Dinuclear Organoruthenium Complexes Exhibiting Antiproliferative

Dec 11, 2018 - complexes with DNA, agarose gel electrophoresis was applied to. Figure 5. ... energy gaps of these complexes vary as1 (1.91 eV) >3(1.79...
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Dinuclear Organoruthenium Complexes Exhibiting Antiproliferative Activity through DNA Damage and a Reactive-Oxygen-SpeciesMediated Endoplasmic Reticulum Stress Pathway Jian Zhao,†,‡ Shuang Li,† Xinyi Wang,† Gang Xu,†,‡ and Shaohua Gou*,†,‡ †

Pharmaceutical Research Center and School of Chemistry and Chemical Engineering and ‡Jiangsu Province Hi-Tech Key Laboratory for Biomedical Research, Southeast University, Nanjing 211189, China

Inorg. Chem. Downloaded from pubs.acs.org by EASTERN KENTUCKY UNIV on 01/24/19. For personal use only.

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ABSTRACT: Subtle ligand modifications on ruthenium arene complexes can lead to different mechanisms of action and result in significant changes in the anticancer efficacy. Herein, four novel dinuclear ruthenium(II) arene complexes were designed and prepared. In vitro tests indicated that complexes 1−3 displayed moderate antiproliferative activity against the tested cancer cells, while the cytotoxicity of complex 4 is superior or comparable to that of cisplatin. Further studies indicated that complexes 1−4 induce cell death through DNA interaction and a reactiveoxygen-species-mediated endoplasmic reticulum (ER) stress pathway, which is the first example of an organometallic ruthenium(II) arene complex to induce ER stress as well as DNA interaction. This kind of dinuclear ruthenium(II) arene complex has unique biological characteristics and is a promising model for new anticancer drug development.

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INTRODUCTION Platinum-based anticancer drugs, including cisplatin, carboplatin, and oxaliplatin, have achieved tremendous successes in cancer therapy, which are still used in more than 50% of all chemotherapeutic regimens.1−3 However, the severe side effects and inevitable drug resistance have limited their clinical application. Ruthenium anticancer agents with relatively lower toxicity are regarded as promising anticancer drug candidates, which may have the potential to overcome the limitations of the platinum drugs.4−12 So far, two ruthenium(III) complexes (NAMI-A, KP1019, and its sodium salt KP1339) have entered clinical trials (Figure 1).13−16 In recent years, ruthenium(II) arene complexes have attracted much attention because several compounds are close to clinical trials, such as RM175 and RAPTA-C from the groups of Sadler and Dyson, respectively.17−19 It was initially anticipated that ruthenium(II) arene complexes exert their anticancer effects by interaction with DNA. However, different mechanisms of action, such as inhibition of the activity of the proteins20,21 or catalytic hydride transfer reactions in cells,22−24 were reported. Moreover, subtle ligand modifications on ruthenium(II) arene complexes can lead to different modes of action and result in significant changes in the anticancer efficacy.25−29 Therefore, the arene rings and coordination ligands of the organoruthenium complexes may provide more opportunities to design anticancer agents with different anticancer mechanisms. Polynuclear metal complexes provide a promising class of compounds for anticancer agent design and discovery, which may display additional advantages over their mononuclear counterparts by offering improved anticancer efficacies and © XXXX American Chemical Society

Figure 1. Representative anticancer ruthenium(II) complexes.

lower toxicities.30−33 The notable examples are the trinuclear platinum(II) compound BBR3464 evaluated in clinical trials and dinuclear rhodium(II) carboxylate complexes with significant in vitro and in vivo anticancer activities by inhibiting nucleic acid and protein synthesis.2,34 Thus, the strategy to modify the action modes and improve the therapeutic properties Received: December 11, 2018

A

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry

Figure 2. Ruthenium(II) complexes 1−4 studied in this work.

Scheme 1. Preparation of Ligands L1 and L2 and Complexes 1−4a

a

Reagents and conditions: (a) MeOH, rt, 24 h; (b) [(arene)RuCl2]2, DCM, rt, 12 h.

Table 1. log POW and Cytotoxicity Data for 1−4 IC50 values (μM) compound

log POW

T24a

MCF-7b

A549c

A549/Cisd

RFe

1 2 3 4 cisplatin

−1.73 ± 0.15 −1.32 ± 0.08 −1.21 ± 0.11 −0.96 ± 0.07 −2.03 ± 0.47f

20.6 ± 1.9 28.0 ± 2.5 29.5 ± 1.7 8.4 ± 1.0 13.5 ± 0.3

23.4 ± 2.1 27.7 ± 1.3 25.8 ± 1.5 8.6 ± 0.6 9.5 ± 0.6

15.5 ± 1.3 19.6 ± 1.2 22.4 ± 2.1 7.5 ± 0.5 9.6 ± 0.3

15.7 ± 1.1 18.6 ± 3.6 24.0 ± 0.9 7.8 ± 0.6 44.0 ± 1.7

1.01 0.95 1.07 1.04 4.58

a

Human bladder carcinoma cell line. bHuman breast carcinoma cell line. cHuman non-small-cell lung cancer cell line. dCisplatin-resistant nonsmall-cell lung cancer cell line. eResistant factor (RF) defined as IC50 in A549/Cis/IC50 in A549. fCited from ref 36.

