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Evaluation of the molecular mechanisms of a palladium(II) saccharinate complex with terpyridine as an anticancer agent

Kacar, Omera; Adiguzel, Zelala; Yilmaz, Veysel T.b; Cetin, Yuksela; Cevatemre, Busec; Arda, Nazlie; Baykal, Ahmet T.a; Ulukaya, Engind; Acilan, Ceydaa

doi: 10.1097/CAD.0b013e328364c6ad
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Metal-based compounds represent promising anticancer therapeutic agents. In this study, the mechanism of action of a novel metal-based drug, a palladium(II) (Pd) complex ([PdCl(terpy)](sac)·2H2O, terpy=2,2′:6′,2′′-terpyridine and sac=saccharinate), was elucidated. The tested compound induced cytotoxicity in nine different human cancer cell lines that originated from various organs, suggesting a broad spectrum of activity. The IC50 values were significantly higher for noncancerous cells when compared with cancer cells. We found that cells treated with the Pd(II) complex exhibited increased caspase 3/7 activities and condensed/fragmented nuclei, as demonstrated by nuclear staining and DNA laddering. Morphological features, such as cellular shrinkage and blebbing, were also observed, indicating that apoptosis was the primary mechanism of cell death. Pd(II) treatment induced DNA double-stranded breaks both in vitro and in vivo, potentially accounting for the source of stress in these cells. Although caspase 3/7 activities were elevated after Pd(II) treatment, silencing or using inhibitors of caspase 3 did not block apoptosis. Other molecules that could potentially play a role in Pd(II)-induced apoptosis, such as p53 and Bax, were also tested using silencing technology. However, none of these proteins were essential for cell death, indicating either that these molecules do not participate in Pd(II)-induced apoptosis or that other pathways were activated in their absence. Hence, this new molecule might represent a promising anticancer agent that exhibits cytotoxicity in p53-mutant, Bax-mutant, and/or caspase 3-mutant cancer cells.

Supplemental Digital Content is available in the text.

aTUBITAK, Marmara Research Center, Genetic Engineering and Biotechnology Institute, Kocaeli

Departments of bChemistry

cBiology, Faculty of Arts and Sciences

dDepartment of Medical Biochemistry, Medical School, Uludag University, Bursa

eDepartment of Molecular Biology and Genetics, Faculty of Arts and Sciences, Istanbul University, Istanbul, Turkey

Supplemental digital content is available for this article. Direct URL citations appear in the printed text and are provided in the HTML and PDF versions of this article on the journal's website (http://www.anti-cancerdrugs.com).

Correspondence to Ceyda Acilan, PhD, TUBITAK, Marmara Research Center, Genetic Engineering and Biotechnology Institute, 41470 Gebze Kocaeli, Turkey Tel: +90 262 677 3354; fax: +90 262 641 2309; e-mail: ceyda.acilan@tubitak.gov.tr

Received February 11, 2013

Accepted July 3, 2013

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Introduction

Among chemotherapeutic drugs, cisplatin is widely recognized for its application in the treatment of a spectrum of cancers, including breast, prostate, small-cell lung, ovarian, and germ cell cancers and neuroblastoma, sarcoma, and lymphoma 1,2. Despite its broad efficacy, cisplatin has major limitations, including toxicity, drug resistance, and poor oral bioavailability 3. These drawbacks have triggered the development of alternative platinum (Pt)(II)-based compounds 4. However, among the thousands of molecules that have been tested, only two (carboplatin and oxaliplatin) have garnered FDA approval 5. Therefore, other metal-based drugs, including palladium(II) (Pd) complexes, have attracted interest as potential anticancer agents 6.

