Dalton Transactions View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

PAPER

Cite this: Dalton Trans., 2014, 43, 17303

View Journal | View Issue

Dual inhibition of topoisomerases I and IIα by ruthenium(II) complexes containing asymmetric tridentate ligands† Kejie Du,a,b Jiewen Liang,a Yi Wang,a Junfeng Kou,a Chen Qian,a Liangnian Jia and Hui Chao*a Five novel ruthenium(II) complexes, [Ru(dtzp)(dppt)]2+ (1), [Ru(dtzp)( pti)]2+ (2), [Ru(dtzp)( ptn)]2+ (3), [Ru(dtzp)( pta)]2+ (4) and [Ru(dtzp)( ptp)]2+ (5) (where dtzp = 2,6-di(thiazol-2-yl)pyridine, dppt = 3-(1,10phenanthroline-2-yl)-5,6-diphenyl-as-triazine), pti = 3-(1,10-phenanthroline-2-yl)-as-triazino-[5,6-f ]isatin, ptn = 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]naphthalene, pta = 3-(1,10-phenanthroline-2yl)-as-triazino[5,6-f ]acenaphthylene, and ptp = 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]-phenanthrene), were synthesised and characterised. The structures of complexes 3–5 were determined by X-ray diffraction. The DNA binding behaviours of the complexes were studied by spectroscopic and viscosity measurements. The results suggested that the Ru(II) complexes, except for complex 1, bind to DNA in an intercalative mode. Topoisomerase inhibition and DNA strand passage assay confirmed that Ru(II) complexes 3, 4, and 5 acted as efficient dual inhibitors of topoisomerases I and IIα. In vitro cytotoxicity assays

Received 15th July 2014, Accepted 23rd September 2014 DOI: 10.1039/c4dt02142h www.rsc.org/dalton

indicated that these complexes exhibited anticancer activity against various cancer cell lines. Ruthenium(II) complexes were confirmed to preferentially accumulate in the nucleus of cancer cells and induced DNA damage. Flow cytometric analysis and AO/EB staining assays indicated that these complexes induced cell apoptosis. With the loss of the mitochondrial membrane potential, the Ru(II) complexes induce apoptosis via the mitochondrial pathway.

Introduction DNA topoisomerases are essential enzymes that control and modify the topological states of DNA to maintain the chromosome superstructure and integrity.1,2 In general, topoisomerases have been shown to be overexpressed in cancer cells; thus, topoisomerases are important targets of anticancer drugs.3 To date, some topoisomerase inhibitors, such as camptothecin (CPT) and its derivatives that target topoisomerase I (Topo I)4,5 and etoposide (VP-16),6 doxorubicin7 and mitoxantrone,8 which target topoisomerase II (Topo II), have been clinically used as potent anticancer drugs. Although these agents have achieved great success in clinical cancer

a MOE Key Laboratory of Bioinorganic and Synthetic Chemistry, State Key Laboratory of Optoelectronic Materials and Technologies, School of Chemistry and Chemical Engineering, Sun Yat-Sen University, Guangzhou 510275, P. R. China. E-mail: [email protected] b School of Chemistry and Chemical Engineering, University of South China, Hengyang 421001, P. R. China † Electronic supplementary information (ESI) available: Scheme S1, Fig. S1–S2, Tables S1–S2. CCDC 879756–879758 for complexes 3–5. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c4dt02142h

This journal is © The Royal Society of Chemistry 2014

chemotherapy, single inhibitors of topoisomerases are limited by several important negative consequences, such as druginduced secondary malignancies.9 Liu et al. also reported that Topo II-based chemotherapy, such as VP-16, is directly associated with treatment-related acute myeloid leukemia.10 In addition, the emergence of drug-resistant tumour cells remains a major problem, which results in the failure of longterm clinical therapies using Topo I- or Topo II-targeting drugs. To overcome the disadvantages of Topo I or II inhibitors, the discovery of novel dual inhibitors has recently emerged as a promising field to identify better antitumour agents. Over the last decade, several classes of dual topoisomerase I and II inhibitors have been identified and described.11,12 Some of these inhibitors, such as acridine DACA (XR5000),13 TAS-103,14 Tafluposide (F11782),15 Batracylin (NSC320846)16 and XR5944,17 have been evaluated in clinical trials. However, most studies have focused on organic compounds and, to a far lesser extent, on metal complexes.18–20 Due to varied coordination forms and their rich photochemical properties, ruthenium(II) complexes have recently received considerable attention as topoisomerase inhibitors.21–23 In our previous study, we found that some ruthenium(II) complexes exhibited

Dalton Trans., 2014, 43, 17303–17316 | 17303

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Paper

Scheme 1

Dalton Transactions

Chemical structures of Ru(II) complexes.

clear dual Topo I and II inhibitory activity.20,24 Most studies have primarily focused on complexes involving bidentate ligands, whereas biological activity studies of ruthenium complexes with tridentate ligands have attracted much less attention. In the present study, we have synthesised and characterised a series of ruthenium(II) complexes containing asymmetric tridentate ligands, ([Ru(dtzp)(dppt)]2+ (1), [Ru(dtzp)( pti)]2+ (2), [Ru(dtzp)(ptn)]2+ (3), [Ru(dtzp)(pta)]2+ (4) and [Ru(dtzp)( ptp)]2+ (5) (where dtzp is 2,6-di(thiazol-2-yl)pyridine, dppt is 3-(1,10-phenanthroline-2-yl)-5,6-diphenyl-as-triazine), pti is 3-(1,10-phenanthroline-2-yl)-as-triazino-[5,6-f]isatin, ptn is 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]naphthalene, pta is 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]acenaphthylene, and ptp is 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]-phenanthrene, Scheme 1), bearing tridentate ligands with different degrees of planarity. DNA binding, topoisomerase inhibitory activity and cytotoxic properties of the five Ru(II) polypyridyl complexes were determined. The experimental results indicated that complexes 4 and 5, which contain greater planar ligands, demonstrated excellent dual Topo I and IIα inhibitory activity and cytotoxicity. The related cellular uptake, DNA damage and apoptosis in HeLa cells were also investigated by ICP-MS, the comet assay, AO/EB staining, Alexa Fluor®488 annexin V/propidium iodide (PI) double staining and cell cycle analysis. In addition, flow cytometric analysis using JC-1 suggested that the complexes induce apoptosis via a mitochondrial pathway.

Results and discussion Synthesis and characterisation Five ligands (dppt, pti, ptn, pta and ptp) were synthesised as previously reported by our group. The synthesis routes of Ru(II) complexes are shown in Scheme S1 (ESI†). Reactions of 1 equiv. of the corresponding ligand with Ru(dtzp)Cl3 in aqueous ethanol at reflux in the presence of an excess of triethylamine afforded the Ru(II) complexes. The five Ru(II) complexes 1–5 were purified by column chromatography and were characterized by 1H NMR spectroscopy, ES-MS and elemental analysis. In the ES-MS spectra for the complexes, only the

17304 | Dalton Trans., 2014, 43, 17303–17316

Fig. 1 ORTEP representations of the complex cations with 30% probability ellipsoid plots. (a) [Ru(dtzp)( ptn)]2+ (3), (b) [Ru(dtzp)( pta)]2+ (4), and (c) [Ru(dtzp)( ptp)]2+ (5).

signals [M − ClO4]+ and [M − 2 ClO4]2+ were observed. The measured molecular weights were consistent with expected values. Single crystals of complexes 3–5 were obtained by slowly evaporating the solvent at room temperature, and the structures were determined by X-ray crystallography. The ORTEP25 views of the cations are illustrated in Fig. 1, and the crystallographic data are summarised in Table 1. Select bond lengths and angles are provided in Table 2.

This journal is © The Royal Society of Chemistry 2014

View Article Online

Dalton Transactions

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Table 1

Paper

Crystallographic data for 3, 4 and 5

Complex

3

4

5

Formula Fw Crystal symmetry Space group T/K a/Å b/Å c/Å α/° β/° γ/° V/Å3 Z μ/mm−1 Dc/g cm−3 Unique reflns/Rint GOF on F2 Final Rindices[I > 2σ(I)]

C68H40Cl4N16O16Ru2S4 1809.34 Monoclinic C2/c 150(2) 18.6833(3) 15.8983(3) 23.6944(4) 90 94.548(2) 90 7015.9(2) 4 6.705 1.713 5209/0.0311 1.072 R1 = 0.0554 wR2 = 0.1610 R1 = 0.0599 wR2 = 0.1663

C40H26Cl2N10O8RuS2 1010.80 Triclinic P1 110(2) 12.006(2) 12.472(3) 15.858(3) 108.327(3) 95.192(3) 112.091(3) 2029.1(7) 2 0.689 1.654 8667/0.0298 1.044 R1 = 0.0379 wR2 = 0.1015 R1 = 0.0468 wR2 = 0.1101

C42H28Cl2N10O8RuS2 1036.83 Triclinic P1 293(2) 12.052(2) 12.336(3) 16.060(3) 97.37(3) 109.39(3) 109.22(3) 2049.7(7) 2 0.685 1.680 8780/0.0508 1.004 R1 = 0.0500 wR2 = 0.1199 R1 = 0.0994 wR2 = 0.1664

