Synthesis, structures, and DNA and protein binding of ruthenium(II)-p-cymene complexes of substituted pyridylimidazo[1,5-a]pyridine: enhanced cytotoxicity of complexes of ligands appended with a carbazole moiety

Abstract

A series of organometallic Ru(II)-arene complexes of the type [(η⁶-p-cymene)Ru(L)Cl](BF₄) 1–6, where L is 3-phenyl-1-pyridin-2-yl-imidazo[1,5-a]pyridine (L1), dimethyl-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)phenyl]-amine (L2), diphenyl-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)phenyl]amine (L3), 9-[4-(1-pyridin-2-yl-imidazo-[1,5-a]pyridin-3-yl)-phenyl]-9H-carbazole (L4), 9-ethyl-3-(1-pyridin-2-yl-imidazo-[1,5-a]pyridin-3-yl)-9H-carbazole (L5), and 10-ethyl-3-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-10H-phenothiazine (L6), has been isolated and characterised by elemental analysis, ESI-MS, NMR and cyclic voltammetry. The photophysical properties of the complexes have been studied by electronic absorption and emission spectral techniques. All the ligands exhibit tuneable photoluminescence behaviour with the emission maximum spanning through the visible region (475–670 nm) in dichloromethane while all the complexes are emissive in acetonitrile. The single crystal X-ray structures of 2, 3 and 4 reveal that the complexes have a “piano stool” coordination geometry, comprising one π-bonded arene centroid, two σ-bonded nitrogen atoms from the chelating ligand and one Cl⁻ ion. From DNA induced EthBr emission quenching experiments the apparent DNA binding constants of the complexes (K_app) have been evaluated, which follows the order, 2 (1.3) < 1 (1.5) < 6 (1.7) < 4 (1.8) < 5 (2.4) < 3 (2.8 × 10⁵ M⁻¹). This trend reveals the role of ligand hydrophobicity on the DNA binding ability of complexes, the non-planar phenothiazine ring (6) and the specific interactions the planar carbazole chromophore (4, 5) make with DNA. The value of K_sv obtained as the slope of the linear plot of F₀/F vs. [complex] follows the order 1 (3.1) < 6 (8.2) < 2 (13.1) < 3 (15.7) < 5 (17.1) ≈ 4 (17.2 L M⁻¹), which supports the inferences from DNA binding experiments. All the complexes, except, 1 and 2 (>100 μM), exhibit in vitro cytotoxicity against A549 small lung cancer cell lines higher than cisplatin (∼69 μM), as revealed by both MTT (11.8–18.1 μM) and crystal violet staining (12.7–23.5 μM) assays, which is in agreement with their DNA and BSA binding affinity. Also, the complexes 3–6 cause higher cell death mainly through the apoptotic mode, as revealed by the observation of a higher percentage of apoptotic cells in AO/EB (36–43%) and Annexin V-Cy3 (36–45%) stained cancer cells.

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Introduction

It is a great challenge for a medicinal inorganic chemist to design and synthesize non-platinum anticancer agents with less toxicity because the most successful platinum-based anticancer drugs like cisplatin and other multi-nuclear platinum compounds, which are useful in the treatment of testicular cancer,^1,2 show several side effects including nephrotoxicity, emetogenesis and neurotoxicity during treatment of cancer.³ The two Ru(III) complexes Na[RuCl₄(Hind)₂] (NKP-1339, Hind = 1H-indazole)⁴ and (H₂im)[RuCl₄(DMSO)(Him)] (NAMI-A, Him = 1H-imidazole),⁵ which show promising anticancer activities, are in the advanced stages of clinical development.⁶ The former complex has shown activity in a variety of advanced and metastasized solid malignancies, notably in non-small cell lung cancer and gastro-intestinal neuroendocrine tumours,⁷ whereas the latter is active against the metastasis form of tumours with a current focus on lung cancer.⁸ It is evident that NKP-1339 acts as a prodrug and is reduced under the hypoxic conditions of the tumor tissue to the reactive Ru(II) species.^9–11 The reactive Ru(II) species can bind rapidly to molecular targets¹² and finally activate apoptosis via the mitochondrial pathway.¹³ In addition to NKP-1339 and NAMI-A, several other Ru(III) and Ru(II) complexes^14–19 have been shown to exhibit cytotoxic activity,⁵ which has now paved the way to the design and study of numerous ruthenium based anticancer agents.

Very recently, attention has been focused on organometallic Ru(II) arene complexes.^20–25 Sadler et al. have found that the Ru(II)-arene–en (en = ethylenediamine) complex exhibits efficient cytotoxic activity and also shows activity against cisplatin-resistant cell lines.^26–30 The Ru(II)-arene-PTA complex (RAPTA-C) acts as an antimetastatic agent and shows a biochemical mode of action against Ehrlich Ascites Carcinoma (EAC) cells, which is different from that of platinum anticancer drugs. It inhibits specific proteases, which play a vital role in anticancer drug development.^31,32 Very recently, Sheldrick and co-workers have prepared half-sandwich Ru(II)- and Rh(III)-arene complexes with methyl-substituted polypyridyl ligands, which strongly bind to DNA and also regulate apoptosis.³³ Pandey have et al. reported water soluble Ru(II)- and Rh(III)-arene complexes, which show DNA binding and topoisomerase II inhibitory activity.³⁴ The antitumor activity of many Ru(II)-arene complexes has been related to their enhanced DNA binding affinity, which involves covalent and/or non-covalent mode of DNA interaction.^35–43 In very recent years, there have been increasing investigations⁴⁴ on the interaction of metallo-drugs with molecular targets other than DNA for understanding their cytotoxic activities. As serum proteins perform the transport, distribution, accumulation and excretion properties of drugs in tumor tissue, special consideration has been given to the study of binding of metallo-drugs to the proteins.⁴⁵ The solubility of hydrophobic drugs in plasma has been found to effectively increase and the drugs bind to serum albumins and modulate their delivery to cells in specific sites. In this connection, the significant binding activity of RAPTA-C compounds with two emerging protein targets, thioredoxin reductase and cathepsin B, has been explored very recently for illustrating their anticancer activity.⁴⁶ Hence, designing of DNA and protein binding Ru(II)-organometallic complexes has currently received attention in the field of metallodrug discovery.

Very recently, one of us has reported DNA and protein binding and cleaving Ru(II) “piano-stool” complexes, which exhibit remarkable cytotoxicity against MCF-7 breast cancer cell lines.⁴⁷ In this article we report a series of organometallic Ru(II)-arene complexes of the type [(η⁶-p-cymene)-Ru(L)Cl]BF₄ 1–6, where L is 3-phenyl-1-pyridin-2-yl-imidazo[1,5-a]pyridine (L1), dimethyl-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-phenyl]amine (L2), diphenyl-[4-(1-pyridin-2-yl-imidazo-[1,5-a]pyridin-3-yl)-phenyl]-amine (L3), 9-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-phenyl]-9H-carbazole (L4), 9-ethyl-3-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-9H-carbazole (L5), and 10-ethyl-3-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-10H-phenothiazine (L6). We have preferred to use p-cymene as the arene ligand in this study as Ru(II)-p-cymene complexes show DNA and protein binding ability higher, and also cytotoxicity higher, than the analogous Ru(II)-benzene complexes.⁴⁷ The molecular structures of three representative complexes 2, 3 and 4 have been determined by using single crystal X-ray crystallography. The ability of the complexes with varying hydrophobicity to bind to calf thymus (CT) DNA and bovine serum albumin (BSA) protein have been studied by using emission spectral methods to understand the role of hydrophobicity on the ability of complexes to bind to these macromolecules. We have incorporated carbazole moiety in the ligands L4 and L5 because certain DNA binding carbazole derivatives have been very recently shown to exhibit potent cytotoxic activity toward P388 leukemia cells.⁴⁸ Also, the L6 ligand has been appended with phenothiazine moiety as phenothiazine is currently considered a prototypical pharmaceutical lead moiety⁴⁹ in medicinal chemistry. Also, the derivatives of phenothiazine are currently under investigation as possible anti-infective drugs^50–53 and in fact, its derivative methylene blue is one of the first antimalarial drugs. And, interestingly, the complexes with carbazole and phenothiazine moieties show cytotoxicity against A549 small lung cancer cell lines higher than cisplatin and those with carbazole moiety cause higher cell death mainly through apoptotic mode.

Experimental

Reagents and materials

Unless and otherwise specified, all the reactions and manipulations were performed under nitrogen atmosphere using standard Schlenk techniques. All of the reagents, benzaldehyde, 4-dimethylaminobenzaldehyde and di-pyridin-2-yl-methanone were procured from commercial sources and used without further purification. 4-Diphenylaminobenzaldehyde, 4-carbazol-9-yl-benzaldehyde, 9-ethyl-9H-carbazole-3-carbaldehyde and 10-ethyl-10H-phenothiazine-3-carbaldehyde were prepared as previously reported.^54–57 The ligands were synthesized with modified procedure as previously reported.⁵⁸ The dimeric complex [Ru(η⁶-p-cymene)Cl₂]₂ was prepared according to a published procedure.^59,60

General synthesis of ligands

The ligands were synthesized by essentially following the same procedure and so an illustrative procedure for L4 is provided below. The procedures for the synthesis of ligands are listed in Scheme 1.

Scheme 1 Structures of the ligands L1–L6.

