II activity in nuclear extracts of cells treated with ATP and DXR, alone or in combination. Where appropriate, data are given as the mean ± SEM. *P < 0.05 compared with DXR alone.

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To elucidate the roles of ROS in DNA damage and p53 accumulation, cells were pretreated with 10 mM NAC for 30 min. Under these conditions, NAC inhibited DXR-induced p53 expression and PARP-1-mediated automodification. Moreover, in the presence of NAC, ATP produced further decreases in the responses to DXR that were of similar extent to the effects of ATP in the absence of NAC (Fig. 5b). These results suggest that both ROS-dependent and -independent mechanisms are involved in DXR-induced activation of PAPR-1 and accumulation of p53. It is also likely that the inhibitory effects of ATP on PARP-1 activation and p53 accumulation are also mediated by both ROS-dependent and -independent mechanisms.

Because inhibition of topoisomerase II activity contributes to the DNA damage and antitumor effects of DXR, we examined the effect of ATP on topoisomerase II. An ex vivo enzymatic study revealed that the decatenated level of kDNA (an index of topoisomerase II activity) was inhibited by DXR (Fig. 5d; lane 2 vs. lane 4), whereas in the presence of ATP decatenated levels of kDNA were restored to control levels (Fig. 5d; lane 5 vs. lane 2). These results suggest recovery of DXR-inhibited topoisomerase II activity in the presence of ATP.

Pyrophosphate mediates the cytoprotective action of nucleotides in CL1.0 cells via reductions in ROS and inhibition of topoisomerases

To determine whether nucleotides and nucleoside can induce similar action as ATP, we compared the effects of ADP, UTP, UDP, and adenosine with ATP on DXR-induced cell death. At concentrations of 2 and 5 mM, UTP, UDP, and ADP did not exert marked effects on cell viability, but did protect CL1.0 cells against DXR to a similar extent as ATP (Fig. 6a,b). In contrast, 1 mM adenosine alone markedly inhibited cell viability by 75% ± 8% (n = 4; data not shown). When cells were treated with 0.1 or 0.3 mM adenosine, concentrations that had no significant effect on cell viability, DXR cytotoxicity was not affected (Fig. 6b).

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Figure 6. Sodium pyrophosphate tetrabasic decahydrate (PPi) is involved in the cytoprotection against doxorubicin (DXR)-induced cell death. (a,b,e) Cells were treated with different concentrations of drugs as indicated. After 24 h, cell viability was assessed using the MTT assay. (c,d) Cells were pretreated with reactive blue-2 (RB-2), pyridoxalphosphate-6-azophenyl-1-2′,4′-disulphonic acid (PPADS), or oxidized ATP (oxATP) for 30 min, followed by exposure to DXR (1 μM) and ATP (5 mM). After 24 h, cell viability was assessed using the MTT assay. (f) Cytosolic reactive oxygen species (ROS) production after treatment with PPi and DXR, alone or in combination, for different times as indicated. (g) Cells were treated with the drugs, as indicated, for 3 h and then topoisomerase activity was determined. Where appropriate, data are given as the mean ± SEM. *P < 0.05 compared with DXR alone.

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To determine whether purinergic receptors mediate the cytoprotective actions of nucleotides, the effects of the non-selective P2 antagonists RB-2 (100 μM) and PPADS (300 μM) on cell viability were evaluated.[7] Neither PPADS nor RB-2 were able to reverse the cytoprotective action of ATP (Fig. 6c). Recently, the P2X7 receptor has been described as a cancer cell biomarker and an activator of NLRP3 inflammasome that is capable of regulating cancer cell growth,[24-26] we determined its involvement in the cytoprotective action of ATP in the present study. To this end, we treated cells with 100 μM oxATP (a P2X7 receptor antagonist). At this concentration, oxATP had no effect on the protection afforded by ATP against DXR-induced cell death (Fig. 6d).

After observing the protective effects of nucleotides on DXR-induced cell death independent of P2 purinoceptors, we sought to determine the possible involvement of PPi in this event. As shown in Figure 6(e), PPi (1 or 3 mM) did not induce obvious cell death of CL1.0 and CL1.5 cells, with cell death observed only at concentrations of 10 mM PPi (data not shown). At 1 and 3 mM, PPi reduced DXR-induced cell death of CL1.0 cells. Similar to the actions of ATP, PPi had no effect on the DXR-induced cell death of CL1.5 cells. Moreover, the cytoprotective effect of PPi in CL1.0 cells was accompanied by reductions in DXR-induced cytosolic ROS production (Fig. 6f) and topoisomerase inhibition (Fig. 6g). These results suggest a contribution of PPi rather than P2 purinoceptors in the cell protective actions of ATP.

Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References

Doxorubicin is a highly active agent in the chemotherapeutic treatment of a number of human malignancies, but is relatively ineffective against non-small cell lung cancer.[27] Because epidemiological and experimental data suggest a close connection between inflammation and tumorigenesis, the high concentrations of nucleotides in the tumor microenvironment, particularly ATP released from necrotic cells during the processes of inflammation and chemotherapy, have gained considerable attention and are believed to regulate tumor biology, immune surveillance, and even the response to chemotherapy.[12, 13, 24] In the present study, using various cancer cell lines, we found no significant interaction between ATP and DXR in terms of cytotoxicity for most cell lines. However, the one exception was that of CL1.0 cells, a non-metastatic non-small cell lung cancer cell line, which proved to be highly susceptible to DXR and in which nucleotide treatment protected against the effects of DXR. Consistent with the type of malignancy, the differential expression of several genes and proteins associated with invasiveness and metastasis has been demonstrated between CL1.0 and CL1.5 cells.[28-30] Currently, we cannot provide further evidence to explain this cancer cell-type specificity, but we did elucidate the mechanisms underlying the cytoprotective effect of ATP in DXR-treated CL1.0 cells, which highlights the role of ATP in chemotherapeutic effectiveness.

Previous studies have demonstrated that caspase activation and ROS production are involved in DXR-induced apoptosis and necrosis, respectively.[3, 31] The results of the present study regarding PI uptake, PARP-1 and caspase 3 cleavage, and ROS production indicate that there are various mechanisms underlying DXR-induced cytotoxicity in CL1.0 cells. Furthermore, the additive cytoprotective effects of NAC and zVAD provide further evidence for DXR-induced cell death being due to multiple mechanisms. The findings of the present study suggest that inhibition of ROS-mediated necrosis and caspase-dependent apoptosis contribute to ATP-induced cell protection. Previous studies indicate that p53 accumulation and activation participate in DXR-induced apoptosis.[4, 6] In the present study, inhibition of DXR cytotoxicity by pifithrin-α, a p53 inhibitor, suggests that p53 activation is involved in the cell death of CL1.0 cells. We also found that ATP inhibits DXR-induced p53 protein accumulation and that this effect is not related to gene transcription, but may result from inhibition of DNA damage, an initiator of p53 activation, and p53 protein stability.

Our data further demonstrate the involvement of ROS-dependent and -independent mechanisms in DXR-induced p53 expression and ATP inhibition of this effect. Both DNA damage and oxidative stress have been shown to mediate DXR-induced activation of the ataxia telangiectasia mutanted (ATM)–p53 apoptosis pathway by causing MDM2 degradation and disruption of the interaction between p53 and MDM2.[32] Because MDM2 is also a target gene of p53, the activity of MDM2 can be restored after p53 stabilization and activation.[33] In the present study, we found early downregulation and late recovery of MDM2 levels after DXR treatment of CL1.0 cells. Furthermore, the observation that ATP prevents MDM2 recovery following DXR treatment suggests an inhibitory action of ATP on p53. An early result of DXR treatment is DNA damage resulting from inhibition of topoisomerase II activity and the induction of oxidative stress.[34] On the basis of results of experiments evaluating PAR formation, as well as the Comet assay, we conclude that ATP is able to reduce the extent of DNA damage. On the basis of our findings, we suggest that the reductions in DNA damage and p53-mediated death signaling are due to ATP inhibition of ROS production and recovery of topoisomerase II activity.

Extracellular ATP may regulate cellular functions through its PPi chain.[35] In the present study, we ruled out an action of ATP via P2 purinoceptors, but found that PPi exerts similar effects as ATP in CL1.0 cells, including cell protection, reductions in ROS production, and attenuation of DXR-induced topoisomerase inhibition. Moreover, consistent with the results obtained for ATP, PPi had no effect affect DXR-induced cytotoxicity in other cancer cell lines (data not shown). Thus, we suggest the cytoprotective action of ATP is not mediated via purinoceptors, but is due to pyrophosphate actions.

In conclusion, ATP and other nucleotides can protect CL1.0 lung cancer cells against DXR-induced cell death via the pyrophosphate moiety. Recovery of topoisomerase II activity, attenuation of ROS production, DNA damage, and p53 accumulation, and activation of caspase may all contribute to the cytoprotective effect of ATP. The findings of the present study highlight cell type-specific actions of ATP in regulating the chemotherapeutic efficacy of DXR.

Acknowledgments

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References

The authors are grateful to Drs P. C. Yang and Z. Yang (National Taiwan University, Taipei, Taiwan) for providing the cell lines. This work was supported by the cooperative research program of National Taiwan University, College of Medicine (NTUCM) and China Medical University, College of Medicine (CMUCM) (grants 99F008-301 and 100F008-402).

References

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgments
  7. Disclosure Statement
  8. References