Rindices (all data)

Table 2

Select bond lengths (Å) and angles (°) for complexes 3, 4 and 5

Ru(1)–N(1) Ru(1)–N(2) Ru(1)–N(3) Ru(1)–N(6) Ru(1)–N(7) Ru(1)–N(8) N(1)–Ru(1)–N(2) N(1)–Ru(1)–N(3) N(1)–Ru(1)–N(6) N(1)–Ru(1)–N(7) N(1)–Ru(1)–N(8) N(2)–Ru(1)–N(3) N(2)–Ru(1)–N(6) N(2)–Ru(1)–N(7) N(2)–Ru(1)–N(8) N(3)–Ru(1)–N(6) N(3)–Ru(1)–N(7) N(3)–Ru(1)–N(8) N(6)–Ru(1)–N(7) N(6)–Ru(1)–N(8) N(7)–Ru(1)–N(8)

3

4

5

2.108(5) 1.973(5) 2.064(4) 2.085(5) 2.001(5) 2.064(5) 79.39(19) 157.17(19) 94.84(18) 98.56(18) 87.43(19) 77.85(18) 103.65(18) 177.52(18) 99.83(19) 92.24(17) 104.15(17) 94.67(18) 77.84(19) 156.4(2) 78.63(19)

2.097(2) 1.967(2) 2.052(2) 2.078(2) 1.999(2) 2.057(2) 79.92(9) 157.39(9) 92.70(8) 95.77(9) 92.30(8) 77.48(9) 104.99(9) 174.57(9) 98.25(9) 93.50(8) 106.77(9) 90.53(9) 78.39(9) 156.74(9) 78.52(9)

2.086(4) 1.966(4) 2.049(4) 2.083(4) 1.982(4) 2.058(4) 80.34(17) 157.72(17) 91.55(17) 93.72(17) 91.57(16) 77.38(17) 106.28(17) 172.37(17) 96.93(17) 94.92(17) 108.45(17) 90.82(17) 78.51(17) 156.78(18) 78.33(17)

All three molecules of complexes 3–5 contain a sixcoordinated ruthenium atom chelated by two tridentate ligands. Complex 3 consists of a [Ru(dtzp)( ptn)]2+ cation coordination sphere and two disordered ClO4− counterions. The naphthyl ring of the main ptn ligand is nearly coplanar with the 1,2,4triazine ring, and they can form a larger π framework. Although the ptn ligand can coordinate to Ru(II) via two different sites, i.e. N(3) of the 1,2,4-triazine ring and the nitrogen atoms of the phen ring or N(5) of the 1,2,4-triazine ring and the nitrogen atoms of the phen ring, the structure confirms that the coordination is at N(3) rather than N(5) due to the steric hindrance between the ligands. This phenomenon is consistent with our previous studies.26,27 The dihedral angle between the main ptn ligand and the ancillary dtzp ligand is

This journal is © The Royal Society of Chemistry 2014

87.89, which is nearly orthogonal. Due to the constrained bite of the ligands, the Ru–N bond lengths to the central ring [1.973(3)–2.001(3) Å] are shorter than those to the terminal rings [2.064(3)–2.108(3) Å], which is typical for coordination of conjugated terminal systems.28 This phenomenon is similar to that of [Ru(tpy)(tmen)OH2]2+ (Ru–N bond lengths: 2.163(7), 2.136(7), 2.152(6) Å).29 The average Ru–N bond length of complex 3 is 2.049 Å, which is in agreement with previously reported values of ([Ru(tpy)(pta)]2+ (2.046 Å) and [Ru(tpy)(ptp)]2+ (2.056 Å).26 The asymmetric units of complexes 4 and 5 are very similar to the asymmetric unit of complex 3, except for the main ligands. Complexes 4 and 5 contain a [Ru(tpy)(L)]2+ (L = pta, ptp) cation coordination sphere, two disordered ClO4− counterions and two acetonitrile solvent molecules in the lattice. The average Ru–N bond lengths of complexes 4 and 5 are 2.040 and 2.037 Å, respectively. As observed for complex 3, the dihedral angles between the main ligand and the ancillary dtzp ligand of complexes 4 and 5 are 89.06 and 89.33. In the three Ru(II) complexes, the main ligands ptn, pta and ptp have a large planar aromatic area and possess intercalating potential for the base pairs of double helical DNA. DNA binding studies To clarify the nature of the interaction between the complexes and CT-DNA, viscosity measurements were performed. These measurements provide sensitive detection of the binding mode of the present complexes. Hydrodynamic measurements that are sensitive to changes in length (i.e. viscosity and sedimentation) are regarded as the least ambiguous and the most critical tests of a binding model in solution in the absence of crystallographic or NMR structural data.30,31 In general, the viscosity of double-stranded DNA increases when a complex binds DNA in an intercalating mode but remains unchanged when a complex binds DNA in an electrostatic mode. If the groove binding mode occurs, there is a little effect on the

Dalton Trans., 2014, 43, 17303–17316 | 17305

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Paper

Fig. 2 Effect of increasing amounts of EB (◀) and complexes 1 (■), 2 (●), 3 (▲), 4 (▼), and 5 (◆) on the relative viscosity of CT-DNA at 30 (±0.1) °C. The total concentration of DNA is 0.5 mM.

viscosity of DNA.31 The effects of Ru(II) complexes 1–5 and ethidium bromide (EB) on the viscosity of rod-like DNA are shown in Fig. 2. As expected, EB, which is a well-known DNA intercalator, strongly increases the relative viscosity by lengthening the DNA double helix through intercalation. The viscosities of the DNA bound to the Ru(II) complexes increase to a greater extent than the viscosity of DNA bound to EB, except for complex 1. The increased degree of viscosity, which may depend on the affinity for DNA, follows the order 5 > 4 > 3 > 2 > EB > 1. Complex 1 essentially exerts no effect on the DNA viscosity at low binding ratios; however, upon additional binding of the complex to DNA, the DNA viscosity decreases. These results suggest that the complexes bind DNA in two different modes: complex 1 in a partial intercalation mode and complexes 2–5 in a classical intercalation mode. The differences in binding strength and mode of complexes 1–5 are due to the main ligands. Complexes 2–5, which contain planar ligands, can deeply intercalate into DNA base pairs and increase the DNA viscosity. In contrast, complex 1 cannot completely intercalate into DNA because the dppt ligand is somewhat sterically hindered from planarity. The partial intercalation may act as a ‘wedge’ to pry one side of a base-pair stack apart, as observed for Δ-[Ru( phen)3].31 Absorption spectra titrations were performed to determine the DNA-binding affinity of complexes 1–5. Binding of a complex to DNA through intercalation typically results in hypochromism and a red shift due to an intercalative mode involving a strong stacking interaction between the aromatic chromophore and the DNA base pairs.32 The absorption spectra of the complexes with an increasing concentration of CT-DNA are shown in Fig. S1 (ESI†). DNA-binding constants and hypochromism of the Ru(II) complexes 1–5 are summarized in Table S1 (ESI†). For complexes 2–5, clear hypochromicities and red shifts were observed with increasing concentrations of DNA. Complexes 3, 4 and 5 exhibit hypochromisms of approximately 24.43, 25.6, and 28.57% and bathochromic shifts of 5, 6 and 6 nm, respectively. To further

17306 | Dalton Trans., 2014, 43, 17303–17316

Dalton Transactions

elucidate the binding strength of the complexes, the intrinsic constants (Kb) for CT-DNA were determined by monitoring the changes of the absorbance in the MLCT band with increasing concentrations of CT-DNA. The Kb values of complexes 3, 4 and 5 are (1.03 ± 0.49) × 106 M−1 (s = 2.21), (4.3 ± 1.33) × 106 M−1 (s = 2.04), and (5.7 ± 2.71) × 106 M−1 (s = 1.75), respectively, which are greater than those of complexes 1 and 2. These values are comparable to those observed for [Ru(bpy)2(dppz)]2+ (>106 M−1)33 and [Ru( phen)2dppz]2+ (5.1 × 106 M−1)34 and higher than those observed for [Ru(tpy)( pta)]2+ (9.5 × 104 M−1),35 [Ru(tpy)( ptp)]2+ (1.6 × 105 M−1)35 and [Ru(tpy)( pti)]2+ (3.0 × 104 M−1).27 These results suggest that the DNA-binding affinities of the complexes for CT-DNA follow the order 1 < 2 < 3 < 4 < 5. The differences in DNA-binding affinities for complexes 1–5 can be attributed to the different degrees of planarity of the substituents. For intercalative ligands with an extended aromatic plane, good conjugation effects from substituted groups can greatly enhance the DNA-binding ability of the complexes. Complex 1 exhibits the lowest binding affinity for CT-DNA because the two phenyl rings in the dppt ligand are non-coplanar. However, the ligands of complexes 2–5 are nearly coplanar with the 1,2,4-triazine ring, and they can form a larger π framework compared with that of dppt. Due to the greater planar area and higher hydrophobicity, complexes 2–5 may intercalate into DNA base pairs with high affinity. Despite all this, for complex 2, which contain a slightly smaller planar ligand pti compared with ptn, pta and ptp, has weaker DNA affinity than 3, 4, 5. Topoisomerase inhibition assay Agarose gel electrophoresis assays were performed to assess the Topo I and IIα inhibitory activities of the five Ru(II) complexes. The results of the Topo I inhibition assay using different concentrations of complexes 1–5 are shown in Fig. 3. As expected, complexes 4 and 5 exhibited significant dual inhibitory activity of Topo I and IIα. Significant inhibition was observed for complexes 4 and 5 at lower concentrations in the

Fig. 3 Effects of different concentrations of complexes 1–5 on the activity of DNA topoisomerase I (Topo I). Form I: supercoiled circular; Form II: nicked circular.