9-[4-(1-Pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-phenyl]-9H-carbazole (L4). To a flask containing a mixture of 4-carbazol-9-yl-benzaldehyde (2.03 g, 7.5 mmol), di-pyridin-2-yl-methanone (0.92 g, 5.0 mmol), and NH₄(OAc) (1.93 g, 25 mmol) was added 25 mL of dry acetic acid. The solution mixture was slowly heated to reflux, stirred for 16 h, and then cooled, and 2 mL of water was added. The solution was pumped dry, and the residue was extracted with dichloromethane : water solvent mixture. The organic layer was dried over magnesium sulphate, filtered, and dried. The residue was chromatographed through silica gel (dichloromethane : hexane = 1 : 3 v/v) to give an yellow powder of L4. Yield: 48%; mp 198–202 °C; ¹H NMR (400 MHz, CDCl₃) δ 8.80–8.77 (d, 1H, J = 12 Hz), 8.67–8.68 (d, 1H, J = 4 Hz), 8.42–8.40 (d, 1H, J = 8 Hz), 8.31–8.29 (d, 1H, J = 8 Hz), 8.19–8.17 (d, 1H, J = 8 Hz), 8.12–8.10 (d, 2H, J = 8 Hz), 7.79–7.75 (t, 4H, J = 8 Hz), 7.53–7.51 (d, 1H, J = 8 Hz), 7.48–7.44 (t, 2H, J = 8 Hz), 7.35–7.31 (t, 3H, J = 8 Hz), 7.15–7.13 (t, 1H, J = 4 Hz), 7.02–6.98 (t, 1H, J = 8 Hz), 6.77–6.74 (t, 1H, J = 6 Hz); ¹³C NMR (400 MHz, CDCl₃) δ: 153.7, 147.9, 139.4, 136.9, 136.0, 135.2, 129.7, 129.3, 128.5, 127.9, 126.3, 125.0, 122.4, 120.8, 120.3, 120.1, 119.5, 119.3, 119.1, 118.8, 113.1, 108.6; anal. found (calcd) for C₃₀H₂₀N₄: C, 82.48 (82.55); H, 4.70 (4.62); N, 12.92 (12.84). HRMS (ESI) calcd for C₃₀H₂₁N₄: 437.1766 [M + H]⁺, found: 437.1761 [M + H]⁺.

3-Phenyl-1-pyridin-2-yl-imidazo[1,5-a]pyridine (L1). Yellow solid; yield: 56%; mp 92–96 °C; NMR (400 MHz, CDCl₃), δ 8.65–8.62 (d, 1H, J = 12 Hz), 8.57–8.56 (d, 1H, J = 4 Hz), 8.20–8.17 (d, 2H, J = 6 Hz), 7.78–7.77 (d, 2H, J = 4 Hz), 7.67–7.63 (t, 1H, J = 8 Hz), 7.50–7.46 (t, 2H, J = 8 Hz), 7.42–7.38 (t, 1H, J = 8 Hz), 7.04–7.02 (t, 1H, J = 4 Hz), 6.86–6.84 (t, 1H, J = 8 Hz), 6.60–6.57 (t, 1H, J = 6 Hz); ¹³C NMR (400 MHz, CDCl₃) δ: 155.0, 148.9, 138.0, 136.2, 130.5, 130.2, 130.1, 129.0, 128.9, 128.3, 121.7, 121.5, 121.0, 120.4, 119.9, 113.9; anal. found (calcd) for C₁₈H₁₃N₃: C, 79.59 (79.68); H, 4.75 (4.83); N, 15.58 (15.49). HRMS (ESI) calcd for C₁₈H₁₄N₃: 272.1188 [M + H]⁺, found: 272.1182 [M + H]⁺.

Dimethyl-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)phenyl]amine (L2). Yellow solid; yield: 71%; mp 176–180 °C; ¹H NMR (400 MHz, CDCl₃) δ 8.7–8.65 (t, 2H, J = 10 Hz), 8.3–8.28 (d, 1H, J = 8 Hz), 8.23–8.21 (d, 1H, J = 8 Hz), 7.75–7.69 (m, 3H), 7.12–7.10 (d, 1H, J = 8 Hz), 6.93–6.85 (m, 3H), 6.64–6.61 (t, 1H, J = 6 Hz), 3.05 (s, 6H); ¹³C NMR (400 MHz, CDCl₃) δ: 155.1, 150.7, 148.8, 139.0, 136.2, 129.7, 129.3, 121.8, 121.6, 120.6, 120.1, 119.9, 117.4, 113.4, 112.3, 40.3; anal. found (calcd) for C₂₀H₁₈N₄: C, 76.55 (76.41); H, 5.80 (5.77); N, 17.52 (17.82). HRMS (ESI) calcd for C₂₀H₁₉N₄: 315.1610 [M + H]⁺, found: 315.1604 [M + H]⁺.

Diphenyl-[4-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)phenyl]amine (L3). Yellow solid; yield: 62%; mp 238–242 °C; ¹H NMR (400 MHz, CDCl₃) δ 8.71–8.69 (d, 1H, J = 8 Hz), 8.63 (s 1H), 8.26 (s, 2H), 7.70–7.68 (d, 3H, J = 8 Hz), 7.32–7.26 (m, 5H), 7.20–7.16 (m, 5H), 7.10–7.08 (d, 3H, J = 8 Hz), 6.92 (s, 1H), 6.67–6.64 (t, 1H, J = 6 Hz); ¹³C NMR (400 MHz, CDCl₃) δ: 153.9, 147.8, 147.4, 146.2, 137.0, 135.1, 129.2, 128.9, 128.3, 128.1, 123.7, 122.3, 122.3, 122.1, 120.7, 120.6, 119.8, 119.3, 118.8, 112.7; anal. found (calcd) for C₃₀H₂₂N₄: C, 82.29 (82.17); H, 5.12 (5.06); N, 12.70 (12.78). HRMS (ESI) calcd for C₃₀H₂₃N₄: 439.1923 [M + H]⁺, found: 439.1917 [M + H]⁺.

9-Ethyl-3-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-9H-carbazole (L5). Yellow solid; yield: 54%; mp 167–170 °C; ¹H NMR (400 MHz, CDCl₃) δ 8.66–8.64 (d, 1H, J = 8 Hz), 8.59–8.58 (d, 1H, J = 4 Hz), 8.47 (s 1H), 8.27–8.22 (t, 2H, J = 6 Hz), 8.10–8.08 (d, 1H, J = 8 Hz), 7.85–7.83 (d, 1H, J = 8 Hz), 7.68–7.65 (t, 1H, J = 6 Hz), 7.50–7.38 (m, 3H), 7.22–7.18 (t, 1H, J = 8 Hz), 7.05–7.02 (t, 1H, J = 6 Hz), 6.88–6.84 (t, 1H, J = 8 Hz), 6.60–6.56 (t, 1H, J = 8 Hz), 4.39–4.34 (m, 2H), 1.41–1.39 (t, 3H, J = 8 Hz); ¹³C NMR (400 MHz, CDCl₃) δ: 155.3, 149.0, 140.4, 140.1, 139.3, 136.2, 130.2, 129.9, 126.2, 123.4, 122.8, 121.7, 121.7, 120.9, 120.8, 120.7, 120.5, 120.3, 119.9, 119.3, 113.6, 109.0, 108.8, 37.7, 13.8; anal. found (calcd) for C₂₆H₂₀N₄: C, 80.52 (80.39); H, 5.10 (5.19); N, 14.35 (14.42). HRMS (ESI) calcd for C₂₆H₂₁N₄: 389.1766 [M + H]⁺, found: 389.1716 [M + H]⁺.

10-Ethyl-3-(1-pyridin-2-yl-imidazo[1,5-a]pyridin-3-yl)-10H-phenothiazine (L6). Yellow solid; yield: 48%; mp 162–165 °C; ¹H NMR (400 MHz, CDCl₃) δ 8.71–8.67 (d, 1H, J = 16 Hz), 8.22 (s, 2H), 7.76 (s, 1H), 7.61 (s, 1H), 7.26–6.94 (m, 9H), 6.68 (s, 1H), 4.0 (s, 2H), 1.47 (s, 3H); ¹³C NMR (400 MHz, CDCl₃) δ: 153.9, 147.8, 144.2, 143.2, 136.2, 135.2, 129.2, 129.0, 126.3, 126.3, 126.2, 125.7, 123.9, 122.9, 122.5, 121.6, 120.6, 120.5, 119.8, 119.3, 118.8, 114.0, 112.8, 40.8, 11.8; anal. found (calcd) for C₂₆H₂₀N₄S₁: C, 74.38 (74.26); H, 4.82 (4.79); N, 13.45 (13.32). ESI-MS m/z 420 (M⁺). HRMS (ESI) calcd for C₂₆H₂₁N₄S: 421.1487 [M + H]⁺, found: 421.1473 [M + H]⁺.

Synthesis of complexes

The [Ru(p-cymene)(L)Cl]BF₄ complexes 1–6 were prepared by essentially following the same procedure and an illustrative example is provided below for 1 (Scheme 2).

Scheme 2 Synthesis of complexes 1–6.

[Ru(η⁶-cymene)(L1)Cl]BF₄ (1). [Ru(p-cymene)(Cl)₂]₂ (0.12 g, 0.2 mmol) and L1 (0.1 g, 0.4 mmol) were suspended in methanol (20 mL) and stirred at room temperature for 2 h. A solution of NaBF₄ (200 mg, 0.60 mmol) in MeOH (10 mL) was added to the initially orange solution, which changed colour to yellow. After 24 h the solution was evaporated and the solid obtained was filtered off. The residue was washed with diethyl ether (40 mL) and dried under vacuum. The desired products were recrystallized from DCM : hexane solvent mixture to give orange coloured microcrystals. Yield: 57%; ¹H NMR (400 MHz, CDCl₃) δ 9.27–9.26 (d, 1H, J = 4 Hz), 8.10–8.08 (d, 2H, J = 8 Hz), 8.03–8.00 (d, 1H, J = 12 Hz), 7.96–7.95 (d, 2H, J = 4 Hz), 7.8 (s, 4H), 7.42–7.41 (d, 1H, J = 4 Hz), 7.34–7.30 (t, 1H, J = 8 Hz), 6.97–6.94 (t, 1H, J = 6 Hz), 5.59–5.58 (d, 1H, J = 4 Hz), 5.35–5.33 (d, 1H, J = 8 Hz), 5.18–5.17 (d, 1H, J = 4 Hz), 4.78–4.76 (d, 1H, J = 8 Hz), 2.51–2.45 (m, 1H), 2.12 (s, 6H), 1.16 (s, 3H). Anal. found (calcd) for C₂₈H₂₇BClF₄N₃Ru: C, 53.30 (53.48); H, 4.24 (4.33); N, 6.56 (6.68). ESI-MS (m/z): 542.13 [M − BF₄]⁺.