Pd(II) and Pt(II) complexes are generally isostructural and exhibit similar coordination chemistry and geometry 7. In chemistry studies, Pd(II) systems have long been used as a model for studying the interactions of Pt(II) with DNA and for modeling its binding properties, because Pd(II) compounds reach equilibrium more quickly than Pt(II) 8. The labile property of Pd(II) complexes has been proposed to be useful for the treatment of cancer types such as those within the gastrointestinal region, wherein cisplatin is therapeutically ineffective 7. More stable Pd(II) complexes have also been synthesized by the addition of less labile groups that produce kinetically inert molecules. Indeed, several studies have reported the antiproliferative and antitumor effects of such Pd(II) complexes 6,9–13. Both the growth inhibitory and lipophilic activities of Pd(II) complexes have been reported to be comparable with, although not more than, those of cisplatin 14,15. The cytotoxicity of Pd(II) complexes has been reported in various cancer cell types, even at low concentrations 10–13,16. Further, Pd compounds appear to be more soluble compared with Pt and thus present as more favorable anticancer drugs 14. Another important feature of Pd complexes is that some derivatives exhibit reduced nephrotoxicity when compared with cisplatin 17. Therefore, Pd(II) complexes represent promising antitumor agents in terms of their chemical and physical properties.

Recently, our group reported significant cytostatic and cytotoxic in-vitro properties of the Pd(II) complex in non-small-lung cancer cells 12. As the modes and extents of cell death can vary between different cell types, in this study, we screened the toxicity of this complex in nine different cancer cell lines. To further elucidate the mechanisms of toxicity, we selected two different cancer lines, MDA-MB-435 and HeLa cells, and investigated the means of cell death using morphological and biochemical methods. Our data indicated that cell death predominantly resulted from apoptosis. To understand the underlying molecular pathways, we assessed whether molecules such as p53, Bax, or caspase 3 were essential for the apoptotic effects of the Pd(II) complex using small interfering RNA (siRNA)-mediated silencing of these candidate genes. We determined that either they are not essential or that other cell death pathways are activated in their absence. Finally, the Pd(II) complex appeared to induce its apoptotic effects by introduction of DNA double-stranded breaks (DSBs), as suggested by our studies.

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Materials and methods

Cell culture

All cells used in this study were maintained in Dulbecco’s modified Eagle medium-F12 (DMEM/F12, #D0547; Sigma-Aldrich, St Louis, Missouri, USA) supplemented with 5% FBS (#S0415; Biochrom, Berlin, Germany) and penicillin (100 U/ml)–streptomycin (100 μg/ml) (#A2212; Biochrom). The cells were cultured at 37°C in 5% CO2. The cells were passaged using trypsin (0.05%, L2143; Biochrom) upon reaching 90–95% confluence.

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Cell viability

The Pd(II) compound was dissolved in DMEM/F12 without serum (stock solution: 500 μmol/l, molecular weight: 593.32 g/mol). Serial dilutions were freshly prepared in cell culture medium on the day of the experiment and 7.5×103 cells/well were seeded onto 96-well plates. The day after seeding, the cells were treated with concentrations of the Pd(II) compound ranging from 0.8 to 50 μmol/l for 24, 48, or 72 h. The medium was then aspirated and WST-1 reagent (Roche Diagnostics, Mannheim, Germany) was added (1 : 10 dilution in DMEM/F12, 5% FBS). Cell viability was measured at 450 nm with a reference wavelength of 650 nm at 2 h after the addition of WST-1. The absorbance from untreated cells was regarded as 100% viability, and cell viability was measured as a percentage of the untreated samples. Each experiment was performed in at least three biological replicates.

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Nuclear condensation/fragmentation analysis

For nuclear condensation and fragmentation analyses, 4×104 cells were seeded onto 12-mm coverslips, treated with the Pd(II) complex at the indicated doses, fixed after 24, 48, or 72 h in −20°C methanol (10 min), and stained with 5–10 μl of Hoechst 33342 dye (100 μg/ml). Images were acquired using a ×20 objective and an A4 filter on a Leica DMI 6000 fluorescence microscope (Leica, Wetzlar, Germany).

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Live cell imaging

The protocol described by Acilan et al.18 was used with minor modifications. Briefly, immediately after addition of the Pd(II) complex, images were acquired at 5-min intervals for 72 h unless stated otherwise. Short movies were made at 25 images/s.