This journal is © The Royal Society of Chemistry 2014

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Dalton Transactions

TopoI-mediated DNA relaxation experiments. The results of the Topo I inhibition assay indicate that complex 5 exhibits the strongest inhibitory activity (IC50 = 0.7 mM), followed by complex 4 (IC50 = 10 μM). In contrast, complexes 1 and 2 exhibit weaker Topo I inhibitory activity (IC50 > 100 μM), and complex 3 exhibits moderate Topo I inhibitory activity. The results indicate that the Topo I inhibitory activities of the complexes correspond to the DNA-binding affinities of the complexes. These results suggest that both complexes may block the DNA strand passage event of the enzyme and may serve as catalytic inhibitors of Topo I. However, as DNA intercalators, the complexes can directly alter the topological state of the negatively supercoiled DNA substrate by inducing constrained negative and unconstrained positive superhelical twists in the plasmid DNA. Because Topo I removes only the unconstrained positive supercoils, the negatively supercoiled DNA product would be identical to the topological state of the original plasmid substrate. In this case, the complexes will also appear to inhibit enzyme catalysis. To determine whether the complexes interfere with the DNA relaxation reaction by inhibiting Topo I catalysis or by altering the apparent topological state of DNA, a DNA strand passage assay was performed.36 The effects of the complexes on enzyme-catalysed DNA strand passage were assessed by comparing the rate of relaxation of the negatively supercoiled plasmid in the absence of the drug with the rate of supercoiling of the relaxed plasmid in the presence of EB. As shown in Fig. 5a, pBR322 DNA was fully relaxed in the presence of the Ru(II) complexes, whereas the relaxed plasmid was converted to supercoiled plasmid in the presence of EB. These findings indicated that the rate of Topo I-catalysed DNA supercoiling was identical to the rate of Topo I-catalysed DNA relaxation in the presence of the Ru(II) complexes. These results suggest that complexes 3, 4 and 5 are catalytic inhibitors of Topo I. Similar results were obtained in inhibition studies of Topo IIα activity using complexes 1–5, and the results are presented in Fig. 4. Complex 5 exhibits the strongest inhibitory activity (IC50 = 0.7 mM). Complex 4 exhibits a stronger inhibition of

Paper

Fig. 5 Effects of EB (30 μM) and Ru(II) complexes 3 (30 μM), 4 (5 μM) and 5 (2 μM) on DNA strand passage assays by Topo I (a) and Topo IIα (b). Lane 0: DNA alone; lane 1: Topo I (Topo IIα) + DNA; EB group: DNA + Topo I (Topo IIα) + EB at different time points; Complex 3 group: DNA + Topo I (Topo IIα) + 3 at different time points; Complex 4 group: DNA + Topo I (Topo IIα) + 4 at different time points; Complex 5 group: DNA + Topo I (Topo IIα) + 5 at different time points;.

Topo II than for Topo I (IC50 = 3 mM). Compared with Ru(II) complexes previously reported by our group as topoisomerase inhibitors,20,24,37 complexes 4 and 5 exhibited much higher topoisomerase inhibitory activity. As described for Topo I, the DNA strand passage assay was also used to distinguish the effects of the Ru(II) complexes on the Topo IIα function from their effects on DNA topology. As shown in Fig. 5b, the supercoiling of the relaxed plasmid in the presence of the Ru(II) complexes was prevented, which suggests that complexes 3, 4 and 5 are catalytic inhibitors (or poisons) of human topoisomerase IIα. MTT assay The antiproliferative activities of complexes 1–5 were evaluated against three human tumour cell lines (HeLa, BEL-7402, and Hep G2) and one normal cell line LO2 using an MTT assay. Cisplatin and 5-fluorouracil were used as positive controls. The IC50 values obtained are summarised in Table 3. The complexes exhibit higher cytotoxicity in vitro against the select tumour cell lines than 5-fluorouracil, which is a widely used clinical antitumor drug, but has a relatively lower cytotoxicity than cisplatin. The five complexes exhibit small differences in the antitumor activity against different evaluated cell lines. As shown in Table 3, the five complexes exhibit higher activity against HeLa cells than against BEL-7402 and Hep G2 cells. A comparison of the IC50 values of these complexes against HeLa cells indicates that complexes 4 and 5 exhibit higher activities than the other complexes under identical experimental conditions. Most importantly, the complexes exhibited much less cytotoxicity in normal cells than three cancer cell lines, whereas cisplatin exhibited higher cytotoxicity in normal cells than in the cancer cells. This suggested that the complexes had high selectivity between tumor cells and normal cells. Cell uptake

Fig. 4 Effects of different concentrations of complexes 1–5 on the activity of DNA topoisomerase IIα (Topo II).

This journal is © The Royal Society of Chemistry 2014

To gain insight into a possible relationship between intracellular levels and in vitro cytotoxicity of the Ru(II) complexes, the uptake of complexes 1–5 was investigated by ICP-MS in HeLa cells. The levels of Ru in different cellular localisations were determined after 24 h treatment of HeLa cells with the

Dalton Trans., 2014, 43, 17303–17316 | 17307

View Article Online

Paper

Dalton Transactions

Table 3 IC50 values for complexes 1–5, cisplatin and 5-fluorouracil against different cell lines

IC50 (μM) Complex

HeLa

HepG2

BEL-7402

LO2

1 2 3 4 5 Cisplatin 5-Fluorouracil

93.2 ± 2.6 84.3 ± 2.8 75.6 ± 3.7 58.8 ± 2.1 42.1 ± 1.7 20.6 ± 1.5 1.2 × 103

181.5 ± 4.1 180.3 ± 5.7 169.2 ± 4.4 149.5 ± 7.2 102.3 ± 5.4 23.4 ± 1.2 2.45 × 103

263.1 ± 8.0 231.2 ± 7.2 220.9 ± 4.7 148.3 ± 5.9 185.3 ± 6.6 35.1 ± 2.8 3.6 × 103

387.3 ± 7.4 362.8 ± 6.2 402.6 ± 9.2 357.1 ± 7.5 346.2 ± 6.8 21.3 ± 1.2 3.3 × 103

and trigger mitochondrial membrane permeabilisation. Complexes 1–5 accumulated preferentially in the nucleus and mitochondria of HeLa cells, and this result was consistent with that of previous studies.37,38

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Cell cycle analysis

Fig. 6 Cellular ruthenium concentrations determined in the nucleus (Nuc), mitochondria (Mit) and cytoplasm (Cyt) in HeLa cells after 24 h of incubation with 50 μM complexes 1–5 by ICP-MS.

complexes. The results are summarised in Fig. 6 and Table S2 (ESI†). The uptake levels of HeLa cells are relatively high for the complexes, except for complex 1, and the results revealed that a majority of the Ru(II) complexes accumulated in the nucleus. The level of Ru in the nucleus (6.233 pg of Ru per cell) is 3.7 fold greater than that in the cytoplasm (1.673 pg of Ru per cell), which indicates that more than 60% of the ruthenium accumulated in the nucleus for complex 5. This result indicated that complex 5, which exhibited the highest antiproliferative activity, was readily taken up by the cells, particularly in the nucleus. Complexes 2, 3 and 4 exhibited a distribution similar to that of complex 5. Complex 2, which exhibited relatively lower cytotoxicity, exhibited the highest uptake level in HeLa cells; this finding may be correlated with lower DNAbinding affinity and inhibitory activity of Topo I and IIα. In addition, the ruthenium concentrations in mitochondria were also determined. Notably, the majority of the Ru(II) complexes in the cytoplasm accumulated in the mitochondrial fraction. This result indicated that apoptotic pathways induced by Ru(II) complexes can converge at the mitochondrial level