[Ru(η⁶-cymene)(L2)Cl]BF₄ (2). Orange solid; yield: 89%; ¹H NMR (400 MHz, DMSO-d₆) δ 9.40–9.39 (d, 1H, J = 4 Hz), 8.46–8.36 (m, 3H), 8.19–8.15 (t, 1H, J = 8 Hz), 7.96–7.94 (d, 2H, J = 8 Hz), 7.54–7.47 (m, 2H), 7.17–7.12 (m, 3H), 5.91–5.90 (d, 1H, J = 4 Hz), 5.60–5.59 (d, 1H, J = 4 Hz), 5.26–5.25 (d, 1H, J = 4 Hz), 4.95–4.93 (d, 1H, J = 8 Hz), 3.17 (s, 6H), 2.56 (s, 6H), 2.46–2.43 (m, 1H), 2.16 (s, 3H). Anal. found (calcd) for C₃₀H₃₂BClF₄N₄Ru: C, 53.65 (53.62); H, 4.93 (4.80); N, 8.25 (8.34). ESI-MS (m/z): 585.32 [M − BF₄]⁺.

[Ru(η⁶-cymene)(L3)Cl]BF₄ (3). Orange solid; yield: 84%; ¹H NMR (400 MHz, DMSO-d₆) δ 9.35–9.33 (d, 1H, J = 8 Hz), 8.42–8.39 (d, 2H, J = 12 Hz), 8.34–8.32 (d, 1H, J = 8 Hz), 8.13–8.09 (t, 1H, J = 8 Hz), 7.92 (s, 1H), 7.45–7.42 (t, 6H, J = 6 Hz), 7.26–7.11 (m, 10H), 5.86 (s, 1H), 5.59–5.58 (d, 1H, J = 4 Hz), 5.36 (s, 1H), 4.84 (s, 1H), 2.38–2.35 (m, 1H), 2.04 (s, 3H), 0.85 (s, 6H). Anal. found (calcd) for C₄₀H₃₆BClF₄N₄Ru: C, 60.40 (60.35); H, 4.39 (4.56); N, 7.10 (7.04). ESI-MS (m/z): 707.94 [M − BF₄]⁺.

[Ru(η⁶-cymene)(L4)Cl]BF₄ (4). Orange solid; yield: 78%; ¹H NMR (400 MHz, CDCl₃) δ 9.40–9.39 (d, 1H, J = 4 Hz), 8.59–8.58 (d, 2H, J = 4 Hz), 8.50–8.31 (m, 5H), 8.18–8.13 (t, 3H, J = 10 Hz), 7.69–7.67 (d, 2H, J = 8 Hz), 7.54–7.53 (d, 4H, J = 4 Hz), 7.39–7.35 (t, 2H, J = 8 Hz), 7.21–7.18 (t, 1H, J = 6 Hz), 5.94–5.93 (d, 1H, J = 4 Hz), 5.62–5.61 (d, 1H, J = 4 Hz), 5.42–5.41 (d, 1H, J = 4 Hz), 4.98 (s, 1H), 2.49 (s, 6H), 2.31–2.28 (m, 1H), 2.12 (s, 3H). Anal. found (calcd) for C₄₀H₃₄BClF₄N₄Ru: C, 60.62 (60.50); H, 4.40 (4.32); N, 7.17 (7.06). ESI-MS (m/z): 707.42 [M − BF₄]⁺.

[Ru(η⁶-cymene)(L5)Cl]BF₄ (5). Orange solid; yield: 88%; ¹H NMR (400 MHz, DMSO-d₆) δ 9.36 (s, 1H), 8.99–8.87 (m, 1H), 8.46–8.09 (m, 7H), 7.78–7.12 (m, 6H), 5.79–5.76 (d, 1H, J = 12 Hz), 5.50 (s, 1H), 5.17 (s, 1H), 4.62 (s, 3H), 2.5 (s, 3H), 2.05–2.03 (d, 1H, J = 8 Hz), 1.43 (s, 3H), 0.8 (s, 6H). Anal. found (calcd) for C₃₆H₃₄BClF₄N₄Ru: C, 57.84 (57.96); H, 4.50 (4.59); N, 7.38 (7.51). ESI-MS (m/z): 659.37 [M − BF₄]⁺.

[RuCl(η⁶-cymene)(L6)Cl]BF₄ (6). Orange solid; yield: 90%; ¹H NMR (400 MHz, DMSO-d₆) δ 9.39–9.38 (d, 1H, J = 4 Hz), 8.46–8.35 (m, 4H), 8.20–8.16 (t, 2H, J = 8 Hz), 7.97–7.90 (m, 2H), 7.53–7.07 (m, 7H), 5.8 (s, 1H), 5.6 (s, 1H), 5.43–5.42 (d, 1H, J = 4 Hz), 4.14–4.12 (d, 2H, J = 8 Hz), 2.38–2.40 (m, 1H), 2.12 (s, 3H), 1.48–1.45 (t, 3H J = 6 Hz), 0.8 (s, 6H). Anal. found (calcd) for C₃₆H₃₄BClF₄N₄RuS: C, 55.68 (55.57); H, 4.21 (4.40); N, 7.28 (7.20). ESI-MS (m/z): 690.23 [M − BF₄]⁺.

Experimental methods

All chromatographic separations were carried out on silica gel (60 M, 230–400 mesh). Dichloromethane and dimethylformamide (DMF) were distilled from calcium hydride under nitrogen atmosphere. Other solvents were purified by routine procedures. ¹H spectra were recorded on a Bruker Avance II 400 MHz spectrometer. Mass spectra (FAB) were recorded on a Thermo LC-MS instrument. Elemental analyses were performed on a Perkin-Elmer 2400 CHN analyzer. Electronic absorption spectra were measured on a Cary 50 Probe UV-visible spectrometer. Emission spectra were recorded on a Hitachi fluorescence spectrometer (F4500). Cyclic voltammetry experiments were performed with a BAS-100 electrochemical analyzer. All measurements were carried out at room temperature with a conventional three-electrode configuration consisting of a glassy carbon working electrode, platinum auxiliary electrode, and a non-aqueous Ag/AgNO₃ reference electrode. The E_1/2 values were determined as 1/2(E_pa + E_pc), where E_pa and E_pc are the anodic and cathodic peak potentials, respectively. All the potentials reported are not corrected for the junction potential. The solvent in all experiments was CH₂Cl₂, and the supporting electrolyte was 0.1 M tetra-n-butylammonium hexafluorophosphate.

Solutions of calf thymus (CT) DNA in the buffer 5 mM Tris–HCl/50 mM NaCl in water gave a ratio of UV absorbance at 260 and 280 nm, A₂₆₀/A₂₈₀, of 1.9,^61,62 indicating that the DNA was sufficiently free of protein. Concentrated stock solutions of DNA (14.8 mM) were prepared in buffer and sonicated for 25 cycles, where each cycle consisted of 30 s with 1 min intervals. The concentration of DNA in nucleotide phosphate (NP) was determined by UV absorbance at 260 nm after 1 : 100 dilutions by taking the extinction coefficient, ε = 260, as 6600 M⁻¹ cm⁻¹. Stock solutions of DNA were stored at 4 °C and used after not more than 4 days. Concentrated stock solutions of metal complexes were prepared by dissolving calculated amounts of ruthenium complexes in respective amounts of solvent and diluted suitably with the corresponding buffer to required concentrations for all experiments.

X-ray crystallographic analysis

Single crystals for X-ray diffraction studies were grown by slow diffusion of hexane in a CH₂Cl₂ solution of complexes 2, 3 and 4 at 20 °C. All crystals were mounted on glass fibers for data collection. Data were collected on an Oxford Diffraction Xcalibur Eos Gemini diffractometer. Crystal data were collected at ambient temperature using graphite-monochromated Mo-Kα radiation (λ = 0.7107 Å). The data were solved using direct methods with SHELXS and refined using SHELXL-2013.⁶³ The graphics interface package used was PLATON, and the figures were generated using the ORTEP 3.07 generation package.⁶⁴ The positions on all the atoms were obtained by direct methods. Metal atoms in each complex were located from the E-maps and non-hydrogen atoms were refined anisotropically. The hydrogen atoms bound to the carbon were placed in geometrically constrained positions and refined with isotropic temperature factors, generally 1.2U_eq. of their parent atoms. The crystallographic data and details of data collection for 2–4 are given in Table 1.

Table 1 Crystallographic data and structure refinement parameters for 2, 3 and 4

a R1 = ∑||Fo| − |Fc||/∑|Fo|.b wR2 = {∑w[(Fo^2 − Fc^2)^2]/∑w[(Fo^2)^2]}1/2.
Compound	2	3	4
Formula	C30H32BN4F4ClRu	C40H36BN4F4ClRu	C40H34BN4F4ClRu
Crystal system	Monoclinic	Monoclinic	Monoclinic
Space group	P21/c	P21/c	P21/n
a (Å)	15.9526 (5)	12.4248 (3)	12.0308 (5)
b (Å)	7.7424 (3)	11.9615 (4)	11.1279 (4)
c (Å)	24.6748 (10)	24.8642 (6)	27.6398 (9)
α (deg)	90	90	90
β (deg)	106.237 (4)	97.198 (2)	102.338 (4)
γ (deg)	90	90	90
V (Å^3)	2926.06 (19)	3666.18 (18)	3614.9 (2)
Z	4	4	4
μ (mm^−1)	0.676	0.556	0.555
ρcalc mg mm^−3	1.5207	1.3873	1.4001
Final R indices	R1 = 0.0426	R1 = 0.0526	R1 = 0.0687
	wR2 = 0.0937	wR2 = 0.1427	wR2 = 0.1785
R1^a	0.0564	0.0742	0.0981
wR2^b	0.0101	0.1613	0.2020

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DNA binding experiments

Concentrated stock solutions of metal complexes were prepared by dissolving them in a 10% DMF–5 mM Tris–HCl/50 mM NaCl buffer (0.5 mL of DMF in 5 mL of buffer) at pH 7.1 and diluting suitably with the corresponding buffer to required concentrations for all the experiments. For emission spectral experiments the DNA solutions were pretreated with solutions of metal complexes to ensure no change in the metal complex concentrations. The tris buffer was used as a blank to make preliminary adjustments. The excitation wavelength was fixed and the emission range was adjusted before measurements. DNA was pretreated with ethidium bromide in the ratio ([NP]/[EthBr]) = 10 for 30 min at 27 °C. The metal complexes were then added to this mixture and their effect on the emission intensity was measured. Fluorescence spectral measurements were performed using a 1 cm quartz cell on a Cary Eclipse Fluorescence Spectrophotometer (G9800A).