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DNA fragmentation assay

A modified protocol described by Zhu and Wang 19 was used for the DNA fragmentation assay. Briefly, 2.5×105 cells were seeded onto 25 cm2 flasks and treated with the Pd(II) compound for 72 h. Both floating and attached cells were collected and combined. The cells were washed with PBS and centrifuged [1000g, 5 min, room temperature (RT)]. The pellet was dissolved in 120 μl of lysis buffer [10 mmol/l Tris (pH 7.4), 100 nmol/l NaCl, 25 mmol/l EDTA, 1% N-lauryl sarcosine, and proteinase K (final concentration: 0.35 μg/μl)] by gentle vortexing and was incubated at 45°C for 2 h. The lysates were further incubated for 1 h at RT after addition of 2 μl of RNAse A (10 μg/ml). The samples were resolved on 2% agarose gels (stained with SYBR Green, 60 V, 4 h) and analyzed using a UVITEC imaging system (UVItec Ltd, Cambridge, UK). The assay was performed in triplicate with similar results for each replicate.

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Caspase 3/7 activity detection

The ApoTox-Glo Triplex Assay kit from Promega Corporation (#G6320; Madison, Wisconsin, USA) was used to measure caspase 3/7 activities. Briefly, 1×104 cells were seeded onto 96-well plates. The next day, the test compound and vehicle controls were added to appropriate wells to achieve a final volume of 100 µl and incubated for 24 h under standard culture conditions. One hundred microliters of caspase-Glo 3/7 reagent was added to each well. After a 30-min incubation at RT, the resulting chemiluminescence was measured using a DTX 880 Multimode Detector (Beckman Coulter, Krefeld, Germany). Ac-DEVD-CHO (A0835, N-acetyl-Asp-Glu-Val-Asp-al; Sigma-Aldrich) was used at a final concentration of 6.5 μmol/l to inhibit caspase activity (24 h incubation). To determine the number of cells, a replica plate was prepared and cell viability was measured using the WST-1 reagent as described above. The graphs indicate caspase 3/7 activity (the ApoTox-Glo Triplex Assay kit from Promega Corporation) after normalization to the number of cells in the wells. Cell number was extrapolated from cell viability graphs, where untreated controls were evaluated as 100% viable.

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γH2AX staining

For γH2AX staining, 4×104 cells were seeded onto 12 mm round coverslips and were treated with 50 μmol/l Pd(II) complex on the day after seeding. At 2 h after the treatment, cells were fixed in 4% paraformaldehyde/PBS for 10 min, washed with PBS, permeabilized in 0.5% (v/v) Triton-X/PBS for 10 min, and rewashed with PBS. The coverslips were blocked using 1% (w/v) BSA (#0332; Amresco, Solon, Ohio, USA) in PBS for 1 h. The cells were stained with γH2AX antibodies (1 : 1000 diluted in 5% BSA/PBS, #ab2893; Abcam, Cambridge, Massachusetts, USA) at +4°C overnight, followed by staining with FITC-conjugated secondary antibodies (#ab97068, Abcam; 1 : 500 diluted in 5% BSA/PBS) for 1 h at RT. The nuclei were counterstained with 5 μg/ml DAPI in 50% glycerol/PBS. The slides were either visualized immediately or kept in the dark at −20°C until subsequent examination using a fluorescence microscope (Leica DMI 6000).

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In-vitro analysis of Pd(II)/DNA interactions

Reactions were performed in a final volume of 20 μl ddH2O. The indicated concentrations of the Pd(II) complex were serially diluted in ddH2O. Plasmid DNA (250 ng) (8454 pCMV-VSV-G, plasmid 8454, 6363 bp; Addgene, Cambridge, Massachusetts, USA) and DNAse inactivation reagent (2 μl) (#3174G; Ambion, Grand Island, New York, USA) were added to each tube. The mixtures were vortexed and then centrifuged at 11 000g for 1.5 min at RT. The supernatants were transferred to a new tube. Sodium azide (0.05 mol/l, #S-2002; Sigma-Aldrich) was added to tubes as indicated in the figure. The tubes were incubated at RT for 20 h. DNA was resolved on 1% SYBR Green-stained agarose gels.