17308 | Dalton Trans., 2014, 43, 17303–17316

To investigate the cellular effects that are induced by complexes 1–5 in terms of cell cycle modification and the induction of apoptosis, a time-dependent evaluation of the cell cycle profile was performed by fluorescence-activated cell sorting (FACS) analysis of the DNA content in HeLa cells. After treatment with the complexes for 24 h, 36 h and 48 h, the cell cycle distribution of HeLa cells was quantitatively determined by flow cytometric analysis of DNA histograms (Fig. S2, ESI†). The results indicated that complexes 1–5 caused a decrease in cells in the S phase (31.59%–22.59%) and the corresponding increase in cells in the G2/M (10.7%–23.16%) phase. For cells treated with the complexes, no significant change in the G0/G1 phase was observed. The increase of varying degrees in the sub-G1 phase cell population was also observed after 36 h or 48 h of treatment, and the increase in the percentage of apoptotic cells was dose dependent. Interestingly, during the treatment of HeLa cells with complex 5, a large increase in the percentage of cells in the S phase (up to 48.33%) and the corresponding reduction in the G2/M phase were observed. In addition, significant increases in DNA debris (grey area) were observed, which indicated that the complex could induce DNA damage within a time that corresponded to the results of a subsequent comet assay. These results indicate that the complexes primarily induced apoptosis because the accumulation of hypodiploid DNA (sub-G1 phase cells) is considered as an indicator of apoptotic cell death.39 Apoptosis assay Cell death can be divided into two types: necrosis (accidental cell death) and apoptosis ( programmed cell death).40 Necrotic cells undergo cell lysis and lose their membrane integrity, and severe inflammation is induced.41 In contrast, apoptotic cells are transformed into small membrane-bound vesicles (apoptotic bodies), which are engulfed in vivo by macrophages, and no inflammatory response is observed.42 The harmless removal of cells (for example, cancer cells) is one consideration in chemotherapy.43 Therefore, the induction of apoptosis is a consideration in the development of anticancer drugs. Morphological analysis of cell death induced by complexes 1–5 was performed using a staining method that utilises acridine orange (AO) and ethidium bromide (EB). This method combines the differential uptake of the fluorescent DNAbinding dyes AO and EB.44 AO is a vital dye and can stain both live and dead cells. EB stains only cells that have lost their membrane integrity. Under a fluorescence microscope, live cells appear green. Necrotic cells stain red but exhibit a nuclear morphology resembling that of viable cells. Apoptotic cells appear green and exhibit morphological changes such as cell blebbing and the formation of apoptotic bodies. Upon treatment of the HeLa cells with the Ru(II) complexes,

This journal is © The Royal Society of Chemistry 2014

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Dalton Transactions

Fig. 7 HeLa cells stained using AO/EB and observed by fluorescence microscopy: (a) HeLa cells without treatment; (b)–(f) HeLa cells incubated in the presence of complexes 1–5 (75 μM), respectively, for 48 h. Arrows indicate the following: L, live cells; A, apoptotic cells; and N, necrotic cells.

apoptotic features such as nuclear shrinkage and chromatin condensation, as well as red necrotic cells, were observed (Fig. 7). Four types of cells could be distinguished according to the fluorescence emission and the morphological aspect of chromatin condensation in the stained nucleus: viable cells, which exhibit uniform bright green nuclei with an organised structure; early apoptotic cells; late apoptotic cells, which exhibit orange to red nuclei with condensed chromatin; and red necrotic cells. These results suggest that HeLa cells treated with complexes 1–5 responded with a higher incidence of apoptosis than necrosis. Under identical conditions, the degree of apoptosis in HeLa cells caused by complex 1 is weaker than that of complexes 2–5, which is in agreement with the DNA binding affinity of the complex.

Paper

The morphological change of the treated cells is a qualitative feature of the apoptosis-inducing activity of Ru(II) complexes. Quantitative analysis of the apoptosis-inducing activity was performed by Alexa Fluor®488-conjugated annexin V/PI staining. Externalisation of phosphatidylserine (PS) on the outer surface of the plasma membrane is a known hallmark of apoptosis or cell necrosis.45 The difference between the two forms of cell death is that during the early stages of apoptosis, the cell membrane remains intact, whereas at the moment when necrosis occurs, the cell membrane loses its integrity. The phospholipid-binding protein, Alexa Fluor®488-conjugated annexinV, binds to cells with exposed PS, whereas PI only stains cells that have lost membrane integrity.46 Doublestaining with annexin V and PI can distinguish four different cell types during apoptosis: live cells (annexin V−/PI+), early apoptotic cells (annexin V+/PI−), late apoptotic cells (annexin V+/PI+) and necrotic cells (annexin V−/PI+). Complexes 4 and 5 were selected for quantitative analysis of apoptosis-inducing activity, and the results are shown in Fig. 8. Treatment with these complexes resulted in a significant dose-dependent enhancement in the total number of apoptotic cells. Following treatment with complex 4 at 50 μM, 46.9% of early and 6.4% of late apoptotic cells were observed. However, at 100 μM, the ratio of early and late apoptotic cells was up to 75.5%. As shown in Fig. 8, complex 5 was more effective than complex 3 in inducing PS exposure and the loss of plasma membrane integrity. Following treatment with complex 5 at 100 μM, up to 31.8% of late apoptotic cells was observed, whereas only 10.6% of the cells exhibited markers of late apoptosis following treatment with complex 4.

Fig. 8 HeLa cells treated with different concentrations of Ru(II) complexes were double-stained with annexinV/PI and analysed by flow cytometry. (a) Control; (b) complex 4 (50 μM); (c) complex 4 (100 μM); (d) complex 5 (50 μM); and (e) complex 5 (100 μM).

This journal is © The Royal Society of Chemistry 2014

Dalton Trans., 2014, 43, 17303–17316 | 17309

View Article Online

Paper

Comet assay

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

To determine whether the complexes induce DNA damage, which is a hallmark of early apoptosis, single cell gel electrophoresis was conducted. DNA strand breaks can be detected with high sensitivity at the single-cell level using the comet assay.47 As shown in Fig. 9, HeLa cells treated with the com-

Dalton Transactions

plexes exhibit significant well-formed comets, whereas the control (untreated) cells exhibit a round shape. This finding clearly indicates that the complexes indeed induce DNA fragmentation, and the length of the comet tail represents the extent of DNA damage. The results of the comet assay indicate that DNA degradation occurred as a consequence of direct DNA damage or rapid apoptosis in HeLa cells. Mitochondrial membrane potential (ΔΨm) analysis

Fig. 9 Drug-induced double-strand DNA breaks in HeLa cells. Cells were (a) untreated or treated with (b) complex 1 (75 μM), (c) complex 2 (75 μM), (d) complex 3 (75 μM), (e) complex 4 (75 μM) and (f ) complex 5 (75 μM) for 48 h.

Mitochondrial dysfunction has been shown to participate in the induction of apoptosis through the release of proapoptotic factors such as cytochrome c and apoptosis-inducing factor and has even been suggested to be central to the apoptotic pathway. The dissipation of the mitochondrial electrochemical potential gradient is known as an early event in apoptosis.48–50 Thus, changes in the mitochondrial membrane potential (MMP, ΔΨm) are monitored using an inverted fluorescence microscope and by flow cytometry after staining live cells with the cationic dye JC-1 (5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolocarbocyanine iodide). JC-1, which preferentially enters into mitochondria, exhibits potential-dependent

Fig. 10 (A) Fluorescence imaging of JC-1 labelled cells was observed using an inverted fluorescence microscope. Cells were untreated or treated with complexes 4 and 5 (50 μM). (B) Effects of complex 4 on MMP were analysed by flow cytometry. Cells were untreated or treated with different concentrations of complex 4 for 24 h.

17310 | Dalton Trans., 2014, 43, 17303–17316

This journal is © The Royal Society of Chemistry 2014

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Dalton Transactions

accumulation in mitochondria. Following the loss of the membrane potential, the dye is dispersed throughout the entire cell leading to a shift from red (J-aggregates) to green fluorescence (JC-1 monomers). Representative images obtained using an inverted fluorescence microscope are shown in Fig. 10A. Following JC-1 accumulation, intact cells exhibited a bright red fluorescence indicating high potential. In contrast, HeLa cells treated with complexes 4 and 5 exhibited a weak yellow or slight green fluorescence, indicating the loss of the MMP. The quantitative analysis of the loss of the MMP was monitored by flow cytometry. As shown in Fig. 10B, complex 4 significantly induced a dose-dependent decrease in the MMP in HeLa cells. Complex 4 altered the MMP in cancer cells, with a rapid increase in depolarisation after 24 h of treatment, as the percentage increased from 15.4% (control) to 66.7%. The results of qualitative and quantitative analyses clearly indicate that the complexes induce apoptosis in cancer cells via a mitochondrial pathway.

Conclusions In summary, five novel ruthenium(II) complexes, [Ru(dtzp)(dppt)]2+ (1), [Ru(dtzp)( pti)]2+ (2), [Ru(dtzp)( ptn)]2+ (3), [Ru(dtzp)( pta)]2+ (4) and [Ru(dtzp)( ptp)]2+ (5), were synthesised and characterised. DNA-binding studies indicated that complexes 2–5 interact with CT-DNA via an intercalative mode, whereas complex 1 binds DNA via a partial intercalative mode. All the results indicated that the ligand planarity of a complex significantly affects the DNA binding affinity of the complex. In addition, a topoisomerase inhibition assay suggested that complexes 3, 4 and 5 exhibit significant dual inhibition of Topo I and Topo IIα. The complexes 3, 4 and 5 exhibited excellent antitumor activity against HeLa cells and much less cytotoxicity in the normal cell line than cancer cell lines. The ICP-MS study indicated that complexes 1–5 preferentially accumulated in the nucleus and mitochondria of HeLa cells and subsequently induced apoptosis. According to the results from fluorescence microscopy and flow cytometric analysis, the complexes primarily induced apoptosis in cancer cells. DNA damage was observed by single cell gel electrophoresis. Mitochondrial membrane potential (ΔΨm) analysis suggests that the complexes induce apoptosis via the mitochondrial pathway. These results may be valuable in understanding the DNA binding and topoisomerase inhibition by Ru(II) complexes and lay the foundation for the discovery of new antitumour agents.