with electrospray ionization mass spectrometry (ESI-MS; Figures S1−S12). The ESI-MS spectra of compounds 1−4 showed the highest isotope at 459.08, 509.04, 487.11, and 537.13, respectively, corresponding to the dication species of [(arene)2Ru2L]2+ (L = L1 or L2). All of the spectral data were compatible with the proposed molecular structures of complexes 1−4. Notably, complexes 1−4 showed similar absorption spectra, except that the absorption peaks of complexes 2 and 4 at about 350 nm showed red shifts in comparison to complexes 1 and 3 (Figure S13). The stability of complexes 1−4 in an aqueous solution was evaluated by UV−vis spectroscopy (Figure S14). No detectable changes of the absorption bands of complexes 1−4 were observed after 12 h, demonstrating that

of complexes by engineering polynuclear scaffolds is an effective way for anticancer drug discovery.35 Here, we report a series of dinuclear ruthenium(II) arene complexes with two metal centers bridged by a relatively rigid aromatic chain that are expected to exert their activities by different anticancer mechanisms.

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RESULTS AND DISCUSSION Synthesis and Characterization. Complexes 1−4 (Figure 2) with different arene and iminopyridyl ligands were prepared by following the procedure shown in Scheme 1. The resulting dinuclear ruthenium(II) complexes were characterized by elemental analysis and 1H and 13C NMR spectroscopy along B

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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the bond distances for complexes 1−4, except that the distances of the Ru−Cl bonds of complexes 3 and 4 are longer than those of complexes 1 and 2, which is due to the increased electron density at RuII from the aromatic ligand of complexes 3 and 4. The EPSs exhibited that the Cl atoms are relatively electron-rich sites of complexes 1−4 (Figure 3). However, the color distributions in the EPSs of complexes 1−4 are rather flat, indicating that these dinuclear ruthenium(II) complexes are relatively nonpolar. In Vitro Cytotoxicity. The antiproliferative activities against three human cancer cell lines (A549, MCF-7, and T24 cancer cell lines) have been investigated for all of the complexes together with cisplatin as a positive agent by the 3-(4,5dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) method. IC50 values were determined from dosesurvival curves (Table 1 and Figure S15). According to the IC50 values, complexes 1−4 showed moderate antiproliferative activities against the tested cancer cell lines. Notably, complex 4 exhibited considerable cytotoxicity against the cancer cell lines with the IC50 range from 7.5 to 8.6 μM, which displayed comparable or superior cytotoxicity to that of cisplatin (range from 9.5 to 13.5 μM for cisplatin). In terms of the strong antiproliferative effects on the tested cancer cell lines, complexes 1−4 were further studied to determine whether these dinuclear ruthenium(II) arene complexes can overcome cisplatin resistance. As shown in Table 1, the IC50 value of cisplatin against A549/Cis was increased to 44.0 μM, whereas the cytotoxicity of complexes 1− 4 against cisplatin-sensitive A549 and cisplatin-resistant A549/ Cis cells was quite similar to that of almost the same resistance factor values ranging from 0.95 to 1.07, indicating that these ruthenium(II) complexes can overcome cisplatin resistance. Significantly, complex 4 (7.8 μM) is 5.6-fold as potent as cisplatin (44.0 μM) against A549/Cis cells. Interaction with DNA. DNA is a critical therapeutic target for metal-based anticancer agents, which may interact with the DNA through covalent binding, intercalation, groove binding, or

these complexes were stable to aquation. Moreover, the lipophilicity of complexes 1−4 was determined using the shake-flask method (Table 1). The log POW values for complexes 1−4 range from −1.73 to −0.96, showing that all compounds were relatively hydrophilic. Electrostatic Potential Surface (EPS). Density functional theory (DFT) calculations were applied to obtain the structural and electronic information on complexes 1−4. The optimized geometries depicted in Figure 3 show very minor differences in

Figure 3. Structures and EPSs for dinuclear ruthenium(II) complexes 1−4. EPS maps (from −0.10 au in red to +0.20 au in blue) drawn onto an electron density isosurface (0.004 au) for the same compounds.

Figure 4. UV−vis specta of complexes 1−4 (25 μM) in Tris-HCl with increasing concentrations of CT-DNA (0−120 μM) at room temperature. Inset: Plot of A0/(A − A0) versus 1/[DNA]: (a) complex 1; (b) complex 2; (c) complex 3; d) complex 4. C

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Figure 5. Fluorescence (λexc = 540 nm) quenching curves of EtBr bound to DNA in the presence of complexes 1−4. [DNA] = 25 μM, [EtBr] = 20 μM, and [complex] = 0−33 μM.