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Transfection

Commercial siRNAs were used to silence Bax (#L-003308-01; Dharmacon, La Fayette, Colorado, USA), TP53 (#L-003329-00; Dharmacon), and caspase 3 (#SI02654603; Qiagen, Valencia, California, USA). Nontargeting siRNA (#D001810-10-05; Dharmacon) was used as a control. The pCasper3-GR vector was purchased from Evrogen (Cat. #FP971). Transfections were performed in 96-well and six-well plates in parallel to determine cell viability after Pd(II) treatment and the efficiency of silencing, respectively. Cells were seeded at 50–60% confluence and at 6 h after seeding, when the cells were newly attached. One microgram of siRNA or plasmid DNA was used to transfect cells in six-well plates using Lipofectamine 2000 (Life Technologies, California, USA) with a 1 : 3 (w : v) ratio following the manufacturer’s instructions (final transfection volume: 1 ml). For the 96-well plates, 0.05 μg of siRNA was used to transfect the cells (final transfection volume: 50 μl). The cells in six-well plates were harvested for RT-PCR analysis at 24 and 48 h after siRNA transfection to confirm the knockdown of these genes. The time points corresponded to the beginning and end of the cell viability experiments that were performed in 96-well plates.

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RT-PCR

Total RNA was isolated using the NucleoSpin RNAII kit (Macherey-Nagel, Düren, Germany) following the manufacturer’s protocols. An additional DNAse treatment was carried out using RNase-free DNase (Promega). The iScript cDNA synthesis kit (Bio-Rad Laboratories, Berkeley, California, USA) was used to reverse transcribe 500 ng of total RNA according to the manufacturer’s protocols (20 μl/reaction). Real-time PCR was performed using the SYBR Green Supermix (Bio-Rad Laboratories) on a Bio-Rad iQ5 PCR Thermal Cycler. The samples were normalized to glyceraldehyde 3-phosphate dehydrogenase expression levels. The list of primers is presented in Table 122,23. Each sample was tested in duplicate. To confirm the amplification specificity, the PCR products were subjected to a melting curve analysis and subsequent agarose gel electrophoresis.

Table 1

Table 1

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Statistical analyses

The SPSS statistical package (17.0 Version; SPSS Inc., Chicago, Illinois, USA) was used for data analysis. The normality of the variables [inhibitory concentration 50 (IC50) values and ‘area under curve’ (AUC) calculations] was analyzed using the Kolmogorov–Smirnov test and was found to exhibit a non-normal distribution (P>0.05). Therefore, we analyzed our data using nonparametric tests. The differences between the control and patient or treatment groups were assessed using the Mann–Whitney U-test. The IC50 values were extrapolated from the cell viability curves based on a regression line in which cell death was within the linear range. The AUC was determined as the mathematical calculation of the area under the cell viability curves. Values with a confidence interval of 95% and a P value less than 0.05 were considered to be statistically significant.

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Results

The molecular structure of the Pd(II) complex used in this study is shown in Fig. 1. The compound was previously synthesized by our group, and the synthesis, characterization, and radiographic structure of this complex have been reported previously elsewhere 20. The antiproliferative and cytotoxic effects of this compound were investigated and compared between various cancer cell lines (MDA-MB-231, MCF-7, PC-3, MDA-MB-435, HeLa, SH-SY5Y, A172, U-87, and C6) and noncancer control cell lines (primary human aortic smooth muscle cells: HASMC-1 and 2; Chinese hamster ovary cells: CHO-K1). All cells were exposed to the Pd(II) complex at concentrations ranging from 0.8 to 50 μmol/l for 24, 48, and 72 h, and cell viability was assessed using a WST-1 assay (Fig. 2). The Pd(II) complex induced cell death in both a dose-dependent and time-dependent manner, and cytotoxicity was observed as early as 24 h, although the lethal dose (IC50) was different for each cell line. The toxicity was most prominent at 72 h, even at the lowest concentrations tested. Hence, the new Pd(II) complex appears to be ubiquitously cytotoxic in different cancer types, despite variable efficacy. The IC50 values of the Pd(II) compound on the tested cell lines at 24, 48, and 72 h are shown in Table 2. The IC50 values of the Pd(II) complex in cancer cells at 72 h of drug exposure were found to be within the range of 2.5–21.7 μmol/l, with an average of IC50 of 7.3 μmol/l. In contrast, the IC50 for normal cells was between 12 and 23 μmol/l, with an average IC50 of 19.6 μmol/l. Thus, it appears that the effects of this complex are more potent in cancer cell lines than in the noncancer control cell lines, at least for the cell types assessed in this study (P=0.021, Mann–Whitney U-test).