Experimental section General procedures All reagents and solvents were of analytical grade except those employed in photophysical experiments, which were of spectroscopic grade. Doubly distilled water was used to prepare all buffers. Calf thymus DNA (CT-DNA) was obtained from the

This journal is © The Royal Society of Chemistry 2014

Paper

Shanghai Sangon Biological Engineering Technology & Services Co., Ltd. A solution of calf thymus DNA in buffer gave a ratio of UV absorbance at 260 nm to that at 280 nm of approximately 1.8–1.9, indicating that the DNA was sufficiently free of protein.51 The DNA concentration per nucleotide was determined by absorption spectroscopy using the molar absorption coefficient (6600 M−1 cm−1) at 260 nm.52 MTT was purchased from Sigma and used without further purification. A mitochondria staining kit (JC-1) was obtained from Multi Sciences Biotech Co., Ltd. An annexin V/PI apoptosis detection kit was purchased from Invitrogen (UK). Other materials were commercially available and used without further purification. Microanalysis (C, H, and N) was carried out using a PerkinElmer 240Q elemental analyzer. 1HNMR spectra were recorded on a Varian Mercury plus 300 NMR spectrometer and a Varian INOVA 500NB spectrometer with (CD3)2SO as the solvent at room temperature and TMS as the internal standard. Fast atom bombardment (FAB) mass spectra were acquired on a VG ZAB-HS spectrometer in a 3-nitrobenzyl alcohol matrix. Electrospray mass spectra (ES-MS) were recorded on an LCQ system (Finnigan MAT, USA). The spray voltage, tube lens offset, capillary voltage and capillary temperature were set at 4.50 kV, 30.00 V, 23.00 V and 200 °C, respectively, and the reported m/z values are for the major peaks in the isotope distribution. UV-Vis spectra were recorded on a PerkinElmer LAMBDA 850 spectrophotometer. Emission spectra were recorded on a PerkinElmer LS55 spectrofluorophotometer at room temperature. Synthesis The ligands 2-cyno-1,10-phenanthroline,53 2-amino(hydrazono)methyl-1,10-phenanthroline (PNH),26 2,6-di(thiazol-2-yl)pyridine (dtzp),54 3-(1,10-phenanthroline-2-yl)-5,6-diphenyl-astriazine (dppt),26 3-(1,10-phenanthroline-2-yl)-as-triazino-[5,6-f]isatin ( pti),55 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]naphthalene ( ptn),27 3-(1,10-phenanthroline-2-yl)-as-triazino[5,6-f ]acenaphthylene) ( pta),26 3-(1,10-phenanthroline-2-yl)as-triazino[5,6-f ]-phenanthrene ( ptp)26 and Ru(dtzp)Cl3 56 were synthesised as previously described. [Ru(dtzp)(dppt)](ClO4)2 (1) A mixture of Ru(dtzp)Cl3 (0.1 g, 0.221 mmol), dppt (0.095 g, 0.221 mmol) and triethylamine (1 cm3) in 50 cm3 ethanol– water (1 : 1, v/v) under argon was refluxed for 12 h to yield a dark purple solution. Upon cooling to room temperature, a dark purple precipitate was obtained by the addition of an aqueous NaClO4 solution. The product was purified by column chromatography on alumina with acetonitrile–toluene (2 : 1, v/v) as the eluent. The major band was collected. The solvent was removed under reduced pressure, and a dark purple powder was obtained. Yield: 0.126 g, 59.4%. Anal. Calcd (%) for C38H24Cl2N8O8S2Ru: C, 47.70; H, 2.53; N, 11.71. Found: C, 47.58; H, 2.71; N, 11.60. 1H NMR ( ppm, DMSO-d6, 300 MHz, 25 °C): δ 9.18 (d, 1H, J = 8.6 Hz), 9.09 (d, 1H, J = 8.6 Hz), 8.88 (d, 2H, J = 8.0 Hz), 8.71 (d, 1H, J = 7.5 Hz), 8.64 (d, 1H, J = 9.0 Hz), 8.48 (t, 1H, J = 8.7 Hz), 8.20 (d, 1H, J = 5.1 Hz), 7.94 (d, 2H,

Dalton Trans., 2014, 43, 17303–17316 | 17311

View Article Online

Paper

J = 3.4 Hz), 7.67 (d, 1H, J = 5.4 Hz), 7.64 (d, 1H, J = 2.7 Hz), 7.56 (d, 2H, J = 7.0 Hz), 7.48 (d, 1H, J = 7.4 Hz), 7.39 (t, 1H, J = 7.8 Hz), 7.26 (t, 2H, J = 7.8 Hz),7.20 (m, 3H), 7.11 (d, 2H, J = 3.3 Hz), 7.04 (d, 1H, J = 7.2 Hz). ES-MS [CH3CN, m/z]: 379.03 ([M − 2ClO4]2+), 857.01 ([M − ClO4]+).

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

[Ru(dtzp)( pti)](ClO4)2 (2) This complex (dark brown) was synthesised as described for complex 1, with pti (0.077 g, 0.221 mmol) instead of dppt. Yield: 0.132 g, 66.7%. Single crystals suitable for an X-ray crystallographic study were grown from acetonitrile–toluene (1 : 1, v/v) at room temperature. Anal. Calcd (%) for C32H19Cl2N9O8S2Ru: C, 43.01; H, 2.14; N, 14.11. Found: C, 42.86; H, 2.46; N, 13.95. 1H NMR ( ppm, DMSO-d6, 300 MHz, 25 °C): δ 9.20 (d, 1H, J = 8.6 Hz), 8.99 (d, 1H, J = 8.6 Hz), 8.86 (d, 2H, J = 8.0 Hz), 8.63–8.53 (m, 2H), 8.48 (t, 1H, J = 8.0 Hz), 8.34 (d, 1H, J = 8.9 Hz), 8.04 (d, 1H, J = 7.5 Hz), 7.93 (dd, 1H, J = 5.1, 1.0 Hz), 7.81 (d, 2H, J = 3.4 Hz), 7.57 (dd, 1H, J = 8.2, 5.2 Hz), 7.38 (t, 1H, J = 7.5 Hz), 7.05 (t, 2H, J = 7.4 Hz), 7.00–6.97 (m, 2H). ES-MS [CH3CN, m/z]: 347.50 ([M − 2ClO4]2+), 794.02 ([M − ClO4]+). [Ru(dtzp)( ptn)](ClO4)2 (3) This complex (dark brown) was synthesised as described for complex 1, with ptn (0.080 g, 0.221 mmol) instead of dppt. Yield: 0.116 g, 58.1%. Single crystals suitable for X-ray crystallographic study were grown from acetonitrile–toluene (1 : 1, v/v) at room temperature. Anal. Calcd (%) for C34H20Cl2N8O8S2Ru: C, 45.14; H, 2.23; N, 12.39. Found: C, 45.21; H, 2.58; N, 12.18. 1H NMR ( ppm, DMSO-d6, 300 MHz, 25 °C): δ 9.49 (d, 1H, J = 4.5 Hz), 9.46 (t, 1H, J = 5.1 Hz), 9.17 (d, 1H, J = 8.7 Hz), 8.94 (d, 2H, J = 8.1 Hz), 8.72 (d, 1H, J = 8.4 Hz), 8.65 (d, 1H, J = 9.0 Hz), 8.55 (t, 1H, J = 7.8 Hz), 8.47 (d, 1H, J = 9.0 Hz), 8.26 (d, 1H, J = 9.0 Hz), 8.18 (d, 1H, J = 5.4 Hz), 8.13 (d, 1H, J = 5.4 Hz), 8.04–8.01 (m, 2H), 7.89 (d, 2H, J = 3.3 Hz), 7.71–7.65 (m, 2H), 7.09 (d, 2H, J = 3.3 Hz). ES-MS [CH3CN, m/z]: 353.02 ([M − 2ClO4]2+), 805.99 ([M − ClO4]+).