Figure 6. Molecular docked model of complexes 1−4 with DNA (PDB 1BNA): (a) 1; (b) 2; (c) 3; (d) 4.

electrostatic interactions.37−43 Thus, the interaction of complexes 1−4 with DNA was studied by UV−vis absorbance and fluorescence spectroscopy. The electronic absorption titration was carried out with results shown in Figure 4. Obviously, the hypochromisms at 207−221 nm and bathochromic shifts (10−14 nm) were observed for all of the complexes with increasing concentrations of CT-DNA (Table S1), probably because of the groove binding as well as intercalation of the dinuclear ruthenium(II) complexes with CT-DNA.44,45 The binding constant Kb was calculated using the Benesi−Hildebrand equation (Table S1). These ruthenium(II) complexes have affinity constants of ∼104 M−1 for DNA. Among them, complexes 2 and 4 with iminoquinoline ligands showed enhanced binding efficacy compared with complexes 1 and 3 with an iminopyridine ligand. In order to study the DNA binding mode further, a competitive binding experiment was carried out by monitoring the emission intensity of DNA-bound ethidium bromide (EtBr) upon the addition of complexes 1−4. Both DNA groove binders and intercalators reduce the fluorescence intensity, but the intercalators can reduce the intensity significantly because of the replacement of EtBr, while the intensity reduction is moderate for groove binders.46 As shown in Figure 5, the observed quenching upon the addition of complexes 1−4 is 22.1%, 50.5%, 43.5%, and 84.5%, respectively. Notably, the fluorescence intensity of the DNA−EtBr complex was reduced more

remarkably by complexes 2 and 4 than by complexes 1 and 3, probably attributed to the intercalation effect of the quinoline ligand. This study hints that complex 1 may interact with DNA mainly through groove binding, while complexes 2−4 can be proposed to interact via the DNA groove binding in addition to intercalation interactions. Molecular docking studies were made to elucidate the DNA binding mode of complexes 1−4. The optimized structures of these complexes were docked with a B-DNA structure (PDB 1BNA) using AutoDock 4.2. The most stable binding conformations of these dinuclear ruthenium(II) complexes and DNA show that complexes 1−4 fit into the minor groove of the DNA in a parallel manner with respect to the DNA backbone (Figure 6), suggesting that complexes 1−4 interact with the minor groove of DNA. Because atomic force microscopy (AFM) is a useful tool to study the interaction of DNA with small molecules and offer an intuitive and direct way to observe the morphological changes of DNA, it was applied to monitor the interaction of complexes 1− 4 with plasmid DNA (pBR322). As shown in Figure S16, DNA condensation was observed after 24 h of incubation of the ruthenium(II) complex with pBR322 plasmid DNA, which is mainly attributed to the insertion of the dinuclear ruthenium(II) complex into DNA chains. With the purpose of confirming the binding mode of these complexes with DNA, agarose gel electrophoresis was applied to D

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Figure 7. Gel electrophoretic mobility pattern of pBR322 plasmid DNA incubated with various concentrations of ruthenium(II) complexes. Lanes 1− 9 (0, 10, 20, 40, 80, 160, 320, 640, and 1280 μM) + DNA: (a) 1; (b) 2; (c) 3; (d) 4.

study the interaction of complexes 1−4 with pBR322 plasmid DNA (Figure 7). A decrease in the rate of migration for closedcircular DNA (form I) was obviously observed for complexes 1 and 2, indicating the covalent binding of the complexes with pBR322 DNA.47 However, no migration was observed for complexes 3 and 4. Moreover, the plasmid DNA gradually disappeared with increasing concentrations of complexes 1−4, indicating that these complexes can inhibit the intercalation of EtBr in plasmid DNA at high concentrations, especially complexes 2 and 4, which is in accordance with the results of an EtBr competitive binding experiment. This study indicates that complexes 1 and 2 can covalently bind to DNA, while complexes 3 and 4 interact with DNA through intercalation interactions. In summary, the interactions of complexes 1−4 with DNA have been investigated by means of UV−vis, EtBr competition, computational docking, AFM, and agarose gel electrophoresis. The results show that complex 1 may interact with DNA through the minor groove binding and covalent binding, while complex 2 binds covalently and intercalatively to DNA, simultaneously. Complexes 3 and 4 interact with DNA through the minor groove binding and intercalation interactions. ROS Generation. Ruthenium(II) arene complexes have been reported to induce ROS generation in cancer cells, which is responsible for the observed cytotoxicity.48 Thus, the levels of ROS in A549 cells induced by complexes 1−4 were determined by flow cytometry. As shown in Figure S17, the dinuclear ruthenium(II) complexes markedly increased the ROS levels in A549 cells, especially complex 4 (Figure 8). The relative order of the ROS levels induced by these complexes is 4 ≫ 3 > 2 > 1. To further elucidate the type of ROS induced by complex 4, some ROS scavengers, such as superoxide dismutase (SOD), tert-butyl alcohol (TBA), and N-acetyl-L-cysteine (NAC), were cotreated with complex 4. As shown in Figures S18 and S19, SOD and TBA treatment inhibited the ROS levels in the cells, indicating that both superoxide (•O2−) and hydroxyl (•OH) radicals were generated. Moreover, the addition of NAC (nonspecific antioxidant) could greatly reduce ROS generation. Taken together, this study suggests that both •O2− and •OH radicals may be generated in A549 cells treated with complex 4. Frontier Molecular Orbital. The highest occupied molecular orbitals (HOMOs) and lowest unoccupied molecular orbitals (LUMOs) of complexes 1−4 were calculated (Figure 9). The HOMOs are localized largely on one Ru atom, Cl− ions, and arene groups in complexes 1−4, while the LUMOs are