Fig. 1

Fig. 1

Fig. 2

Fig. 2

Table 2

Table 2

To determine the mode of cell death, two cell lines, HeLa and MDA-MB-435, were selected for further analysis. The cells were exposed to different concentrations of the Pd(II) complex and fixed at 24, 48, and 72 h for visualization of their nuclear morphologies (Fig. 3; Supplementary Fig. 1, Supplemental digital content 1, http://links.lww.com/ACD/A35). As expected, the untreated cells appeared healthy with normal nuclei and mitotic figures (Fig. 3, 0 μmol/l; Supplementary Fig. 1, Supplemental digital content 1, http://links.lww.com/ACD/A35). In contrast, after Pd(II) treatment, both cell lines exhibited nuclear condensation and fragmentation as early as 12.5 μmol/l treatment for 24 h. There was also an increase in cell number per field at 72 h in the untreated samples, which reduced significantly as cells detached and started floating upon exposure to the complex.

Fig. 3

Fig. 3

To further evaluate when and how this compound affects the morphology of cells, both HeLa and MDA-MB-435 cell lines were treated with the drug (at a concentration of 25 μmol/l) and imaged every 5 min for 72 h, whereas the untreated cells remained attached and divided during the time course (Fig. 4a; Supplementary Fig. 2a, Supplemental digital content 1, http://links.lww.com/ACD/A35; Supplemental Movie 1a, Supplemental digital content 2, http://links.lww.com/ACD/A36; and Supplemental Movie 2a, Supplemental digital content 3, http://links.lww.com/ACD/A37). Both cell lines exhibited typical apoptotic characteristics such as shrinkage, blebbing, and the formation of spikes (Fig. 4b; Supplementary Fig. 2b, Supplemental digital content 1, http://links.lww.com/ACD/A35; Supplementary Movie 1b, Supplemental digital content 4, http://links.lww.com/ACD/A38; and Supplementary Movie 2b, Supplemental digital content 5, http://links.lww.com/ACD/A39). Apoptosis started between 4 and 6 h and occurred at various time points, unlike necrosis, in which most cells simultaneously die at once. Some cells formed membrane blisters upon prolonged exposure to the agent. Thus, the morphological assessment of cells treated with the Pd(II) complex strongly implicated apoptosis as the form of cell death.

Fig. 4

Fig. 4

In addition to the morphological analyses, we evaluated the activities of caspase 3/7 using microscopic and biochemical methods. Initially, cells were transfected with a plasmid that encodes for a GFP tag containing the caspase 3/7 cleavage sequence (DEVD, pCasper3-GR), which fluoresces green only after being cleaved. After transfection, the cells were treated with 25 μmol/l of the Pd(II) complex and were imaged every 5 min for 48 h. Whereas there was no green fluorescence at 0 h, after the addition of the Pd(II) complex, accumulation of green signal started at 4 h, consistent with the time at which the cells started shrinking and blebbing (Fig. 5a; Supplementary Movie 3, Supplemental digital content 6, http://links.lww.com/ACD/A40). The fluorescence started fading by 28 h, indicating a peak time for caspase activity between 4–28 h. This experiment most likely underestimates the number of cells undergoing caspase 3/7-dependent apoptosis because not all cells in the field were transfected with the plasmid. To quantitatively measure caspase 3/7 activity, cell lysates were prepared at 24 h Pd(II) exposure, a time point at which caspase 3/7 was active (Fig. 5a), and significant cell death occurred at higher doses (Fig. 2). Indeed, there was an increase in caspase 3/7 activities in both cell lines, although the precise fold-change varied between the two cell types (Fig. 5b, black columns). The activity could also be prevented by the addition of caspase 3/7 inhibitors to the reaction mixture, confirming the specificity of the assay (Fig. 5b, gray columns).

Fig. 5

Fig. 5

Finally, DNA fragmentation was visualized by agarose gel electrophoresis (Fig. 6). Interestingly, whereas the HeLa cells consistently exhibited DNA laddering, the MDA-MB-435 cells did not. Because all of the other data implicated the apoptotic pathway as the form of death in MDA-MB-435 cells, we concluded that these cells undergo apoptosis without exhibiting DNA laddering.