BSA binding experiments

The protein binding study was performed by tryptophan fluorescence quenching experiments using bovine serum albumin (BSA, 5 μM) as the substrate in phosphate buffer (pH 6.8). Quenching of the emission intensity of tryptophan residues of BSA at 344 nm (excitation wavelength at 295 nm) was monitored using complexes 1–6 as quenchers with increasing complex concentration. The F₀/F versus [complex] plot was constructed using the corrected fluorescence data taking into account the effect of dilution. The excitation and emission slit widths were 2.5 and 2.5 nm, respectively. Fluorescence measurements were performed using a 1 cm quartz cell on a Cary Eclipse Fluorescence Spectrophotometer (G9800A).

Cell culture

The A549 small lung cancer cell lines were obtained from the National Center for Cell Science (NCCS), Pune, India. The cells were cultured in RPMI 1640 medium (Biochrom AG, Berlin, Germany) supplemented with 10% fetal bovine serum (Sigma, USA), cisplatin (Getwell Pharmaceuticals, India), mitomycin C (Sigma, USA) and 100 U mL⁻¹ penicillin and 100 μg mL⁻¹ streptomycin as antibiotics (Himedia, Mumbai, India) in 96 well culture plates at 37 °C in a humidified atmosphere of 5% CO₂ in a CO₂ incubator (Heraeus, Hanau, Germany). All experiments were performed using cells from passage 15 or less.

MTT assay

The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay was carried out as described previously.⁶⁵ The ruthenium complexes, in the concentration range of 0–100 μM, dissolved in 100% dimethyl sulfoxide (DMSO) (Sigma-Aldrich) and prepared to final dilution of DMSO to 0.02%, was added to the well, 24 h after seeding 5000 cells (A549) per well of 96-well plate. DMSO (0.02%) was used as the solvent control and cisplatin was used as the positive control. After 24 h incubation, 20 μL of MTT solution [5 mg mL⁻¹ in phosphate-buffered saline (PBS)] was added to each well, and the plates were wrapped with aluminium foil and incubated for 3 h at 37 °C. The purple formazan product was dissolved by addition of 100 μL of 100% DMSO to each well. The absorbance was monitored at 570 nm (measurement) and 630 nm (reference) using a 96-well plate reader (Bio-Rad, Hercules, California, USA). Data were collected for three replicates, each in triplicate wherein the three average values were used to calculate the means and the standard deviations. The percentage inhibition was calculated from this data using the following formula:

From the values thus obtained, the IC₅₀ values for 24 h treatment, for A549 cells, were deduced from the curves obtained by plotting percentage inhibition against concentration.

Crystal violet assay

The cytotoxicity of complexes was measured by crystal violet assay (CVS) according to the work published in the literature.⁶⁶ The different concentrations of working solution was prepared by dissolving ruthenium complexes in 100% dimethyl sulfoxide (DMSO) (Sigma-Aldrich) and added to the well, 24 h after seeding 5000 cells (A549) per well of 96-well plate. DMSO (0.02%) was used as the solvent control and cisplatin was used as the positive control. After 24 h incubation, the medium was removed and washed the cells twice in a gentle stream of tap water. The plate was allowed to dry completely and 50 μL of 0.5% crystal violet staining solution was added to the well. After, 20 minutes incubation at room temperature, the plate was washed four times in a stream of tap water. The plate was allowed to dry for 2 h at room temperature. Finally, 200 μL of methanol was added to the wells, and incubated for 20 minutes at room temperature. The optical density of each well at 570 nm (OD₅₇₀) was measured with plate reader (Bio-Rad, Hercules, California, USA). Data were collected for three replicates, each in triplicate, wherein the three average values were used to calculate the means and standard deviations. The average OD₅₇₀ of non-treated cells was set as 100%. Then the percentage of viability in treated samples was measured by comparing the average OD₅₇₀ values of treated cells with the OD₅₇₀ values of non-treated cells.

Acridine orange (AO) and ethidium bromide (EB) fluorescent assay

AO and EB staining was performed as described by Spector et al.⁶⁷ A549 cells were cultured separately in 6-well plates and treated with IC₅₀ concentrations of the complexes for 24 h, when DMSO (0.02%) was used as solvent control. The treated and untreated cells (25 μL of suspension containing 5000 cells) were incubated with acridine orange (AO) and ethidium bromide (EB) solutions (1 part of 100 μg mL⁻¹ each of AO and EB in PBS) and examined in a fluorescent microscope (Carl Zeiss, Jena, Germany) using a UV filter (450–490 nm). Three hundred cells per sample were counted, in triplicate, for each time point and scored as viable or dead, and if dead, whether by apoptosis or necrosis as judged from the nuclear morphology and cytoplasmic organization. The percentages of apoptotic and necrotic cells were then calculated. Morphological features of interest were photographed.

Annexin V-Cy3 staining

Phosphatidylserine translocation from the inner to outer leaflet of the plasma membrane is one of the early apoptotic features. Cell surface phosphatidylserine was detected by phosphatidylserine binding protein annexin V conjugated with Cy3 using the commercially available Annexin V-Cy3 apoptosis detection kit (APOAC, Apoptosis Detection Kit, Sigma).⁶⁸ Cells were cultured on cover slips and treated with IC₅₀ concentrations of complexes 3–6 and incubated for 24 h. The cell pellet was washed with PBS and then with 1× binding buffer. The washed cell pellet was suspended in 50 μL of double label staining solution (Ann-Cy3 and 6-CFDA) and kept in dark for 10 min. After the incubation, the excess label was removed by washing the cells with 1× binding buffer. The annexin-Cy3 and 6-CFDA-labelled cells were observed in the fluorescent microscope. 300 cells at random were observed. This assay facilitated detection of live cells (green), necrotic cells (red), and apoptotic cells (red nuclei and green cytoplasm). The percentage of cells reflecting cell death (apoptotic and necrotic, separately) was calculated. Data were collected from three individual experiments, each in triplicate, and used to calculate the respective means and the standard deviations.

Results and discussions

The pyridylimidazo[1,5-a]pyridine based bidentate ligands (L1–L6) were synthesized by condensing di-pyridin-2-yl-methanone with aldehydes in acetic acid solution. The ligands were characterized by ¹H NMR spectra. Mononuclear arene ruthenium complexes [Ru(p-cymene)(L)Cl]BF₄ have been prepared by treating a slight excess of the corresponding ligand with [Ru(p-cymene)Cl₂]₂ in methanol as solvent. The products were isolated as tetrafluoroborate salts in good yields. The complexes have been isolated as yellow coloured powders. Based on elemental analysis and ESI-MS the complexes were formulated as [Ru(p-cymene)(L)Cl]BF₄, and the stoichiometry of 2, 3 and 4 is confirmed by single crystal X-ray structure determination. The ESI-MS data reveal that the complexes retain their identity even in solution, which is supported by values of molar conductivity in acetonitrile (Λ_M/Ω⁻¹ cm² mol⁻¹: 82–89), falling in the range for 1 : 1 electrolytes. The complexes 1–6 are moderately soluble in common organic solvents such as MeCN and MeOH. They are stable in the solid state as well as in solution. They were further characterised in DMF solvent due to their higher solubility at various interval of time. The ¹H NMR spectra of 1–6 in CDCl₃ show downfield shifts in the positions of arene protons due to the incorporation of the L1/L2 ligand, which modifies the electron density around metal centre as compared to the corresponding precursor complexes. The cyclic voltammetric (CV) responses obtained in acetonitrile solution reveal that the Ru(II)/Ru(III) redox couples of 1–6 are far from reversible (E_1/2, 0.326–0.344 V).

Description of the crystal structures of [Ru(η⁶-cymene)L(Cl)]BF₄ (2, 3 and 4)

Crystal structure determination data are summarized in Table 1. The selected bond distances and bond angles are tabulated in Table 2. The perspective views of the crystal structures of 2, 3 and 4 are depicted in Fig. 1. The asymmetric units of all the complexes contain a cationic complex molecule and a BF₄⁻ counter ion. In the complex cations Ru(II) is coordinated to all the six carbon atoms (C1–C6) of the p-cymene ring, both the nitrogen atoms of bidentate ligands and one chloride ion (Cl⁻). The coordination geometry around Ru(II) in the complexes is best described as pseudo-octahedral with the arene ring occupying the three coordination sites in a η⁶-fashion and the two nitrogen atoms of the bidentate ligands and a chloride ion occupying the remaining coordination sites. The complex cations adopt the familiar “three leg piano-stool” structure^69,70 with the η⁶-arene ring forming the seat and the two nitrogen atoms of ligand and one Cl⁻ constituting the three legs of the stool, as evident from the nearly 90° bond angles N_py–Ru–Cl (84.21–86.78°) and N_im–Ru–Cl (86.19–86.29°). The Ru–N bond lengths fall in the range 2.093–2.106 Å (2: 2.106, 2.093; 3: 2.099, 2.098; 4: 2.096, 2.104 Å) and the Ru–Cl bond length in the range 2.393–2.405 Å (2: 2.405, 3: 2.393, 4: 2.401 Å).^71–73 The six Ru–C bonds have almost comparable bond distances (2: 2.181–2.231; 3: 2.156–2.251; 4: 2.165–2.251 Å) with an average Ru–C bond lengths of 2.201 (2); 2.196 (3); 2.199 (4) Å and the ruthenium-arene centroid ring distance (Ru–Ct, 1.689 Å) is almost the same for all the complexes and is similar to those in related Ru(II)-arene complexes.^74–77

Table 2 Selected bond lengths (Å) and angles (°) for 2, 3 and 4

	2	3	4
Bond lengths
Ru–Npy	2.106 (2)	2.099 (3)	2.096 (5)
Ru–Nim	2.093 (2)	2.098 (3)	2.104 (4)
Ru–Cl	2.405 (8)	2.393 (11)	2.400 (2)
Ru–C1	2.231 (3)	2.251 (4)	2.250 (6)
Ru–C2	2.205 (3)	2.210 (4)	2.233 (6)
Ru–C3	2.181 (3)	2.156 (4)	2.165 (8)
Ru–C4	2.219 (3)	2.194 (5)	2.189 (9)
Ru–C5	2.185 (3)	2.179 (4)	2.166 (7)
Ru–C6	2.190 (3)	2.191 (4)	2.195 (7)
Ru–Cave	2.201 (3)	2.196 (4)	2.199 (7)
Bond angles
Npy–Ru–Nim	77.15 (10)	76.480 (11)	76.56 (15)
Npy–Ru–Cl	84.21 (7)	86.786 (9)	84.37 (17)
Nim–Ru–Cl	86.19 (7)	86.286 (10)	86.29 (14)

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Fig. 1 ORTEP drawings of 2, 3 and 4. Hydrogen atoms and BF₄ anion are omitted for clarity.