Figure 8. Quantification of the flow cytometric results in Figure S17 showing the percentage of cells with increased intracellular DCF oxidation compared to that of control cells. The data are representative of three independent experiments. The results are mean ± SD (n = 3). (**) p < 0.01 compared with the value of the control.

localized largely on ligand L1 or L2. The HOMO−LUMO energy gaps of these complexes vary as 1 (1.91 eV) > 3 (1.79 eV) ≈ 2 (1.75 eV) > 4 (1.62 eV), suggesting that electrons in complex 4 are easier to excite from the HOMO to LUMO than in complexes 1−3. The lower HOMO−LUMO energy gap in complex 4 may result in easier electron transfer and ROS generation, which is in accordance with the data of ROS generation. The study implicates that electron transfer of these complexes can be responsible for ROS generation. Western Blot. To further investigate the impact of ROS on the cancer cells, we applied a Western blot technique to study the expression of the transcription factor Nrf2, which can regulate the expression of some antioxidant genes.49 As shown in Figure 10, the levels of Nrf2 protein were increased after the treatment of complexes 1−4. Because some reported ROS induced by chemotherapeutic agents can trigger endoplasmic reticulum (ER) stress via the IRE1/XPB-1s pathway,49 the expression of two crucial ER stress markers, XBP-1s and CHOP, was examined as well. It was observed that complexes 1−4 increased the accumulation of XBP-1s and CHOP, indicating that these complexes can induce ER stress in cancer cells. Cell Cycle Distribution. The influence of complex 4 on the cell cycle progression of A549 and A549/Cis cells was evaluated in the presence and absence of NAC by flow cytometry. As shown in Figure S20, complex 4 induced the cell cycle at the S phase in both A549 and A549/Cis cells with respect to the untreated control. Moreover, the addition of NAC had a negligible influence on the cell cycle distribution in A549/Cis E

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Figure 9. Frontier molecular orbital diagram and energy profiles for the HOMOs and LUMOs of complexes 1−4.

Figure 10. (a) Western blot analysis of the ER stress protein markers in A549 cells after treatment with complexes 1−4 at 20 μM. The data are representative of three independent experiments. (b) Densitometric analysis of the expression of proteins normalized with GAPDH. The relative expression of each protein was represented by the density of the protein band/density of the GAPDH band. (**) p < 0.01 compared with the value of control.

Figure 11. Flow cytometry analysis for apoptosis of A549 and A549/Cis cells induced by complex 4 and cisplatin.

Apoptosis Study. Apoptosis is a programmed cell death process in multicellular organisms, which is the main cell death pathway induced by metal-based anticancer agents. With the purpose of investigating the cause of the different cytotoxicities of complex 4 and cisplatin, equimolar concentrations (20 μM) were chosen for a comparison to elucidate the underlying

cells. As for A549 cells, the percentage of the cells arrested at the S phase was slightly decreased after the addition of NAC. Taken together, complex 4 inhibited the cell cycle in the S phase in A549 and A549/Cis cells compared with the control cells, and the addition of NAC had little effect on the cell cycle distribution. F

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry mechanisms. Besides, an extra concentration (10 μM) was evaluated for complex 4. As shown in Figure 11, both complex 4 and cisplatin induced an improved incidence of early-to-latestage apoptosis in A549 cells compared with untreated cells (control). Moreover, the effect of complex 4 (43.5%) on the cancer cell apoptosis is stronger than that of cisplatin (30.6%) at 20 μM in A549 cells. As for A549/Cis cells, the population of apoptotic cells induced by cisplatin (20.6%) decreased, while complex 4 produced a population of apoptotic cells (42.4%) similar to that in the A549 cells (43.5%) at 20 μM, which is in accordance with the cytotoxicity results. Besides, the apoptotic rates of the A549 cells were increased with increasing concentration of complex 4, suggesting that complex 4 could induce cancer cell death through an apoptotic pathway in a concentration-dependent manner. To explore the role of ROS in apoptosis induced by complex 4, the A549 and A549/Cis cells were preincubated with the antioxidant NAC before exposure to complex 4. As shown in Figure S21, NAC preincubation caused an approximate 12% and 15% decrease in the percentage of apoptotic cells in A549 and A549/Cis cells, respectively, indicating that ROS plays a crucial role in cell apoptosis.