Fig. 6

Fig. 6

Cisplatin induces toxicity predominantly by DNA cross-linking, which triggers a cascade of events that lead to apoptosis 21. To understand whether the new Pd(II) complex also acts through DNA damage, we performed an in-vitro assay in which plasmid DNA was incubated with the Pd(II) complex. The ability of the agent to interact with DNA was tested by agarose gel electrophoresis to detect a possible change in the migration pattern of plasmid DNA. Interestingly, we observed smearing of DNA at 50–100 μmol/l of Pd(II) treatment and loss of open, closed, and supercoiled forms (Fig. 7a; Supplementary Fig. 3, Supplemental digital content 1, http://links.lww.com/ACD/A35), indicating that the Pd(II) complex might induce DNA DSBs. Further, the addition of sodium azide to the reaction mixture prevented DNA smearing, suggesting that the induction of DSBs by the Pd(II) complex involved hydroxyl radical formation. To investigate whether this was the case in vivo, cells were stained for γH2AX, which is a phosphorylated histone H2A variant that is present at the sites of DSBs and at DNA damage foci and is used as a marker for DSBs 24. Indeed, cells exhibited foci formation after exposure to Pd (Fig. 7b), suggesting that the mechanism of action for the toxicity of this new Pd(II) complex is through DSB induction.

Fig. 7

Fig. 7

Because the agent appeared to induce DSB, the involvement of candidate molecules such as caspase 3, p53, and Bax, which are known to play a role in response to DNA damage-induced apoptosis 25, was evaluated by siRNA-mediated silencing in both cell lines (Supplementary Fig. 4, Supplemental digital content 1, http://links.lww.com/ACD/A35, silencing efficiency: 70–90% knockdown as determined by RT-PCR analysis). Although all the genes were efficiently silenced, there was no significant difference in cell viability between the silenced cells and the control cells that were transfected with nontargeting siRNA or were mock transfected (analysis based on AUC, Mann–Whitney U-test, P>0.1 for all genes, Fig. 8a). Using inhibitors against caspase 3/7 (Ac-DEVD-CHO), similar results were obtained. Hence, we silenced both p53 and Bax genes, as well as simultaneously inhibited caspase 3/7 activities, to evaluate whether there is a combinatorial effect. However, there was no significant change in cell viability when compared with the cells transfected with either nontargeting siRNA or vehicle control. To assess whether the cause of cell death was still apoptosis, HeLa cells were analyzed for Pd(II)-induced DNA fragmentation after silencing (Fig. 8b). Our data indicated that DNA laddering still occurred in the absence of caspase 3, Bax, or p53, indicating that the cells died by apoptosis.

Fig. 8

Fig. 8

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Discussion

To overcome cancer, metal-based agents have emerged as promising anticancer drugs. Pd(II) complexes have gained significant interest, particularly after the success of cisplatin, which is a Pt-based drug that is widely used in clinical practice. In this study, we further investigated a newly synthesized Pd(II) complex that was previously found to be cytotoxic in lung cancers 11. Here, we report our findings on the mechanism of action of this novel agent using in-vitro models.

In this study, we expanded the previous cytotoxicity analyses to breast, prostate, cervix, neuroblastoma, and glioma cells and observed that our compound was toxic to all cancer types, indicating a broad efficacy of the agent regardless of the cell type. Pd(II) elicited increasing cytotoxicity on the tested cell lines that was proportionate to increasing concentrations and durations of exposure. For some cell lines, there was a critical concentration after which a sharp decrease in viability was detected. Hence, it appears that the toxic effect of the Pd(II) complex induces an initial damage that cells attempt to resist or repair, subsequently dying when the damage exceeds the cellular repair capacity. In support of this hypothesis, at low doses, there was increased proliferation in some cells, which was followed by cell death at increased doses (Fig. 2).

Damage-induced proliferation has previously been detected in response to several other agents, including hydrogen peroxide, which stimulates cell growth at low levels and causes death at higher doses 26. DNA DSBs are known to induce proliferation to facilitate homologous recombination repair 27. Thus, our result that Pd(II) induces DSBs (Fig. 7) is consistent with the finding of break-induced replication at low doses and lethality at nonreparable doses.