Dalton Transactions

[Ru(dtzp)( ptp)](ClO4)2 (5) This complex (dark brown) was synthesised as described for complex 1, with ptp (0.090 g, 0.221 mmol) instead of dppt. Yield: 0.162 g, 76.74%. Single crystals suitable for X-ray crystallographic study were grown from acetonitrile–toluene (1 : 1, v/v) at room temperature. Anal. Calcd (%) for C42H28Cl2N10O8S2Ru: C, 48.65; H, 2.72; N, 13.51. Found: C, 48.72; H, 2.63; N, 13.57. 1H NMR ( ppm, DMSO-d6, 500 MHz, 25 °C): δ 9.48 (d, 1H, J = 8.5 Hz), 9.45 (d, 1H, J = 7.0 Hz), 9.20 (d, 1H, J = 8.5 Hz), 9.02 (d, 2H, J = 8.0 Hz), 8.83 (dd, 1H, J = 8.5, 8.5 Hz), 8.77 (d, 1H, J = 8.5 Hz), 8.68 (d, 1H, J = 9.0 Hz), 8.65 (d, 1H, J = 8.0 Hz), 8.51 (d, 1H, J = 9.0 Hz), 8.33 (d, 1H, J = 5.5 Hz), 8.22 (d, 1H, J = 7.0 Hz), 8.05 (t, 1H, J = 7.0 Hz), 7.97–7.93 (m, 2H), 7.91 (d, 2H, J = 3.5 Hz), 7.75–7.70 (m, 3H), 7.12 (d, 2H, J = 3.5 Hz). ES-MS [CH3CN, m/z]: 378.03 ([M − 2ClO4]2+), 856.0 ([M − ClO4]+). Caution: Perchlorate salts of metal complexes with organic ligands are potentially explosive, and only small amounts of the material should be prepared and handled with great care. X-ray crystallography Diffraction data for compound 3 were recorded on an Oxford Diffraction Gemini R CCD diffractometer with Cu-Kα radiation at 150 K. Diffraction data for complexes 4 and 5 were collected on a Bruker SMART Platform CCD diffractometer with Mo-Kα radiation at 110 K and 293 K, respectively. The absorption correction was applied using the SADABS program.57 The temperature was controlled using Oxford Cryostream cooling apparatus. Structure determination and full-matrix leastsquares refinement based on F2 for complexes 3–5 were performed using the SHELXS 97 and SHELXL 97 program packages, respectively.58 All non-hydrogen atoms were anisotropically refined. Hydrogen atoms of the organic ligands were geometrically generated (C–H 0.96 Å). Detailed crystallographic data for the crystal structural analysis have been deposited in the Cambridge Crystallographic Data Centre; 879756 (3), 879757 (4), and 879758 (5) contain the supplementary crystallographic data for the present paper.

[Ru(dtzp)( pta)](ClO4)2 (4) This complex (dark brown) was synthesised as described for complex 1, with pta (0.084 g, 0.221 mmol) instead of dppt. Yield: 0.152 g, 74.14%. Single crystals suitable for X-ray crystallographic study were grown from acetonitrile–toluene (1 : 1, v/v) at room temperature. Anal. Calcd (%) for C40H26Cl2N10O8S2Ru: C, 47.53; H, 2.59; N, 13.86. Found: C, 47.62; H, 2.47; N, 13.91. 1H NMR ( ppm, DMSO-d6, 500 MHz, 25 °C): δ 9.26 (d, 1H, J = 8.5 Hz), 9.14 (d, 1H, J = 8.5 Hz), 8.99 (d, 2H, J = 8 Hz), 8.72 (t, 1H, J = 7.0 Hz), 8.69 (d, 1H, J = 9.0 Hz), 8.61 (t, 1H, J = 8.0 Hz), 8.51 (d, 1H, J = 8.5 Hz), 8.48 (d, 1H, J = 9.0 Hz), 8.44 (d, 1H, J = 8.0 Hz), 8.17 (d, 1H, J = 5.0 Hz), 8.11–8.07 (t, 2H), 7.94 (d, 2H, J = 3.0 Hz), 7.91 (d, 1H, J = 7.0 Hz), 7.68 (d, 1H, J = 5.5 Hz), 7.66 (d, 1H, J = 5.0 Hz), 7.10 (d, 2H, J = 3.5 Hz). ES-MS [CH3CN, m/z]: 365.02 ([M − 2ClO4]2+), 829.98 ([M − ClO4]+).

17312 | Dalton Trans., 2014, 43, 17303–17316

DNA binding experiments DNA binding experiments were performed at room temperature. All spectroscopic titrations and viscosity measurements were carried out in buffer A (5 mM Tris-HCl, 50 mM NaCl, pH 7.0; Tris = Tris(hydroxymethyl)aminomethane). Thermal DNA denaturation was carried out in buffer B (1.5 mM Na2HPO4, 0.5 mM NaH2PO4, 0.25 mM Na2EDTA; EDTA = ethylene diaminetetraacetic acid, pH = 7.0). Viscosity measurements were carried out using an ubbelohde viscometer maintained at 29.0 ± 0.1 °C in a thermostatic bath. The flow time was measured using a digital stopwatch; each sample was measured three times, and an average flow time was calculated. Data are presented as (η/η0)1/3 versus the binding ratio,59 where η is the viscosity of DNA in the presence of the complex and η0 is the viscosity of CT-DNA alone.

This journal is © The Royal Society of Chemistry 2014

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Dalton Transactions

Paper

The absorption titration experiments of the Ru(II) complexes were performed at the fixed complex concentration (10 μM) until the absorbance did not change with increasing DNA. The complex–DNA solutions were allowed to incubate for 5 min before the absorption spectra were recorded. The intrinsic binding constants Kb to DNA were determined using eqn (1), as follows:60 pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi b  ðb2  2Kb 2 Ct ½DNA=sÞ εa  εf ¼ ð1aÞ 2KCt εb  εf b ¼ 1 þ K b Ct þ K b ½DNA=2s

ð1bÞ

where [DNA] is the concentration of DNA in base pairs and the apparent absorption coefficients εa, εf, and εb correspond to Aobsd/[Ru], the extinction coefficient for the free ruthenium complex, and the extinction coefficient for the ruthenium complex in the fully bound form, respectively. Kb is the equilibrium binding constant in M−1, Ct is the total metal complex concentration, and s is the binding size.

pBR322 plasmid DNA and Topo I (5 units) in 40 μL of Topo I reaction buffer. Reactions were carried out in the absence of the drug or in the presence of 40 μM Ru(II) complexes or EB. Following a 5 min incubation of DNA with the drug or water, Topo I was added, and the reactions were incubated up to 20 min at 37 °C. Reactions were terminated, processed, and subjected to gel electrophoresis as previously described. The Topo IIα DNA strand passage activity was examined in the presence of EB and the Ru(II) complexes. DNA strand passage assays contained 0.3 μg of pBR322 plasmid DNA (relaxed or supercoiled, as previously described) and Topo IIα (10 units) in 40 μL of Topo IIα reaction buffer. Reactions were carried out in the absence of the drug or in the presence of 40 μM Ru(II) complexes or EB. Following a 5 min incubation of DNA with the drug or water, Topo IIα was added, and the reactions were incubated up to 20 min at 30 °C. Reactions were terminated, processed, and subjected to gel electrophoresis as previously described. Cell culture

Topoisomerase inhibition assay

Human cancer cell lines (including cervical carcinoma HeLa, hepatocellular carcinoma HepG2, human hepatoma BEL-7402) and a normal cell line (normal hepatic cell LO2) were obtained from the American Type Culture Collection (ATCC, Rockville, MD). All cell lines were cultured as monolayers and maintained in Dulbecco’s modified Eagle’s medium (DMEM, Gibco BRL) supplemented with 10% foetal bovine serum and 1% penicillin-streptomycin solution at 37 °C in a humidified incubator under a 5% CO2 atmosphere. All complexes were dissolved in DMSO and diluted in tissue culture medium before use.

DNA topoisomerase I (Topo I) from calf thymus was purchased from MBI Fermentas and used without further purification. One unit of the enzyme was defined as the amount that completely relaxes 1 μg of the negatively supercoiled pBR322 DNA in 30 min at 37 °C under standard assay conditions. The reaction mixture (20 μL) contained 35 mM Tris-HCl ( pH 8.0), 72 mM KCl, 5 mM MgCl2, 5 mM DTT, 2 mM spermidine, 0.1 mg ml−1 BSA, 0.1 μg of pBR322 DNA, 1 unit of Topo I, and the Ru(II) complexes. The reaction mixtures were incubated at 37 °C for 30 min, and the reaction was terminated by the addition of 4 μL of 5 × stop solution consisting of 0.25% bromophenol blue, 4.5% SDS, and 45% glycerol. The samples were electrophoresed using 1% agarose in TBE at 80 V for 2 h. The gel was stained with EB (1 μg mL−1) and then photographed and analysed on a Tanon-3500 fluorescence chemiluminescence and visible imaging system. Recombinant human DNA topoisomerase IIα (Topo IIα) was purchased from Affymetrix Inc. (USB) and used without further purification. One unit of the enzyme was defined as the amount that completely relaxes 0.3 μg of negatively supercoiled pBR322 plasmid DNA in 15 min at 30 °C under standard assay conditions. The reaction mixture (20 μL) contained 10 mM Tris-HCl ( pH 7.9), 50 mM NaCl, 50 mM KCl, 5.0 mM MgCl2, 0.1 mM Na2H2EDTA, 15 μg ml−1 BSA, 1.0 mM ATP, 0.1 μg of pBR322 DNA, 2 units of Topo IIα, and the Ru(II) complexes. The reaction mixtures were incubated at 30 °C for 15 min. Reactions were terminated, processed, and subjected to gel electrophoresis as previously described.

In vitro cytotoxicity was assessed using a standard MTT (3-(4,5dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide) colorimetric assay.63 Cells were plated in 96-well microassay culture plates (1 × 104 cells per well) and grown overnight at 37 °C in a 5% CO2 incubator. The cells were then incubated with the test compounds at different concentrations for an additional 48 h. Control wells were prepared by the addition of DMSO (1%). At the end of this incubation, 20 μL of MTT (5 mg mL−1 in PBS) was added to each well. After a 4 h incubation, the formazan crystals were dissolved in 150 μL of dimethylsulphoxide (DMSO), and the absorbance was determined at 595 nm using a microplate spectrophotometer. The IC50 value was determined from plots of % viability against the dose of the compound added. Three different human carcinomas were analysed: HeLa (cervical cancer cells), BEL-7402 (hepatic cancer cells), and HepG2 (hepatocellular carcinoma).