Absorption and emission spectral properties

The absorption and emission spectra of the ligands L1–L6 and their Ru(II) complexes 1–6 were investigated in dichloromethane and acetonitrile solutions respectively, and the pertinent data are collected in Table 3. In the absorption spectra, all the ligands display a band located in the range 336–400 nm, which possibly originates from the π → π* transitions^78,79 located in the chromophores such as substituted phenyl, carbazole and phenothiazine moieties (Fig. 2). Within the ligand series the wavelength of this absorption band adopts the trend L1 < L2 > L3 > L4 < L5 > L6. The incorporation of –NMe₂ group on the phenyl ring of L1 to obtain L2 causes red-shift in the band position illustrating that the electron-releasing group –NMe₂ group enhances the electron density on the aromatic rings.^80–82 On the other hand, the replacement of methyl groups on –NMe₂ moiety by two phenyl groups leads to a blue-shift, which is expected of the electron-withdrawing phenyl groups. The linking of the phenyl groups in L3 to get L4 enhances the electron-withdrawing effect and causes a further blue-shift. As expected, the incorporation of one phenyl group as in L5 causes a lower blue-shift compared to L2, and the separation of two phenyl rings in L5 by a sulphur atom to get L6 causes a higher blue shift. Additional more intense bands in the lower wavelength region (326–223 nm) are attributable to the n → π*/π → π* and other related transitions located in the imidazo[1,5-a]pyridine core. All the ligands are moderately emissive in DCM solution with the emitted colour varying from yellow to green depending on the substituents on the imidazo[1,5-a]pyridine core. The variation observed in the emission maximum L1 < L2 > L3 > L4 < L5 < L6 (Fig. 3) is the same as that observed for the π → π* transition, as expected, confirming the manifestation of the effect of different organic structural components.^83,84 The electronic absorption spectra of 1–6 (Fig. 4) obtained in acetonitrile solution contains features in the range 238–400 nm. All the complexes exhibit intense absorption bands in the range 290–320 nm, assigned to intraligand π–π* transitions. The lowest-energy absorption in each case (385–410 nm) is tentatively assigned as metal-to-ligand charge transfer (MLCT) transition from the 4d orbitals of Ru(II) to the empty π* ligand orbitals.^85–87 All of the metal complexes exhibit a red-shift in the longer wavelength band (336–400 nm) of the corresponding ligands; however, interestingly, the trend in the band maximum is the same as that observed for the ligands. This illustrates that the variation in ligand moieties determine the position of the bands even after complexation. All the complexes exhibit moderate luminescence at room temperature in CH₃CN solution (Fig. 5). Within the series, the wavelength of emission band maximum follows the trend, 1 < 2 > 3 > 4 > 5 < 6, which is similar to that discerned in the emission spectra of the corresponding ligands L1–L6. This illustrates the role of ligand moieties in determining the emission properties even after complexation.

Table 3 Absorption and emission spectral and electrochemical properties of ligands and complexes

Compound	λabs^a, nm (ε, M^−1 cm^−1)	λem^a, nm	E1/2^b (V)
a Ligands in dichloromethane and complexes in acetonitrile solution.b Potential measured vs. Ag(s)/Ag^+ (0.01 M, 0.10 M TBAP).
L1	387, 325, 295(sh), 229	460
L2	400, 326, 269, 227	495
L3	346, 304, 232, 224	480
L4	336, 293, 283(sh), 235	465
L5	395, 315, 287, 270, 242, 223	475
L6	365, 322, 260, 227	480
1	398 (7170), 378 (10 620), 360 (8330), 311 (9050), 292 (10 160), 247 (8830)	460	1.44
2	388 (25 785), 301 (52 595), 247 (26 160)	505	0.97, 1.22
3	381 (27 105), 337 (40 770), 305 (43 770), 229 (40 280)	485	1.13, 1.34, 1.48
4	401 (17 250), 379 (25 985), 337 (33 500), 325 (34 910), 310 (35 795), 291 (41 325)	465	1.29, 1.35
5	407 (16 280), 384 (22 575), 291 (52 040), 266 (42 865), 238 (60 915)	500	1.28, 1.43
6	405 (19 545), 383 (26 895), 287 (39 565), 261 (48 940)	480	0.83, 1.35

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Fig. 2 Absorption spectra of ligands L1–L6 in dichloromethane.

Fig. 3 Emission spectra (normalized) of the ligands L1–L6 in dichloromethane.

Fig. 4 Absorption spectra of complexes 1–6 (5 × 10⁻⁵ M) in acetonitrile.

Fig. 5 Emission spectra (normalized) of complexes 1–6 in acetonitrile.

Electrochemical studies

The electrochemical properties of the present complexes were studied by cyclic voltammetry at a platinum electrode. The measurements were performed on ca. 0.5 mM acetonitrile solutions containing 0.1 M [Bu₄N][PF₆] as the supporting electrolyte. The complexes exhibit a single-electron oxidation wave assigned to Ru(II) → Ru(III) oxidation^88–91 (0.83–1.44 V, Table 3, Fig. 6). The Ru(II)/Ru(III) redox potentials of the complexes vary in the order 1 (1.44) > 2 (0.97) < 3 (1.03) < 4 (1.33) > 5 (1.28) > 6 (0.83 V), illustrating the influence of ligand σ-donating and π-accepting properties on the Ru(II) → Ru(III) oxidation. Thus the incorporation of –NMe₂ group with electron-donating methyl substituents on amine nitrogen in 1 to give 2 facilitates oxidation of Ru(II) center while that of –NPh₂ group with electron-withdrawing phenyl substituents to give 3 discourages it. Similarly, 4 with two electron-withdrawing but linked phenyl groups on nitrogen shows an oxidation wave at a more positive potential (1.33 V), and 5 and 6, both with one ethyl and one phenyl groups on nitrogen, show lower redox potentials of 1.28 (5) and 0.83 V (6). Additional quasi-reversible waves corresponding to oxidation of amine moieties are discernible for 3 and 4. Thus incorporation of electron-donating and electron-withdrawing groups on the bidentate ligand results in variation in electron density at Ru(II) centre, which in turn manifests on the redox propensity of the complexes.

Fig. 6 Cyclic voltammogram of 1 recorded in acetonitrile solution (1 × 10⁻³ M) at 25 °C. Supporting electrolyte: 0.1 M TBAP. Scan rate of 100 mV s⁻¹.

Interaction of Ru(II) complexes with DNA and protein

Competitive DNA-binding studies. A competitive DNA binding study was performed to understand the mode of DNA interaction of the Ru(II) complexes 1–6. All the present complexes, which exhibit emission in acetonitrile solution, fail to show steady-state emission in 10% DMF–5 mM Tris–HCl–50 mM NaCl buffer at pH 7.1 at 25 °C and lack emission even in the presence of CT DNA (R = 25). So the extent of DNA binding of the complexes has been evaluated using competitive DNA binding studies involving ethidium bromide (EthBr), which is known to emit strongly (λ_ex, 450; λ_em, 595 nm) due to its intercalative interaction with DNA. Addition of a complex molecule, which binds to DNA more strongly than EthBr, quenches the DNA-induced EthBr emission.⁹² Upon adding 1–6 (0–60 μM) to CT DNA (125 μM) with EthBr (12.5 μM) in 10% DMF–5 mM Tris–HCl–50 mM NaCl buffer at pH 7.1, the emission intensity of DNA-bound EthBr decreases enormously (Fig. 7), and the emission intensity is quenched completely at higher concentrations. This reveals the ability of complexes to bind to DNA, displace DNA-bound EthBr and quench the DNA induced EthBr emission. To quantify the displacement, the concentration of the complex at which EthBr fluorescence decreases by 50% (assumed to be 50% displacement of EthBr) is calculated.⁹³ The apparent DNA binding constant (K_app) was calculated⁹⁴ from a plot of the observed intensities against complex concentration used to titrate with DNA bound with EthBr using the equation,

K_EthBr[EthBr] = K_app[complex]

where the value of K_EthBr is 4.94 × 10⁵ M⁻¹, is the DNA binding constant of EthBr, [EthBr] is the concentration of EthBr (12.5 μM) and [complex] is the concentration of the complex used to obtain 50% reduction in fluorescence intensity of EthBr bound to DNA.⁹⁵ The observed ability of the Ru(II)-arene complexes to quench the DNA induced EthBr emission and hence to bind to DNA, that is, the apparent DNA binding constant (K_app) vary in the order, 1 (1.5) > 2 (1.3) < 3 (2.8) > 4 (1.8) < 5 (2.4) > 6 (1.7 × 10⁴ M⁻¹). The values observed are of the same order as those determined for the analogous [(η⁶-p-cymene)Ru(L)Cl]⁺ complexes, where L is 1-(anthracen-10-ylmethyl)-4-methylhomo-piperazine,⁴⁷ suggesting that the methyl and isopropyl groups on coordinated cymene contribute to the higher DNA binding affinity of the complexes. Also, for the latter complexes and certain other complexes⁹⁶ the experimental values of K_app parallel the observed values of intrinsic DNA binding constant K_b and so the present values of K_app can be reliably used to illustrate the DNA binding affinity of the present complexes. Thus the DNA binding affinity of 3 and 5 is higher than other complexes; obviously, the coordinated ligands containing freely rotating N-phenyl (3) and N-ethyl (5) groups are involved in stronger hydrophobic interaction (cf. below) with the hydrophobic surface of DNA. The higher DNA binding affinity of 4 and 5 is consistent with the presence of planar carbazole ring of the ligands, which may fit with the minor groove surface of DNA or stack on DNA base pairs.⁴⁸ The DNA binding affinity of 3 is higher than that of 4 as the two phenyl rings in the former are expected to show higher hydrophobic interaction. The complex 6 shows DNA binding affinity lower than 5, which is expected of the inability of non-planar phenothiazine ring⁹⁷ in the former to involve itself in partial intercalative DNA interaction. As expected, the complexes 1 and 2, which have no hydrophobic or partially intercalating bidentate ligand moieties, exhibit lower DNA binding affinities. It is possible that the complexes interact with DNA leading to destabilization of exciplex of EthBr bound to DNA by electron-transfer from Ru(II) complex to the excited state, but the observed redox potentials of the complexes do not parallel their DNA binding affinity, confirming the involvement of displacement mechanism in quenching EthBr emission.