obtained from Jiangsu KeyGEN BioTECH Co. (China). Cell apoptosis and ROS experiments were measured by flow cytometry (FAC Scan, Becton Dickenson). General Procedure for the Synthesis of Complexes 1−4. A methanol (MeOH) solution (50 mL) of 4,4′-methylenedianiline (1.2 mmol) and 2-quinolinecarbaldehyde or 2-pyridinecarbaldehyde (2.4 mmol) was stirred at room temperature for 24 h. Then RuL2Cl2 (1.0 mM) was added, and the resulting mixture was stirred at room temperature for 12 h. The solvent was then removed by evaporation under reduced pressure. The crude product was collected and recrystallized from MeOH. Complex 1. Yield: 0.45 g (45.5%); yellow-brown powder. Anal. Calcd for C45H48Cl4N4Ru2: C, 54.66; H, 4.89; N, 5.67. Found: C, 54.63; H, 4.79; N, 5.58; ESI-MS: m/z 459.08 ([M/2 − Cl]+). 1H NMR (600 MHz, CD3OD): δ 1.08−1.09 (m, 12H), 2.25 (s, 6H), 2.58−2.62 (m, 2H), 4.28 (s, 2H), 5.52−5.53 (d, 2H, J = 5.8 Hz), 5.62−5.63 (d, 2H, J = 6.1 Hz), 5.70−5.71 (d, 2H, J = 6.1 Hz), 6.01−6.02 (d, 2H, J = 6.1 Hz), 7.56−7.58 (m, 4H), 7.80−7.82 (d, 4H, J = 8.2 Hz), 7.84−7.86 (m, 2H), 8.25−8.30 (m, 4H), 8.81−8.82 (m, 2H), 9.53−9.55 (m, 2H). 13C NMR (150 MHz, CD3OD): δ 17.58, 20.90, 21.01, 31.02, 40.48, 85.39, 85.56, 86.31, 86.80, 103.99, 106.28, 122.64, 128.86, 129.81, 129.92, 139.70, 143.13, 150.60, 154.93, 155.84, 166.77. Complex 2. Yield: 0.41 g (37.7%); yellow-brown powder. Anal. Calcd for C53H52Cl4N4Ru2: C, 58.46; H, 4.81; N, 5.15. Found: C, 58.60; H, 4.74; N, 5.23. ESI-MS: m/z 509.04 ([M/2 − Cl]+). 1H NMR (600 MHz, CD3OD): δ 0.86−0.87 (d, 6H, J = 6.8 Hz), 1.01−1.02 (d, 6H, J = 6.9 Hz), 2.31−2.32 (d, 6H, J = 2.9 Hz), 2.38−2.45 (m, 2H), 4.33−4.34 (d, 2H, J = 2.1 Hz), 5.36−5.37 (d, 2H, J = 6.1 Hz), 5.77− 5.78 (d, 2H, J = 6.1 Hz), 5.88−5.89 (d, 2H, J = 6.2 Hz), 6.04−6.05 (d, 2H, J = 6.2 Hz), 7.64−7.66 (m, 4H), 8.00−8.02 (m, 6H), 8.20−8.23 (m, 2H), 8.26−8.28 (d, 2H, J = 8.2 Hz), 8.34−8.35 (m, 2H, J = 7.3 Hz), 8.82−8.84 (m, 4H), 9.08−9.09 (d, 2H, J = 3.7 Hz). 13C NMR (150 MHz, CD3OD): δ 17.59, 20.47, 21.22, 31.06, 40.55, 85.30, 85.93, 86.70, 86.87, 105.08, 106.22, 122.63, 129.02, 129.14, 129.78, 130.04, 130.31, 133.18, 140.78, 143.47, 149.04, 150.93, 155.90, 167.39, 167.41. Complex 3. Yield: 0.52 g (49.8%); yellow-brown powder. Anal. Calcd for C49H56Cl4N4Ru2: C, 56.32; H, 5.40; N, 5.36. Found: C, 56.43; H, 5.29; N, 5.31. ESI-MS: m/z 487.11 ([M/2 − Cl]+). 1H NMR (600 MHz, CD3OD): δ 1.93−1.94 (m, 36H), 4.23 (s, 2H), 7.53−7.55 (m, 4H), 7.74−7.75 (d, 2H, J = 8.3 Hz), 7.84−7.87 (m, 2H), 8.19−8.24 (m, 4H), 8.66−8.67 (d, 4H, J = 3.4 Hz), 9.00−9.01 (d, 2H, J = 5.5 Hz). 13 C NMR (150 MHz, CD3OD): δ 14.19, 40.36, 96.64, 122.63, 128.67, 129.49, 129.85, 129.88, 139.19, 143.20,143.23, 149.26, 153.23, 155.04, 167.20. Complex 4. Yield: 0.48 g (42.0%), yellow-brown powder. Anal. Calcd for C57H60Cl4N4Ru2: C, 59.79; H, 5.28; N, 4.89. Found: C, 59.68; H, 5.37; N, 4.97. ESI-MS: m/z = 537.13 ([M/2 − Cl]+). 1H NMR (600 MHz, CD3OD): δ 1.81 (s, 36H), 4.22−4.24 (m, 2H), 7.55−7.56 (d, 4H, J = 7.6 Hz), 7.89−7.93 (m, 2H), 7.98−8.00 (m, 4H), 8.07−8.09 (m, 2H), 8.15−8.18 (m, 2H), 8.23−8.27 (m, 2H), 8.48− 8.50 (d, 2H, J = 8.8 Hz), 8.66−8.69 (d, 2H, J = 8.5 Hz), 8.88−8.91 (m, 2H). 13C NMR (150 MHz, CD3OD): δ 14.37, 40.42, 96.95, 123.24, 124.29, 128.84, 128.85, 129.01, 129.99, 130.02, 130.26, 132.24, 140.03, 140.05, 143.71, 143.76, 148.93, 167.35, 167.40. log POW Determination. The log POW determination of complexes 1−4 was conducted using the shake-flask method. Excess 1−4 were dissolved in double-distilled water presaturated with n-octanol for 24 h at 37 °C. The solution was filtered to remove undissolved ruthenium complexes. Subsequently, the solution was added to an equal volume of n-octanol (presaturated with water). The heterogeneous mixture was shaken vigorously for 2 h before centrifuging for 15 min to achieve phase separation. The initial and final concentrations of compounds in an aqueous phase were determined by the UV−vis spectrum method, and the water−octanol partition coefficients (log POW) were calculated. DFT Calculation. All calculations were performed using the Gaussian 09 program package.51 Full geometry optimizations were carried out for complexes 1−4 at the M06-L/6-31G*//LanL2DZ level in the gas phase at 310.15 K and 1 atm.52,53 Docking Study. Docking studies were carried out using AutoDock 4.2.54 The crystal structure of B-DNA (PDB 1BNA) has been taken