The primary biological target of cisplatin appears to be DNA. Cisplatin induces DNA adducts predominantly by forming intrastrand cross-links and, to a reduced extent, interstrand cross-links 28. Cisplatin also induces DSBs, although the formation of cisplatin-induced DSBs is thought to occur during DNA replication 29. A detailed analysis of the interactions between the Pd(II) complex and fish sperm DNA has indicated that this complex tightly binds to DNA by both intercalation and covalent binding 30. Our analysis supports this finding, as we observed a slight shift in the migration pattern of supercoiled DNA when incubated with the complex (Fig. 7a; Supplementary Fig. 3, Supplemental digital content 1, http://links.lww.com/ACD/A35). After a specific titration of Pd(II) complex/DNA, a significant degradation of plasmid DNA was observed, suggesting that this complex could also induce DSBs. The addition of sodium azide to the reaction mixture reverted the degradation of the DNA, implying that the Pd(II) complex induced DSBs by hydroxyl radical formation. We also confirmed the formation of DSBs in vivo by immunostaining for γH2AX, a well-established marker for DSBs 24. Thus, the toxicity of the Pd(II) complex involves DNA damage.

Several approaches were undertaken to elucidate the form of cell death induced by the Pd(II) complex. Our data indicated that cells predominantly died by apoptosis, as suggested by our morphological and molecular analyses. Both MDA-MB-435 and HeLa cells exhibited shrinkage and blebbing, which is a distinct marker for apoptosis. The DNA staining of these cells revealed nuclear condensation and fragmentation in both cell types. However, no DNA laddering was observed in the MDA-MB-435 cells. This was not unexpected, as other studies have reported that different cells can exhibit different apoptotic properties 31. One of the explanations includes the cleavage of DNA to 50 kB, which was not detectable by the assay used in this study. The enzyme responsible for DNA fragmentation, caspase-activated DNase (CAD), is normally inhibited by the inhibitor of caspase-activated DNase (ICAD) 32. During apoptosis, ICAD is cleaved by caspase 3, releasing active CAD. As MDA-MB-435 cells exhibited enhanced caspase 3/7 activity (Fig. 5b), an alternative possible explanation for the lack of DNA laddering might be the lack of ICAD or the expression of a caspase-resistant mutant ICAD.

Because the Pd(II) complex appeared to damage DNA, we hypothesized that Pd-induced apoptosis would involve apoptotic molecules such as p53, Bax, and caspase 3 31. However, the knockdown of any of these proteins did not rescue the cell death phenotype, indicating that these molecules were not essential for Pd(II)-induced apoptosis. As most cancer cells harbor mutations in p53, caspase 3, or Bax, this result suggests that the Pd(II) complex will still be effective in inducing apoptosis in mutant cancer cells, increasing its potential value as an anticancer agent. Interestingly, although caspase activity was clearly elevated in response to Pd(II) treatment in both cell lines, neither its knockdown nor its inhibition prevented cell death. It is known that in biological systems, when a gene is silenced or lost, the expression of another gene might increase and compensate for the loss of activity. For example, in Chinese hamster ovary cells, the silencing of caspase 3 alone does not elicit effects on sodium butyrate-induced cell death because caspase 7 expression increases, compensating for its apoptotic effects. Thus, apoptosis can only be effectively abrogated when both genes are knocked down 33. The simultaneous knockdown of p53 and Bax together, in addition to the inhibition of caspases 3/7 using Ac-DEVD-CHO inhibitors, did not rescue cell death in response to Pd(II) treatment. Thus, other molecules might be activated during Pd(II)-induced apoptosis, and the silencing of two or more genes might be required to prevent cell death.

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Conclusion

The new Pd(II) complex appears to represent a new potent anticancer agent that acts through DNA damage and can induce apoptosis independent of p53, Bax, or caspase 3, suggesting that this agent is toxic even to cancer cells that harbor mutations in these genes. Indeed, further studies in animal models are required for proof-of-concept before undertaking clinical studies.

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Acknowledgements

This work was supported by the internal TUBITAK funding.

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Conflicts of interest

There are no conflicts of interest.

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Keywords:

apoptosis; cancer; cytotoxicity; metal-based anticancer agents; palladium(II) complex

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