Fig. 7 (A) Effect of addition of 3 on the emission intensity of CT DNA-bound ethidium bromide (12.5 μM) at different concentrations in Tris buffer (pH 7.1). (B) Plots of relative integrated emission intensity (F/F₀) vs. [complex] for 1–6.

Tryptophan quenching studies. To investigate the protein binding affinity of the Ru(II)-arene complexes 1–6, tryptophan emission-quenching experiments were carried out by exciting the tryptophan emission of bovine serum albumin (BSA, 5 μM) at 295 nm, adding the complex in increasing concentration (0–30 μM) to it at 298 K with the incubation period of 5 min and following the decrease in fluorescence intensity. The emission intensity of BSA, measured as described in the Experimental section, is found to decrease in the presence of 1–6. As the emission intensity of BSA depends on the degree of exposure of the two hydrophobic tryptophan side chains, 134 and 212 of the protein, to polar solvents and also on their proximity to specific quenching groups, such as protonated carbonyl, protonated imidazole, deprotonated-amino groups, and tyrosinate anions,⁹⁸ it is evident that changes in the protein secondary structure causing changes in tryptophan environment of BSA occur upon binding of the complexes.^99,100 Generally, the fluorescence quenching is illustrated by the well-known Stern–Volmer equation,

F₀/F = 1 + K_sv[Q]

where F₀ and F are the fluorescence intensities of BSA in the absence and presence of the complexes respectively, and K_sv is the Stern–Volmer quenching constant. The value of K_sv obtained (Table 4) as the slope of the linear plot of F₀/F vs. [complex] (Fig. 8, 0–30 μM) follows the order, 1 (3.1) < 2 (13.1) > 3 (15.7) < 4 (17.2) ≈ 5 (17.1) > 6 (8.2 L M⁻¹). This suggests that the protein-binding affinity of 4 and 5 are the highest on account of the enhanced hydrophobicity of their carbazole moiety and that the ligand hydrophobicity contributes to the enhanced BSA protein binding affinity of the complexes.¹⁰¹ These observations are in good agreement with those from competitive DNA binding studies (cf. above), confirming that 4 and 5 show specific hydrophobic forces of interactions involving the carbazole moiety and the hydrophobic DNA surface as well as tryptophan pockets in BSA.

Table 4 DNA binding properties, Stern–Volmer quenching constants and in vitro cytotoxicity assay for complexes 1–6

Compound	Kapp^a (10^5 M^−1)	Ksv^b (L mol^−1)	IC50^c^,^d (μM)	IC50^c^,^e (μM)
a Apparent DNA binding constant from ethidium bromide displacement assay using increasing concentrations (0–60 μM) of 1–6.b [BSA] = 5 μM; increasing concentration (0–50 μM) of 1–6.c IC50 = concentration of the drug required to inhibit the growth of 50% of cancer cells (in μM).d Data obtained from MTT assay at 24 h incubation.e Data obtained from CVS assay at 24 h incubation.
1	1.5	3.1 ± 0.3	>100	>100
2	1.3	13.1 ± 1.6	>100	>100
3	2.8	15.7 ± 2.7	18.1 ± 0.2	16.3 ± 0.2
4	1.8	17.2 ± 2.4	15.0 ± 0.2	18.7 ± 0.5
5	2.4	17.1 ± 1.4	17.5 ± 0.5	12.7 ± 0.2
6	1.7	8.2 ± 1.9	11.8 ± 0.5	23.5 ± 0.5
Cisplatin			71.0 ± 2.0	67.4 ± 2.0

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Fig. 8 (A) Effect of addition of 5 on the emission intensity of BSA (2.5 μM) at different concentrations in phosphate buffer (pH 7.0). (B) Plots of relative integrated emission intensity (F/F₀) vs. [complex] for 1–6.

Cytotoxicity of Ru(II) complexes

MTT and crystal violet staining (CVS) assay. The cytotoxicity of the present complexes against A549 small lung cancer cell lines has been investigated in aqueous buffer solution in comparison with the widely used drug cisplatin under identical conditions by using the MTT and CVS assay. The cytotoxicity of 1 and 2 are lower than that of cisplatin and does not exhibit efficient cytotoxicity till 100 μM. The IC₅₀ values (Table 4) obtained by plotting the cell viability determined using MTT assay against different concentrations of complexes reveal that the cytotoxicity of 3–6 are higher than that of cisplatin for 24 h incubation and follows the order 6 > 4 > 5 > 3. In contrast, the IC₅₀ values of the complexes determined by CVS assay follows the reverse order 5 > 3 > 4 > 6. Very recently it has been shown¹⁰² that the cytotoxicity, as determined by CVS assay, is quick and reliable as it is a non-enzymatic assay, which involves determination of amount of crystal violet dye absorbed by the total DNA content and hence the quantity of viable adherent cells in the culture under diverse stimulation conditions.¹⁰² On the other hand, though one of the most popular tests to assess the activity of potential anticancer compounds, the MTT assay, which is based on the assumption that reduction of MTT tetrazolium salt to formazan occurs in the mitochondria of living cells due to the enzymatic activity of mitochondrial dehydrogenases,¹⁰³ may be influenced by compounds that modify cell metabolism by increasing the NADPH level or the activity of lactate dehydrogenase (LDH).¹⁰⁴ So it is possible that the present complexes may influence the loss of mitochondrial membrane potential and modify the metabolism of the cells, leading to affect the IC₅₀ value determined by MTT assay. Thus the 1C₅₀ value based on CVS assay is consistent with the DNA and protein binding studies (cf. above), and it is evident that the complex 5, which shows higher DNA and BSA binding ability, exhibits the lowest IC₅₀ value and hence the highest cytotoxicity among the complexes in the series, and also six times higher than cisplatin. It is noteworthy that the compound 6, which exhibits the highest cytotoxicity in the MTT assay, displays the highest IC₅₀ value in CVS assay. In this regard, it may be noted that phenothiazine has attracted considerable interest as a lead structural moiety in current medicinal chemistry investigations, and phenothiazine derivatives have been found to show promising antitumor activities.⁵⁰ So, it is evident that the phenothiazine moiety appended in 6 contributes to the decreased cytotoxicity as well as apoptotic potential (cf. below) due to complexation. Also, generally, complexes with higher ligand hydrophobicity and higher DNA and protein binding affinity are expected⁴⁷ to display higher ability to penetrate the cell membrane and hence show higher ability to kill cancer cells. We have previously shown⁴⁷ that Ru(II)-arene complexes with higher ligand hydrophobicity and hence lipophilicity show higher ability to cross the cell membrane and exhibit higher cytotoxicity. The enhanced hydrophobicity of p-cymene may also contribute to the prominent cytotoxicity of the complexes. Also, very recently, it has been demonstrated that hydrophobicity of certain Ru(III) complexes correlate with their lipophilicity, which is a determining factor for cytotoxicity, and hence pharmacological behaviour.¹⁰⁵ So the highest cytotoxicity of 5 may be attributed, in addition to its higher DNA and protein binding affinity, to specific molecular interactions its ligand carbazole moiety exhibits with cell membrane so that it penetrates it easily and binds with the cellular components.

Acridine orange (AO) and ethidium bromide (EB) fluorescent assay method. The ability of 3–6 to induce apoptotic morphologies has been investigated by using acridine orange/ethidium bromide (AO/EthBr) staining and adopting fluorescence microscopy (Fig. 9). The cytological changes observed may be classified¹⁰⁶ into the following four types according to the fluorescence emission and morphological features of chromatin condensation in the AO/EthBr stained nuclei: (i) viable cells having uniformly green fluorescing nuclei with highly organized structure; (ii) early apoptotic cells (which still have intact membranes but have started undergoing DNA fragmentation) having green fluorescing nuclei, and peri-nuclear chromatin condensation visible as bright green patches or fragments; (iii) late apoptotic cells having orange to red fluorescing nuclei with condensed or fragmented chromatin; and (iv) necrotic cells, swollen to large sizes, having uniformly orange to red fluorescing nuclei with no indication of chromatin fragmentation. These morphological changes and manual counting data (Fig. 10) observed for 3–6, which exhibit higher cytotoxicity, suggest that the cells are committed to very efficient apoptotic cell death and necrosis to a certain extent. The complexes may also bind to the cellular proteins involved in inducing cancer and cause cell death mainly through the apoptotic mode. Interestingly, the complex 5, which shows the highest DNA and BSA binding ability, exhibits a higher percentage of apoptotic cell death than other complexes.

Fig. 9 AO/EB staining of A549 lung cancer cells treated with complexes 3–6 at 24 h of incubation.

Fig. 10 Graph showing manual count of apoptotic and necrotic cells in percentage when complexes 3–6 treated with A549 lung cancer cells at 24 h of incubation, as revealed by AO/EB fluorescent assay (data are mean% ± SD% of each triplicate).

Annexin V-Cy3 staining. An early indicator of apoptosis is the rapid translocation and accumulation of the membrane phospholipid phosphatidylserine from the cytoplasmic interface to the extracellular surface.¹⁰⁷ The combination of Annexin V-Cy3 (emits red) and 6-CFDA (emits green) allows the differentiation among early apoptotic cells (annexin V positive, 6-CFDA positive), necrotic cells (annexin V positive, 6-CFDA negative), and viable cells (annexin V negative, 6-CFDA positive). The results obtained with annexin V binding assay of control and treated cells are represented in Fig. 11. When treated with 1–6, there is a significant increase in number of cells positive for both annexin V and 6-CFDA indicating early stage of apoptosis. The results of Annexin V-Cy3 substantiate that the complexes might selectively trigger apoptotic mode of cell death than necrosis (Fig. 12). It is noteworthy that the complex 5 exhibits higher percentage of apoptotic cell death than other complexes, as evident from both AO/EB and Annexin V-Cy3 staining assays.

Fig. 11 A549 lung cancer cells stained with Annexin V-Cy3 and 6-CFDA. Cells were treated with 3, 4, 5 and 6 for 24 h.

Fig. 12 Percentage of A549 cells in apoptosis and necrosis, control and 3, 4, 5 and 6 treated, as evaluated by Annexin V-Cy3 staining studies (data are mean% ± SD% of each triplicate).