DNA strand passage assay

ICP-MS analysis

The DNA strand passage activity of Topo I was determined by monitoring the ability of the enzyme to relax negatively supercoiled plasmid molecules in the absence of drug61 or to supercoil relaxed plasmid substrates in the presence of intercalative agents.62 DNA strand passage assays contained 0.3 μg of

For uptake studies, exponentially growing cells were harvested, and the resulting single-cell suspension was plated in 100 mm tissue culture plates (Costar) at 1 × 105 cells per plate. After24 h at 37 °C, the medium of subconfluent cells was replaced with a fresh medium containing the desired test

This journal is © The Royal Society of Chemistry 2014

MTT assay

Dalton Trans., 2014, 43, 17303–17316 | 17313

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Paper

compounds at an appropriate concentration. After 24 h of drug treatment, the cells were washed three times with cold saline solution (0.9% NaCl). After digestion, the cells were counted and divided into three portions. In the first portion, the nuclei were extracted using a nucleus extraction kit; in the second portion, the cytoplasm was extracted using a cytoplasm extraction kit; and in the third portion, the mitochondria were extracted using a mitochondria extraction kit. The samples were digested with 60% HNO3 at RT for one day. Each sample was diluted with MilliQ H2O to obtain 2% HNO3 sample solutions.64 The ruthenium concentrations in the three portions were determined using an ICP-MS (Thermo Elemental Co., Ltd).

Dalton Transactions

stained cells were analysed by flow cytometry on a FACS Canto II system (BD Biosciences, USA). Single cell gel electrophoresis (comet assay)

HeLa cells at a density of 1 × 106 cells ml−1 were treated with Ru(II) complexes 1–5 for the indicated times. After treatment, cells were collected by trypsinisation and centrifugation, washed with PBS, and fixed with ice-cold 70% ethanol at −20 °C overnight. Fixed cells were resuspended in 0.5 mL of PBS containing 50 μL of RNase (Sigma; 1 mg mL−1 in PBS) and 50 μL of propidium iodide (PI) (Sigma, 500 mg mL−1 in PBS) for 30 min at 37 °C in the dark. The cell cycle distribution was analysed using the FACS Canto II system (BD Biosciences, USA). The data were acquired and analysed using the ModFit LT software (BD).

This single cell electrophoresis assay was performed to measure the extent of DNA fragmentation induced by the complexes.65 HeLa cells were incubated with metal complexes 1–5 at a concentration that allows for 75% viability. Following exposure, cells were harvested by centrifugation, resuspended in a culture medium at a density of 1 × 105 cells mL−1, and mixed with 1% low melting point agarose at 37 °C at a ratio of 1 : 2. The mixture (100 μL) was spread on slides precoated with 1% normal melting point agarose. After solidification of the agarose, the embedded cells were lysed in a precooled lytic solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris base, 1% Triton X-100, 10% DMSO, pH = 10) at 4 °C for 2 h. Slides were then denatured in an electrophoresis tank containing alkaline electrophoresis buffer (0.3 M NaOH and 1 mM EDTA, pH = 13) for 30 min. Next, electrophoresis was carried out at 25 V/ 300 mA for 20 min, and the slides were neutralised with 0.4 M Tris ( pH 7.5) and immersed in 70% ethanol for 30 min. Samples were allowed to air-dry and were stained with 50 μL of PI (5 μg mL−1). Samples were observed under an inverted fluorescence microscope (Zeiss Axio Observer D1). All of these procedures were performed under low-light conditions to avoid light-induced DNA damage.

Acridine orange (AO) and ethidium bromide (EB) staining

Mitochondrial membrane potential analysis

Apoptosis studies were performed with a staining method utilising acridine orange (AO) and ethidium bromide (EB). According to the difference in membrane integrity between necrosis and apoptosis, AO can pass through the cell membrane, but EB cannot. Under a fluorescence microscope, live cells appear green. Necrotic cells stain red but exhibit a nuclear morphology resembling that of viable cells. Apoptotic cells appear green and exhibit morphological changes such as cell blebbing and the formation of apoptotic bodies. A monolayer of HeLa cells was incubated in the absence or presence of the Ru(II) complexes at a concentration of 75 μM at 37 °C and 5% CO2 for 48 h. After 48 h, cells were stained with an AO/EB solution (100 μg ml−1 AO, 100 μg ml−1 EB). Samples were observed under an inverted fluorescence microscope (Zeiss Axio Observer D1).

The mitochondrial membrane potential was determined using the fluorescent probe JC-1, which produces green fluorescence in the cytoplasm and red fluorescence when accumulated in mitochondria that have a negative internal potential. Qualitative analysis of mitochondrial dysfunction was examined by fluorescence microscopy. A monolayer of HeLa cells in 12-well plates was incubated in the absence or presence of the Ru(II) complexes at 37 °C and 5% CO2. After cells were stained with 10 μM JC-1 for 5 min, the cell layer was washed three times with warm PBS. Fluorescence imaging of JC-1-labelled cells was observed using an inverted fluorescence microscope (Zeiss Axio Observer D1). Quantitative analysis of mitochondrial dysfunction was analysed by flow cytometry. Cells treated with complexes in 6-well plates were trypsinised and resuspended in 0.5 mL of PBS buffer containing 10 μM JC-1. After incubation for 10 min at 37 °C, the supernatant was immediately removed by centrifugation. After washing and resuspension in 500 μL of warm PBS, the stained cells were immediately analysed on a flow cytometer with 488 nm excitation using emission filters appropriate for the Alexa FLUOR®488 dye and R-phycoerythrin.

Cell cycle analysis

Annexin V-FITC/PI assay Apoptosis was quantified by the detection of phosphatidylserine surface exposure on apoptotic cells using an annexin V-FITC/PI ( propidium iodide) apoptosis detection kit. HeLa cells were incubated with or without the Ru(II) complexes for 48 h. The adherent and floating cells were combined and washed twice with cold PBS. Subsequently, the cells were resuspended in 100 μL of annexin V binding buffer (10 mM HEPES, 140 mM NaCl, 2.5 mM CaCl2; pH 7.4). Annexin V (5 μL) and 10 μL of propidium iodide were added to the reaction mixture and incubated for 15 min at room temperature in the dark. After the addition of 400 μL of binding buffer, the

17314 | Dalton Trans., 2014, 43, 17303–17316

Acknowledgements This work was supported by the 973 program (2014CB845604), the National Natural Science Foundation of China (no. 21172273, 21171177, 91122010), the Program for Changjiang

This journal is © The Royal Society of Chemistry 2014

View Article Online

Dalton Transactions

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Scholars and Innovative Research Team in University of China (no. IRT1298), the National High Technology Research and Development Program of China (863 Program, 2012AA020305), and the Research Fund for the Doctoral Program of Higher Education (20110171110013).

Notes and references 1 J. C. Wang, Annu. Rev. Biochem., 1985, 54, 665–697. 2 J. C. Wang, Annu. Rev. Biochem., 1996, 65, 635–692. 3 H. Turley, M. Comley, S. Houlbrook, N. Nozaki, A. Kikuchi, I. D. Hickson, K. Gatter and A. L. Harri, Br. J. Cancer, 1997, 75, 1340–1346. 4 Y. Pommier, Nat. Rev. Cancer, 2006, 6, 789–802. 5 D. Demarquay, M. Huchet, H. Coulomb, L. L. Ginot, O. Lavergne, P. G. Kasprzyk, C. Bailly, J. Camara and D. C. Bigg, Anticancer Drugs, 2001, 12, 9–19. 6 E. L. Baldwin and N. Osheroff, Curr. Med. Chem.: AntiCancer Agents, 2005, 5, 363–372. 7 Y. L. Lyu, J. E. Kerrigan, C. P. Lin, A. M. Azarova, Y. C. Tsai, Y. Ban and L. F. Liu, Cancer Res., 2007, 67, 8839–8846. 8 K. C. Murdock, R. G. Child, P. F. Fabio, R. B. Angier, R. E. Wallace, F. E. Durr and R. V. Citarella, J. Med. Chem., 1979, 22, 1024–1030. 9 C. A. Felix, Biochim. Biophys. Acta, 1998, 1400, 233–255. 10 M. Azarova, Y. Lyu, C. Lin, Y. Tsai, J. Y. Lau, J. C. Wangand and L. F. Liu, Proc. Natl. Acad. Sci. U. S. A., 2007, 107, 11014–11019. 11 S. Salerno, F. D. Settimo, S. Taliani, F. Simorini, C. La Motta, G. Fornaciari and A. M. Marini, Curr. Med. Chem., 2010, 17, 4270–4290. 12 W. A. Denny and B. C. Baguley, Curr. Med. Chem., 2003, 3, 339–353. 13 M. R. McCrystal, B. D. Evans, V. J. Harvey, P. I. Thompson, D. J. Porter and B. C. Baguley, Cancer Chemother. Pharmacol., 1999, 44, 39–44. 14 T. Utsugi, K. Aoyagi, T. Asao, S. Okazaki, Y. Aoyagi, M. Sano, K. Wierzba and Y. Yamada, Jpn. J. Cancer Res., 1997, 88, 992–1002. 15 D. Perrin, B. V. Hille, J. M. Barret, A. Kruczynski, C. Etievant, T. Imbert and B. T. Hill, Biochem. Pharmacol., 2000, 59, 807–819. 16 V. A. Rao, K. Agama, S. Holbeck and Y. Pommier, Cancer Res., 2007, 67, 9971–9979. 17 J. Stewart, P. Mistry, W. Dangerfield, D. Bootle, M. Baker, B. Kofler, S. Okiji, B. C. Baguley, W. A. Denny and P. A. Charlton, Anticancer Drugs, 2001, 12, 359–367. 18 Y. C. Lo, T. P. Ko, W. C. Su, T. L. Su and A. H. Wang, J. Inorg. Biochem., 2009, 103, 1082–1092. 19 F. Arjmand and M. Muddassir, J. Photochem. Photobiol., B, 2010, 101, 37–46. 20 K. J. Du, J. Q. Wang, J. F. Kou, G. Y. Li, L. L. Wang, H. Chao and L. N. Ji, Eur. J. Med. Chem., 2011, 46, 1056–1065. 21 F. Gao, H. Chao, J. Q. Wang, Y. X. Yuan, B. Sun, Y. F. Wei, B. Peng and L. N. Ji, J. Biol. Inorg. Chem., 2007, 12, 1015–1027.