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CONCLUSION In summary, four novel dinuclear ruthenium(II) arene complexes were designed and synthesized. In vitro tests indicated that the resulting ruthenium(II) complexes displayed moderate antiproliferative activity against the tested cancer cells, while the cytotoxicity of complex 4 is superior or comparable to that of cisplatin. Further experimental and computational studies showed that the ruthenium(II) complexes have affinity constants of ∼104 M−1 for DNA by interacting with DNA mainly through minor groove binding and/or intercalation interactions. The agarose gel electrophoresis study indicates that complexes 1 and 2 can covalently bind to DNA, while complexes 3 and 4 interact with DNA through intercalation interactions. Moreover, complexes 1−4 can generate significant levels of ROS and further induce ER stress. Hence, we proposed that these dinuclear ruthenium(II) complexes induce cell death through DNA binding and a ROS-mediated ER stress pathway, which is the first example of an organometallic ruthenium(II) arene complex to induce ER stress as well as DNA interaction. The findings of the present study help to unravel the mechanisms of action of ruthenium(II) arene complexes. It is noteworthy that complexes 1−4 have no cross-resistance with cisplatin due to the different mechanisms of action. Overall, this kind of dinuclear ruthenium(II) arene complex has unique biological characteristics and is a promising model for new anticancer drug development.

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EXPERIMENTAL SECTION

Materials and Measurements. All chemicals and solvents were of analytical reagent grade and were used without further purification. Ligands L1 and L2 were prepared according to previous reports.50 1H and 13C NMR spectra were measured on a Bruker Avance III-HD 600 MHz spectrometer. UV−vis spectra and kinetic traces were recorded on a Shimadzu UV2600 instrument. The concentration of DNA in the base pairs was determined by UV−vis absorbance at 260 nm taking ε260 nm as 13100 M−1 cm−1. Fluorescence measurements were performed using a FluoroMax-4 fluorometer. Scans were run at room temperature with excitation and emission slit widths of 2.5 nm. Mass spectrometry was measured by an Agilent 6224 ESI/TOF MS instrument. Elemental analysis of C, H, and N used a Vario MICRO CHNOS elemental analyzer (Elementar). Human cancer cells were G