Conclusions

In this study, a family of half-sandwich Ru(II)-p-cymene complexes of the type [(η⁶-p-cymene)Ru(L)Cl]BF₄ 1–6 has been isolated and well-characterized. The complexes 2, 3 and 4 adopt the familiar pseudo-octahedral “three-leg piano-stool” structure. All the complexes are emissive in acetonitrile solution, but not in aqueous solution at room temperature. The hydrophobic interaction of complexes with –NPh₂ (3), planar carbazole ring (4, 5) and –NEt group on carbazole ring (5) and that of methyl and isopropyl groups on the p-cymene ligand lead to enhanced DNA and protein binding affinities for the complexes. The complexes 3–6 are highly cytotoxic in vitro, and induce early apoptosis in A549 small lung cancer cells. The most active complex 5, appended with ligand carbazole moiety, displays cytotoxicity and ability to induce apoptosis higher than other complexes, and so, it is suggested as a potential anticancer drug candidate demanding extended in vitro and in vivo studies. Also, a suitable ligand moiety such as carbazole, along with the coordinated p-cymene and other DNA as well as protein recognition elements on ligands, may be incorporated into ruthenium-based organometallic compounds when designing efficient cytotoxic drugs.

Acknowledgements

Themmila Khamrang thanks University Grants Commission (UGC), New Delhi, India for providing financial support in the form of UGC-JRF. We thank DST-PURSE, NEHU-SAIF Shillong, India for providing XRD and NMR spectral facilities. One of us (MP) thanks Indian National Science Academy (INSA) for Senior Scientist position, Science and Engineering Research Board (SERB), New Delhi for a research scheme EMR/2015/002222 and Council of Scientific & Industrial Research (CSIR), India for a research scheme CSIR/01(2462)/11/EMR-II. One of the authors M. V. express his thanks to UGC, MRP-MAJOR-CHEM-2013-5144, (69/2014 F.No. 10-11/12) for financial assistance in the form of a major sponsored project. One of us (VR) thanks SERB, New Delhi for a Start-Up Research Grant (Young Scientists) – Chemical Sciences (SB/FT/CS-187/2011). Mrs Gowdhami Balakrishnan, Mahatma Gandhi-Doerenkamp Center for Alternatives to Use of Animals in Life Science Education, Bharathidasan University, Tiruchirappalli is thanked for performing a few biological studies.