This journal is © The Royal Society of Chemistry 2014

Paper

22 F. A. Beckford, J. Thessing, M. J. Shaloski, P. C. Mbarushimana, A. Brock, J. Didion, J. Woods, A. G. Sarrías and N. P. Seeram, J. Mol. Struct., 2011, 992, 39–47. 23 X. Chen, F. Gao, Z. X. Zhou, W. Y. Yang, L. T. Guo and L. N. Ji, J. Inorg. Biochem., 2010, 104, 576–582. 24 Y. C. Wang, C. Qian, Z. L. Peng, X. J. Hou, L. L. Wang, H. Chao and L. N. Ji, J. Inorg. Biochem., 2014, 130, 15–27. 25 C. K. Johnson, ORTEP II, Report ORNL-5138, Oak Ridge National Laboratory, Oak Ridge, TN, 1976. 26 H. Chao, G. Yang, G. Q. Xue, H. Li, H. Zang, I. D. Williams, L. N. Ji, X. M. Chen and X. Y. Li, J. Chem. Soc., Dalton Trans., 2001, 8, 1326–1331. 27 L. Y. Li, H. N. Jia, H. J. Yu, K. J. Du, Q. T. Lin, K. Q. Qiu, H. Chao and L. N. Ji, J. Inorg. Biochem., 2012, 113, 31–39. 28 N. W. Alcock, P. R. Barker, J. M. Haider, M. J. Hannon, C. L. Painting, Z. Pikramenou, E. A. Plummer, K. Rissanen and P. Saarenketo, J. Chem. Soc., Dalton Trans., 2000, 1447–1461. 29 N. Grover, N. Gupta, P. Singh and H. H. Thorp, Inorg. Chem., 1992, 31, 2014–2020. 30 S. Satyanarayana, J. C. Dabroniak and J. B. Chaires, Biochemistry, 1992, 31, 9319–9324. 31 S. Satyanarayana, J. C. Daborusak and J. B. Chaires, Biochemistry, 1993, 32, 2573–2584. 32 T. M. Kelly, A. B. Tossi, D. J. McConnell and T. C. Strekas, Nucleic Acids Res., 1985, 13, 6017–6034. 33 E. Friedman, J. C. Chambron, J. P. Sauvage, N. J. Turro and J. K. Barton, J. Am. Chem. Soc., 1990, 112, 4960–4962. 34 R. B. Nair, E. S. Teng, S. L. Kirkland and C. J. Murphy, Inorg. Chem., 1998, 37, 139–141. 35 H. Chao, W. J. Mei, Q. W. Huang and L. N. Ji, J. Inorg. Biochem., 2002, 92, 165–170. 36 J. M. Fortune, L. Velea, D. E. Graves, T. Utsugi, Y. Yamada and N. Osheroff, Biochemistry, 1999, 38, 15580–15586. 37 P. Y. Zhang, J. Q. Wang, H. Y. Huang, L. P. Qiao, L. N. Ji and H. Chao, Dalton Trans., 2013, 42, 8907–8917. 38 C. Qian, J. Q. Wang, C. L. Song, L. L. Wang, L. N. Ji and H. Chao, Metallomics, 2013, 5, 844–854. 39 C. C. Sprenger, M. E. Vail, K. Evans, J. Simurdak and S. R. Plymate, Oncogene, 2002, 21, 140–147. 40 I. Vermes and C. Haanen, Adv. Clin. Chem., 1994, 31, 177–246. 41 R. V. Furth and T. L. Van Zwet, J. Immunol. Methods, 1988, 108, 45–51. 42 S. Savill, A. H. Wyllie, J. E. Henson, M. J. Walport, P. M. Henson and C. Haslett, J. Clin. Invest., 1989, 83, 865–875. 43 N. A. Thornberry and Y. Lazebnik, Science, 1998, 281, 1312– 1316. 44 H. L. Chan, D. L. Ma, M. Yang and C. M. Che, ChemBioChem, 2003, 4, 62–68. 45 B. Schutte, R. Nuydens, H. Geerts and F. Ramaekers, J. Neurosci. Methods, 1998, 86, 63–69. 46 I. Vermes, C. Haanen, H. S. Nakken and C. Reutelingsperger, J. Immunol. Methods, 1995, 184, 39–51. 47 A. Hartmann and G. Speit, Mutat. Res., 1995, 346, 49–56.

Dalton Trans., 2014, 43, 17303–17316 | 17315

View Article Online

Published on 24 September 2014. Downloaded by Ondoku Mayis Universitesi on 09/11/2014 06:48:58.

Paper

48 R. S. Hotchkiss, A. Strasser, J. E. McDunn and P. E. Swanson, N. Engl. J. Med., 2009, 361, 1570–1583. 49 C. Wang and R. J. Youle, Annu. Rev. Genet., 2009, 43, 95–118. 50 A. Cossarizza, M. B. Contri, G. Kalashnikova and C. Franceschi, Biochem. Biophys. Res. Commun., 1993, 197, 40–45. 51 J. Marmur, J. Mol. Biol., 1961, 3, 208–218. 52 M. E. Reichmann, S. A. Rice, C. A. Thomas and P. Doty, J. Am. Chem. Soc., 1954, 76, 3047–3053. 53 E. J. Corey, A. L. Borror and T. Foglia, J. Org. Chem., 1965, 30, 288–290. 54 T. Baker, P. Singh and V. Vignevich, Aust. J. Chem., 1991, 44, 1041–1048. 55 Y. J. Liu, J. C. Chen, F. H. Wu and K. C. Zheng, Transition Met. Chem., 2009, 34, 297–305. 56 P. Sullivan, J. M. Calvert and T. J. Meyer, Inorg. Chem., 1980, 19, 1404–1407. 57 R. H. Blessing, Acta Crystallogr., Sect. A: Found. Crystallogr., 1995, 51, 33–38.

17316 | Dalton Trans., 2014, 43, 17303–17316

Dalton Transactions

58 G. M. Sheldrick, SHELXS 97, Program for X-Ray Crystal Structure Determination, University of Göttingen, 1997; SHELXL 97, Program for X-Ray Crystal Structure Refinement, University of Göttingen, 1997. 59 G. Cohen and H. Eisenberg, Biopolymers, 1969, 8, 45–49. 60 M. T. Carter, M. Rodriguez and A. Bard, J. Am. Chem. Soc., 1989, 111, 8901–8911. 61 N. Osheroff, E. R. Shelton and D. L. Brutlag, J. Biol. Chem., 1983, 258, 9536–9543. 62 M. Fortune and N. Osheroff, J. Biol. Chem., 1998, 273, 17643–17650. 63 C. Alley, D. A. Scudiero, A. Monks, M. L. Hursey, M. J. Czerwinski, D. L. Fine, B. J. Abbott, J. G. Mayo, R. H. Shoemaker and M. R. Boyd, Cancer Res., 1988, 48, 589–601. 64 S. Po, Y. Li, T. Siu, M. Tang, K. Shek, M. Yiu, K. Kam and W. Lo, Chem. – Eur. J., 2012, 18, 13342–13354. 65 P. Singh, M. T. MacCoy, R. R. Tice and E. L. Schneider, Exp. Cell Res., 1988, 175, 184–191.

This journal is © The Royal Society of Chemistry 2014

Dual inhibition of topoisomerases I and IIα by ruthenium(II) complexes containing asymmetric tridentate ligands.

Five novel ruthenium(II) complexes, [Ru(dtzp)(dppt)](2+) (1), [Ru(dtzp)(pti)](2+) (2), [Ru(dtzp)(ptn)](2+) (3), [Ru(dtzp)(pta)](2+) (4) and [Ru(dtzp)(...
4MB Sizes 0 Downloads 5 Views