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry

by centrifugation (5 min, 25 °C, 2000 rpm). Then, the cells were washed twice with cold water and resuspended in a binding buffer (10 mM HEPES, 140 mM NaCl, 2.5 mM CaCl2, pH 7.4). The cells were stained with 5 μL of Annexin V-FITC and then with 5 μL of propidium iodide (PI; 20 μg/mL) for 15 min in the dark at room temperature. The fluorescence of the cells was detected by an Annexin V-FITC apoptosis detection kit (Roche) according to the manufacturer’s protocol, and the cells were quantified by system software (Cell Quest; BD Biosciences). MTT Assay. The cytotoxicities of complexes 1−4 and cisplatin (positive control) against A549, MCF-7, T24, and A549/Cis cells were determined by means of MTT assay. Cells (105/well) with better vitality were seeded in 96-well plates. The compounds were dissolved by DMF (40 mM) and diluted with a medium to various concentrations (0.16, 0.63, 2.5, 10, 40, and 160 μM; the final concentration of DMF was less than 0.4%). Cisplatin was diluted to the tested concentration with a medium (0.16, 0.63, 2.5, 10, 40, and 160 μM). As for the negative control, 0.4% DMF was added. The cells were incubated in the dark for 72 h. After that, the cells were stained with MTT (5 mg/mL) for another 5 h, and then the medium was thrown away and replaced by 150 mL of dimethyl sulfoxide. The inhibition of cell growth induced by the tested complexes was detected by measuring the absorbance of each well at 570/630 nm using an enzyme-labeling instrument. The IC50 values were calculated by SPSS software after three parallel experiments. Cell Cycle Measurement. A549 and A549/Cis cells were transferred into 6-well plates, with a density of 105/well, and cultured overnight at 37 °C. Then, complex 4 was dissolved by DMF (10 mM) and diluted with a medium to 20 μM (the final concentration of DMF was 0.2%). As for the negative control, the same volume of DMF was added. For NAC experiments, the cells were preincubated with a NAC solution (2 mM in a growth medium) for 30 min. All adherent and floating cells were collected and washed twice with phosphate-buffered saline. Then, the cells were fixed with 70% ethanol at 4 °C for 24 h. After being centrifuged, the cells were stained with a 50 μg/mL PI solution containing 100 μg/mL RNase for 0.5 h at 37 °C. The samples were measured by flow cytometry (FAC Scan, Becton Dickenson) using Cell Quest software and recording PI in the FL2 channel. ROS Detection. A549 cells were cultured in black 96-well culture plates (20000 cells/well) for 24 h. For a ROS quencher experiment, the cells were preincubated with SOD (1500 units/mL), TBA (10 mM), and NAC (10 mM) for 30 min. Thereafter, complexes 1−4 were dissolved by DMF (10 mM) and diluted with a medium to 20 μM (the final concentration of DMF was 0.2%). The ROS levels induced by arene−ruthenium complexes increased initially but decreased with time.26 Hence, 6 h of incubation was selected for the study. As for the negative control, the same volume of DMF was added. Concentrated hydrogen peroxide was diluted to 20 μM in a RPMI-1640 medium with 10% FBS and was added to the cells as a positive control. As for the negative control, only a RPMI-1640 medium was added. Then, the cells were incubated with a ROS probe, carboxy-H2DCFDA (20 μM), for 30 min at 37 °C in the dark. The fluorescence was measured with a microplate reader using an excitation at 485 nm and an emission at 535 nm. All experiments were done in triplicate, and data are depicted as mean ± standard deviation (SD). Western Blot. A549 cells were grown in a 6-well plate at a density of 2 × 105 cells/well and cultured until the cell density reached 80%. Complexes 1−4 were dissolved by DMF (10 mM) and diluted in media to give the required concentration (20 μM) for the addition to the cells (the final concentration of DMF was 0.2%), and the cells were cultured for 24 h at 37 °C. As for the negative control, 0.2% DMF was added. A549 cells were lysed in a cell lysis buffer and collected by centrifugation at 13 000 rpm for 20 min at 4 °C. Proteins from cell lysates were separated by 8−12% sodium dodecyl sulfate−polyacrylamide gel electrophoresis (SDS−PAGE) and transferred onto a poly(vinylidine difluoride) membrane (Amersham Biosciences). The membrane was blocked with PBS with Tween 20 containing 5% nonfat dry milk for 1 h and further incubated with primary antibodies (Nrf-2, XBP-1s, and CHOP) overnight at 4 °C under gentle shaking. After that, the membrane was incubated with the secondary antibody (1:2000) for 1 h at room temperature (25 °C). Protein blots were detected with chemiluminescence reagent (Thermo Fischer Scientifics Ltd.).