References

1. E. Wiltshaw, Platinum Met. Rev., 1979, 23, 90–98.
2. N. J. Wheate and J. G. Collins, Curr. Med. Chem.: Anti-Cancer Agents, 2005, 5, 267–279.
3. C. X. Zhang and S. J. Lippard, Curr. Opin. Chem. Biol., 2003, 7, 481–489.
4. R. Trondl, P. Heffeter, C. R. Kowol, M. A. Jakupec, W. Berger and B. K. Keppler, Chem. Sci., 2014, 5, 2925–2932.
5. E. Alessio, G. Mestroni, A. Bergamo and G. Sava, Curr. Top. Med. Chem., 2004, 4, 1525–1535.
6. P.-S. Kuhn, V. Pichler, A. Roller, M. Hejl, M. A. Jakupec, W. Kandioller and B. K. Keppler, Dalton Trans., 2015, 44, 659–668.
7. D. S. Thompson, G. J. Weiss, S. Fields Jones, H. A. Burris, R. K. Ramanathan, J. R. Infante, A. Ogden and D. D. Von Hoff, J. Clin. Oncol., 2012, 30, 3033.
8. A. Bergamo, C. Gaiddon, J. H. M. Schellens, J. H. Beijnen and G. Sava, J. Inorg. Biochem., 2012, 106, 90–99.
9. P. Schluga, C. G. Hartinger, A. Egger, E. Reisner, M. Galanski, M. A. Jakupec and B. K. Keppler, Dalton Trans., 2006, 1796–1802.
10. M. J. Clarke, Coord. Chem. Rev., 2003, 236, 209–233.
11. M. A. Jakupec, E. Reisner, A. Eichinger, M. Pongratz, V. B. Arion, M. Galanski, C. G. Hartinger and B. K. Keppler, J. Med. Chem., 2005, 48, 2831–2837.
12. E. Reisner, V. B. Arion, B. K. Keppler and A. J. L. Pombeiro, Inorg. Chim. Acta, 2008, 361, 1569–1583.
13. P. Heffeter, K. Böck, B. Atil, M. A. R. Hoda, W. Körner, C. Bartel, U. Jungwirth, B. K. Keppler, M. Micksche, W. Berger and G. Koellensperger, J. Biol. Inorg. Chem., 2010, 15, 737–748.
14. B. S. Murray, M. V. Babak, C. G. Hartinger and P. J. Dyson, Coord. Chem. Rev., 2016, 306, 86–114.
15. V. Rajendiran, M. Murali, E. Suresh, S. Sinha, K. Somasundaram and M. Palaniandavar, Dalton Trans., 2008, 148–163.
16. P. Kumar, R. K. Gupta and D. S. Pandey, Chem. Soc. Rev., 2014, 43, 707–733.
17. M. R. Gill and J. A. Thomas, Chem. Soc. Rev., 2012, 41, 3179–3192.
18. S. Medici, M. Peana, V. M. Nurchi, J. I. Lachowicz, G. Crisponi and M. A. Zoroddu, Coord. Chem. Rev., 2015, 284, 329–350.
19. V. Rajendiran, M. Murali, E. Suresh, M. Palaniandavar, V. S. Periasamy and M. A. Akbarsha, Dalton Trans., 2008, 2157–2170.
20. A. K. Singh, D. S. Pandey, Q. Xu and P. Braunstein, Coord. Chem. Rev., 2014, 270–271, 31–56.
21. A. A. Nazarov, C. G. Hartinger and P. J. Dyson, J. Organomet. Chem., 2014, 751, 251–260.
22. S. K. Singh and D. S. Pandey, RSC Adv., 2014, 4, 1819–1840.
23. W. H. Ang, A. Casini, G. Sava and P. J. Dyson, J. Organomet. Chem., 2011, 696, 989–998.
24. Q. Wu, K. Zheng, S. Liao, Y. Ding, Y. Li and W. Mei, Organometallics, 2016, 35, 317–326.
25. G. S. Fink, Dalton Trans., 2010, 39, 1673–1688.
26. A. F. A. Peacock and P. J. Sadler, Chem.–Asian J., 2008, 3, 1890–1899.
27. C. G. Hartinger, N. M. Nolte and P. J. Dyson, Organometallics, 2012, 31, 5677–5685.
28. I. R. Canelon, L. Salassa and P. J. Sadler, J. Med. Chem., 2013, 56, 1291–1300.
29. R. Carter, A. Westhorpe, M. J. Romero, A. Habtemariam, C. R. Gallevo, Y. Bark, N. Menezes, P. J. Sadler and R. A. Sharma, Sci. Rep., 2016, 6, 20596.
30. J. J. S. Barreda and P. J. Sadler, Curr. Opin. Chem. Biol., 2015, 25, 172–183.
31. S. Chatterjee, S. Kundu, A. Bhattacharyya, C. G. Hartinger and P. J. Dyson, J. Biol. Inorg. Chem., 2008, 13, 1149–1155.
32. B. Wu, M. S. Ong, M. Groessl, Z. Adhireksan, C. G. Hartinger, P. J. Dyson and C. Davey, Chem.–Eur. J., 2011, 17, 3562–3566.
33. Y. Geldmacher, R. Rubbiani, P. Wefelmeier, A. Prokop, I. Ott and W. S. Sheldrick, J. Organomet. Chem., 2011, 696, 1023–1031.
34. S. K. Singh, S. Joshi, A. R. Singh, J. K. Saxena and D. S. Pandey, Inorg. Chem., 2007, 46, 10869–10876.
35. Y. K. Yan, M. Melchart, A. Habtemariam and P. J. Sadler, Chem. Commun., 2005, 4764–4776.
36. H. Chen, J. A. Parkinson, S. Parsons, R. A. Coxall, R. O. Gould and P. J. Sadler, J. Am. Chem. Soc., 2002, 124, 3064–3082.
37. G. Gasser, I. Ott and N. M. Nolte, J. Med. Chem., 2011, 54, 3–25.
38. M. Melchart and P. J. Sadler, Bioorganometallics, ed. G. Jaouen, Wiley-VCH, Weinheim, 2006, pp. 39–64.
39. H.-K. Liu and P. J. Sadler, Acc. Chem. Res., 2011, 44, 349–359.
40. A. M. Pizarro, A. Habtemariam and P. J. Sadler, Top. Organomet. Chem., 2010, 32, 21–56.
41. G. Jaouen and N. M. Nolte, Medicinal Organometallic Chemistry, Springer-Verlag, Heidelberg, 2010, vol. 32, pp. 21–56.
42. E. Alessio, Bioinorganic Medicinal Chemistry, Wiley VCH, Weinheim, 2011, p. 151.
43. C. G. Hartinger and P. J. Dyson, Chem. Soc. Rev., 2009, 38, 391–401.
44. I. Ascone, L. Messori, A. Casini, C. Gabbiani, A. Balerna, F. D. Unto and A. C. Castellano, Inorg. Chem., 2008, 47, 8629–8634.
45. A. R. Timerbaev, C. G. Hartinger, S. S. Aleksenko and B. K. Keppler, Chem. Rev., 2006, 106, 2224–2248.
46. A. Casini, C. Gabbiani, F. Sorrentino, M. P. Rigobello, A. Bindoli, T. J. Geldbach, A. Marrone, N. Re, C. G. Hartinger, P. J. Dyson and L. Messori, J. Med. Chem., 2008, 51, 6773–6781.
47. M. Ganeshpandian, R. Loganathan, E. Suresh, A. Riyasdeen, M. A. Akbarsha and M. Palaniandavar, Dalton Trans., 2014, 43, 1203–1219.
48. U. Jacquemard, S. Routier, A. Tatibouët, J. Kluza, W. Laine, C. Bal, C. Bailly and J.-Y. Mérour, Org. Biomol. Chem., 2004, 2, 1476–1483.
49. A. Jaszczyszyn, K. Gąsiorowski, P. Świątek, W. Malinka, K. C. Boczula, J. Petrus and B. C. Matusewicz, Pharmacol. Rep., 2012, 64, 16–23.
50. K. Pluta, B. M. Mlodawska and M. Jeleń, Eur. J. Med. Chem., 2011, 46, 3179–3189.
51. N. Motohashi, M. Kawase, S. Saito and H. Sakagami, Curr. Drug Targets, 2000, 1, 237–245.
52. D. Zong, K. Z. Chomej, T. Juntti, B. Mörk, R. Lewensohn, P. Hååg and K. Viktorsson, Cell Death Dis., 2014, 5, e1111.
53. C. M. Abuhaie, E. Bîcu, B. Rigo, P. Gautret, D. Belei, A. Farce, J. Dubois and A. Ghinet, Bioorg. Med. Chem. Lett., 2013, 23, 147–152.
54. S.-L. Deng, T.-L. Chen, W.-L. Chien and J.-L. Hong, J. Mater. Chem. C, 2014, 2, 651–659.
55. N.-G. Kang, B. Cho, B.-G. Kang, S. Song, T. Lee and J.-S. Lee, Adv. Mater., 2012, 24, 385–390.
56. C. Bian, G. Jiang, H. Tong, Y. Cheng, Z. Xie, L. Wang, X. Jing and F. Wang, J. Polym. Sci., Part A: Polym. Chem., 2011, 49, 3911–3919.
57. X. Yang, R. Lu, P. Xue, B. Li, D. Xu, T. Xu and Y. Zhao, Langmuir, 2008, 24, 13730–13735.
58. J. Wang, L. Dyers, R. Mason, P. Amoyaw and X. R. Bu, J. Org. Chem., 2005, 70, 2353–2356.
59. M. A. Bennett and A. K. Smith, J. Chem. Soc., Dalton Trans., 1974, 233–241.
60. M. A. Bennett, T.-N. Huang, T. W. Matheson and A. K. Smith, Inorg. Synth., 1982, 21, 74.
61. C. R. Merril, D. Goldman, S. A. Sedman and M. H. Ebert, Science, 1980, 211, 1437–1438.
62. R. Loganathan, S. Ramakrishnan, E. Suresh, A. Riyasdeen, M. A. Akbarsha and M. Palaniandavar, Inorg. Chem., 2012, 51, 5512–5532.
63. G. M. Sheldrick, Acta Crystallogr., Sect. A: Found. Crystallogr., 2008, 64, 112–122.
64. M. N. Burnett and G. K. Johnson, ORTEP III, Report ORNL-6895, OAK Ridge National Laboratory, Tennessee, USA, 1996.
65. M. V. Blagosklonny and W. S. E. Deiry, Int. J. Cancer, 1996, 67, 386–392.
66. M. Feoktistova, P. Geserick and M. Leverkus, Crystal violet assay for determining viability of cultured cells, Cold Spring Harbor Protocols, 2015a, 343–346.
67. Cells: A Laboratory Manual, Culture and Biochemical Analysis of Cells, ed. D. L. Spector, R. D. Goldman and L. A. Leinwand, Cold Spring Harbor Laboratory Press, New York, 1998, vol. 1.
68. A. Riyasdeen, V. S. Periasamy, P. Preethy, A. A. Alshatwi and M. A. Akbarsha, J. Evidence-Based Complementary Altern. Med., 2012, 2012, 136527.
69. L. He, S.-Y. Liao, C.-P. Tan, R.-R. Ye, Y.-W. Xu, M. Zhao, L.-N. Ji and Z.-W. Mao, Chem.–Eur. J., 2013, 19, 12152–12160.
70. J. J. S. Barreda, P. C. A. Bruijnincx, A. Habtemariam, G. J. Clarkson, R. J. Deeth and P. J. Sadler, Organometallics, 2012, 31, 5958–5967.
71. R. E. Morris, R. E. Aird, P. S. Murdoch, H. Chen, J. Cummings, N. D. Hughes, S. Parsons, A. Parkin, G. Boyd, D. I. Jodrell and P. J. Sadler, J. Med. Chem., 2001, 44, 3616–3621.
72. S. Betanzos-Lara, O. Novakova, R. J. Deeth, A. M. Pizarro, G. J. Clarkson, B. Liskova, V. Brabec, P. J. Sadler and A. Habtemariam, J. Biol. Inorg. Chem., 2012, 17, 1033–1051.
73. M. V. Babak, S. M. Meier, A. A. Legin, M. S. Adib Razavi, A. Roller, M. A. Jakupec, B. K. Keppler and C. G. Hartinger, Chem.–Eur. J., 2013, 19, 4308–4318.
74. H. Türkmen, I. Kani and B. Çetinkaya, Eur. J. Inorg. Chem., 2012, 4494–4499.
75. C. Aliende, M. P. Manrique, F. A. Jaloń, B. R. Manzano, A. M. Rodríguez and G. Espino, Organometallics, 2012, 31, 6106–6123.
76. C. Romain, S. Gaillard, M. K. Elmkaddem, L. Toupet, C. Fischmeister, C. M. Thomas and J. Renaud, Organometallics, 2010, 29, 1992–1995.
77. Y. Fu, A. Habtemariam, A. M. Pizarro, S. H. Rijt, D. J. Healey, P. A. Cooper, S. D. Shnyder, G. J. Clarkson and P. J. Sadler, J. Med. Chem., 2010, 53, 8192–8196.
78. S. B. Lara, L. Salassa, A. Habtemariam, O. Novakova, A. M. Pizarro, G. J. Clarkson, B. Liskova, V. Brabec and P. J. Sadler, Organometallics, 2012, 31, 3466–3479.
79. C. Garino, T. Ruiu, L. Salassa, A. Albertino, G. Volpi, C. Nervi, R. Gobetto and K. I. Hardcastle, Eur. J. Inorg. Chem., 2008, 3587–3591.
80. S. J. Dougan, M. Melchart, A. Habtemariam, S. Parsons and P. J. Sadler, Inorg. Chem., 2006, 45, 10882–10894.
81. L. Salassa, C. Garino, A. Albertino, G. Volpi, C. Nervi, R. Gobetto and K. I. Hardcastle, Organometallics, 2008, 27, 1427–1435.
82. M. Velusamy, Y.-C. Hsu, J. T. Lin, C.-W. Chang and C.-P. Hsu, Chem.–Asian J., 2009, 5, 87–96.
83. G. A. Ardizzoia, S. Brenna, S. Durini and B. Therrien, Polyhedron, 2015, 90, 214–220.
84. G. Volpi, C. Garino, L. Salassa, J. Fiedler, K. I. Hardcastle, R. Gobetto and C. Nervi, Chem.–Eur. J., 2009, 15, 6415–6427.
85. K. S. Singh and W. Kaminsky, Inorg. Chim. Acta, 2011, 365, 487–491.
86. W. F. Schmid, R. O. John, V. B. Arion, M. A. Jakupec and B. K. Keppler, Organometallics, 2007, 26, 6643–6652.
87. T. Tsolis, M. J. Manos, S. Karkabounas, I. Zelovitis and A. Garoufis, J. Organomet. Chem., 2014, 768, 1–9.
88. R. K. Gupta, R. Pandey, G. Sharma, R. Prasad, B. Koch, S. Srikrishna, P.-Z. Li, Q. Xu and D. S. Pandey, Inorg. Chem., 2013, 52, 3687–3698.
89. F. Marchetti, C. Pettinari, R. Pettinari, A. Cerquetella, A. Cingolani, E. J. Chan, K. Kozawa, B. W. Skelton, A. H. White, R. Wanke, M. L. Kuznetsov, L. M. D. R. S. Martins and A. J. L. Pombeiro, Inorg. Chem., 2007, 46, 8245–8257.
90. X. Shang, T. F. S. Silva, L. M. D. R. S. Martins, Q. L. M. Fátima, C. G. Silva, M. L. Kuznetsov and A. J. L. Pombeiro, J. Organomet. Chem., 2013, 730, 137–143.
91. Y. Sun, Y.-C. Dai, X. Wang, J.-M. Cheng and W.-G. Jia, J. Coord. Chem., 2016, 69, 48–56.
92. P. Uma Maheswari and M. Palaniandavar, J. Inorg. Biochem., 2004, 98, 219–230.
93. M. Lee, A. L. Rhodes, M. D. Wyatt, S. Forrow and J. A. Hartley, Biochemistry, 1993, 32, 4237–4245.
94. B. C. Baguley and M. LeBret, Biochemistry, 1984, 23, 937–943.
95. S. Satyanarayana, J. C. Dabrowiak and J. B. Chaires, Biochemistry, 1992, 31, 9319–9324.
96. V. Rajendiran, R. Karthik, M. Palaniandavar, H. S. Evans, V. S. Periasamy, M. A. Akbarsha, B. S. Srinag and H. Krishnamurthy, Inorg. Chem., 2007, 46, 8208–8221.
97. T. Kurihara, K. Shinohara, M. Inabe, H. Wakabayashi, N. Motohashi, H. Sakagami and J. Molnar, Top. Heterocycl. Chem., 2008, 15, 253–279.
98. T. Peters, Adv. Protein Chem., 1985, 37, 161–245.
99. C. J. Halfman and T. Nishida, Biochim. Biophys. Acta, 1971, 243, 284–293.
100. C. J. Halfman and T. Nishida, Biochim. Biophys. Acta, 1971, 243, 294–303.
101. N. S. Quiming, R. B. Vergel, M. G. Nicolas and J. A. Villanueva, J. Health Sci., 2005, 51, 8–15.
102. K. Chiba, K. Kawakami and K. Tohyama, Toxicol. in Vitro, 1998, 12, 251–258.
103. T. Mosmann, J. Immunol. Methods, 1983, 65, 55–63.
104. D. T. Vistica, P. Skehan, D. Scudiero, A. Monks, A. Pittman and M. R. Boyd, Cancer Res., 1991, 1, 2515–2520.
105. S. W. Chang, A. R. Lewis, K. E. Prosser, J. R. Thompson, M. Gladkikh, M. B. Bally, J. J. Warren and C. J. Walsby, Inorg. Chem., 2016, 55, 4850–4863.
106. S. Ramakrishnan, V. Rajendiran, M. Palaniandavar, V. S. Periasamy, B. S. Srinag, H. Krishnamurthy and M. A. Akbarsha, Inorg. Chem., 2009, 48, 1309–1322.
107. Y. Wu, N. Tibrewal and R. B. Birge, Trends Cell Biol., 2006, 16, 189–197.

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Footnote

† Electronic supplementary information (ESI) available: Crystallographic data for compounds 2, 3 and 4, cif files. CCDC 1484099, 1484100 and 1484101. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c6ra23663d

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