from the protein data bank.55 Visualization of the docked molecules has been made by the software PyMOL. The docking simulation was performed with the Lamarckian genetic algorithm for as much as 150 docking runs. Each run of the docking operation was terminated after a maximum of 2500000 energy evaluations. During docking studies, the DNA structure was kept rigid. Rotation in the complexes 1−4 was permitted about all single bonds. Electronic Absorption Titration Studies. Absorption spectral titration experiments were performed by electronic spectroscopy under physiological conditions (5 mM Tris-HCl/10 mM NaCl buffer solution, pH 7.2) with 5 min of equilibration time. Spectra were collected from 195 to 600 nm after the successive addition of CT-DNA (0−120 μM) into a 3 mL solution of the complex (25 μM). Because the electronic absorption of N,N-dimethylformamide (DMF) in 200−300 nm may overlap with the absorbance of compounds 1−4, they were initially dissolved in MeOH (5 mM) and diluted with buffer to the required concentration (the final concentration of MeOH was 0.5%). Fluorescence Emission Spectrometry. The experiments were carried out by the addition of compounds 1−4 to samples containing 25 μM CT-DNA (nucleotide) and 20 μM EtBr in a Tris buffer solution (5 mM Tris-HCl/10 mM NaCl buffer solution, pH 7.2). The stock solutions of the complexes were prepared using MeOH (5 mM), and they were further diluted to the required concentration using a Tris buffer solution (the final concentration of MeOH was less than 2%). The influence of the addition of complexes 1−4 (0−33 μM) to the EtBr−DNA mixture was measured by recording variations of the fluorescence emission spectra with an excitation at 540 nm and an emission between 550 and 720 nm. AFM. The pBR322 plasmid DNA (200 ng/μL) was diluted to 5 ng/ μL with a 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer (40 mM HEPES, 5 mM MgCl2, pH 7.2). Opencircular and linear plasmid molecules were obtained after incubation of a solution of supercoiled pBR322 plasmid DNA (5 ng/μL) at 333 K for 30 min. Complexes 1−4 were first dissolved in DMF (10 mM) and then diluted to 0.5 mM with a HEPES buffer. The solutions of complexes 1− 4 (25 μL) and pBR322 (25 μL) were mixed together (the final concentration of DMF was 2.5%). As for the negative control, 2.5% DMF was added. After 24 h of incubation, the mixture was dropped onto freshly cleaved mica. Then the samples were rinsed for 10 s with deionized water and dried with nitrogen gas. The images were obtained in air at room temperature on areas of 1.0 × 1.0 μm2. Gel Electrophoresis Study. DNA binding properties of complexes 1−4 were investigated by agarose gel electrophoresis, and pBR322 plasmid DNA (50 ng/mL) was used as the target. Complexes 1−4 were first dissolved in DMF (10 mM) and then diluted to the desired concentrations with Tris-H3PO4 (100 mm) buffer. pBR322 DNA (5 μL) and complexes 1−4 (5 μL) were added to each tube, and the mixtures of ruthenium complexes and pBR322 plasmid DNA were then incubated at 37 °C for 24 h. As for the negative control, the same volume of DMF was added. Subsequently, the agarose gel [made up to 1% (w/v)] was prepared with a TA buffer (50 mm Tris-acetate, pH 7.4). The mixtures with a loading buffer (1 mL) were submitted to electrophoresis in agarose gel in a TA buffer at 100 V for 90 min. Agarose gels were then dyed with EtBr (0.5 mg/L) for 20 min. Bands were imaged by using a Molecular Imager (Bio-Rad, USA) under UV light. Apoptosis Analysis by Flow Cytometry. On the basis of the IC50 values of our compounds, 10 and 20 μM were selected for the apoptosis studies, which are 1.3−2.7-fold of the IC50 values of complex 4 against A549 and A549/Cis cells. A549 and A549/Cis cells were cultured in a RPMI-1640 medium with 10% fetal bovine serum (FBS). All media were also supplemented with 100 mg/mL penicillin and 100 mg/mL streptomycin. Cells were grown in a 6-well plate at a density of 2 × 105 cells/well and cultured overnight. Complex 4 was dissolved in DMF (10 mM) and diluted with a medium to the desired concentrations (the final concentration of DMF was 0.1% for 10 μM and 0.2% for 20 μM). Cisplatin was diluted to 20 μM with a medium. As for the negative control, the same volume of DMF was added. For NAC experiments, cells were preincubated with a NAC solution (2 mM in a growth medium) for 30 min. After incubation for 24 h, the cells were collected H

DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX

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Inorganic Chemistry Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as the loading control. Statistics. The student’s t test was applied to evaluate the significance of the differences measured. Results were expressed as the mean ± SD and considered to be significant when p < 0.05. The data are representative of three independent experiments.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.inorgchem.8b03447. 1 H and 13C NMR, ESI-MS, and UV−vis spectra of complexes 1−4, dose-dependent cell viability curves, AFM images, analysis of the ROS levels by flow cytometry, cell cycle distribution, Flow cytometry analysis, and absorption spectroscopic properties (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Shaohua Gou: 0000-0003-0284-5480 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We are grateful to the National Natural Science Foundation of China (Grant 21601034) and Jiangsu Province Natural Science Foundation (Grant BK20160664) for financial aid to this work. Fundamental Research Funds for the Central Universities (Projects 2242016K30020 and 2242017K41025) and Priority Academic Program Development of Jiangsu Higher Education Institutions for the construction of fundamental facilities are also appreciated.

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REFERENCES

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DOI: 10.1021/acs.inorgchem.8b03447 Inorg. Chem. XXXX, XXX, XXX−XXX