Protein Binding, and Anticancer Activity of

Article pubs.acs.org/Organometallics

Synthesis, Structure, DNA/Protein Binding, and Anticancer Activity of Some Half-Sandwich Cyclometalated Rh(III) and Ir(III) Complexes Sujay Mukhopadhyay,† Rakesh Kumar Gupta,† Rajendra Prasad Paitandi,† Nishant Kumar Rana,‡ Gunjan Sharma,‡ Biplob Koch,‡ Love Karan Rana,§ Maninder Singh Hundal,§,∥ and Daya Shankar Pandey*,† †

Department of Chemistry and ‡Department of Zoology, Faculty of Science, Banaras Hindu University, Varanasi 221005, Uttar Pradesh, India § Department of Chemistry, Guru Nanak Dev University, Amritsar 143005, Punjab, India S Supporting Information *

ABSTRACT: The Schiff base ligands benzylidene(4-tertbutylphenyl)amine 4-methyl ester (L1), (4-nitrobenzylidene)(4tert-butylphenyl)amine (L2), and (4-cyanobenzylidene)(4-tertbutylphenyl)amine (L3) and the new series of cyclometalated mononuclear piano-stool complexes [(η5-C5Me5)RhCl(L1)] (1), [(η5-C5Me5)RhCl(L2)] (2), [(η5-C5Me5)RhCl(L3)] (3), [(η5C5Me5)IrCl(L1)] (4), [(η5-C5Me5)IrCl(L2)] (5), and [(η5C5Me5)IrCl(L3)] (6) have been synthesized. The ligands L1−L3 and complexes 1−6 have been thoroughly characterized by satisfactory elemental analyses, spectral studies (ESI-MS, IR, 1H and 13C NMR, UV−vis), and structures of 1−3 authenticated by Xray single-crystal analyses. Efficient binding of 1−6 with calf thymus DNA (CT DNA) have been established by UV−vis and emission spectroscopic studies. Protein binding (bovine serum albumin, BSA) has been investigated by UV−vis, fluorescence, synchronous, and 3D fluorescence spectroscopy. Binding of the complexes with DNA through minor groove and hydrophobic interaction with proteins via sub domain IIA cavity has been substantiated by molecular docking studies. The complexes exhibited significant cytotoxicity against the human lung cancer cell line (A549), and 1 and 2 showed better activity than cisplatin. The cytotoxicity, morphological changes, and apoptosis have been assessed by MTT assay, Hoechst 33342/PI staining, cell cycle analysis by fluorescence-activated cell sorting (FACS), and reactive oxygen species (ROS) generation by DCFH-DA dye. The complexes 1−6 induce apoptosis in the order 2 > 1 > 4 > 3 > 5 > 6.

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INTRODUCTION Platinum based anticancer drugs, viz. cisplatin, oxaliplatin, carboplatin, etc., have proved to be indispensable in cancer chemotherapy and cover nearly 50% cancer therapeutic drugs worldwide.1 Unfortunately, these are associated with adverse side effects, severe toxicity, drug resistance, and lack of selectivity.2 To overcome the negative aspects of the platinum-based drugs, intense activity has been shown by various research groups to discover anticancer agents based on metals other than platinum that exhibit lower toxicity.3 In this context, ruthenium-based complexes have drawn special attention, due to their favorable kinetic aspects, variable oxidation states, and low toxicity.4 Simultaneously, anticancer properties of complexes derived from the neighboring group 9 members Rh(III) and Ir(III) have not been explored much, despite showing variable oxidation states (particularly MI, MIII, MIV; M = Rh, Ir), largely due to thier sluggish kinetic activity and lability toward ligand exchange.5 Recently it has been shown that half-sandwich Rh(III) and Ir(III) complexes display high flexibility toward promising anticancer activity via intercalating target DNA in the cancer cells.6 It has been © XXXX American Chemical Society

established that high activity can be achieved by alteration of ligands about the metal center and that minute variation in the structure by fine tuning functional groups can have a pronounced effect on biological activity.6 Introduction of an electron-rich pentamethylcyclopentadienyl group (Cp*) enhances the rate of ligand exchange many times over, and the labile −Cl attached to the metal center hydrolyzes rapidly.7 Further, incorporation of C,N-donor in place of N,N-donor ligands also improves the potency of the complexes toward tumor cells by enhancing electron density at the metal center and makes it labile for ligand exchange, stabilizes higher oxidation states, and enhances oxidizing ability.7 Highly oxidizing drugs generate reactive oxygen species (ROS), which cause death of the cancer cells oxidatively. Generation of the ROS for cellular signaling in various physiological processes is mostly governed by mitochondria;6 however, uncontrolled generation or reduced ability of the cells to eradicate ROS creates an oxidative stress, subsequently Received: June 10, 2015

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DOI: 10.1021/acs.organomet.5b00475 Organometallics XXXX, XXX, XXX−XXX

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Organometallics

(CDRI), Lucknow, India. IR and electronic absorption spectra were acquired on Varian 3300 FT-IR and Shimadzu UV-1601 spectrophotometers, respectively. Fluorescence spectra were recorded on a PerkinElmer LS 55 spectrofluorimeter equipped with a xenon lamp source and 10.0 mm quartz cell. The excitation and emission slit widths were set at 10.0 and 5.0 nm, and the temperature was maintained by recycling water using a Peltier system. 1H (300 MHz) and 13C (75.45 MHz) NMR spectra at room temperature were obtained on a JEOL AL300 FT-NMR spectrometer using tetramethylsilane (Si(CH3)4) as an internal reference. Electrospray ionization mass spectrometric (ESI-MS) measurements were made on a THERMO Finningan LCQ Advantage Max ion trap mass spectrometer. Samples (10 μL) were dissolved in dichloromethane/ acetonitrile (3/7, v/v) and introduced into the ESI source through a Finningan surveyor auto sampler. The mobile phase (MeCN/H2O, 90/10) flowed at a rate of 250 μL/min. The ion spray voltage was set at 5.3 kV and capillary voltage at 34 V. The MS scan was run up to 2.5 min, and spectral printouts were an average of over 10 scans. Synthesis of Benzylidene(4-tert-butylphenyl)amine 4-Methyl Ester (L1). Methyl 4-formylbenzoate (1.0 mmol 0.164 g) and 4-tertbutylaniline (1.0 mmol, 0.149 g) were dissolved in dry ethanol (25 mL) and stirred at room temperature for 1 h. A white precipitate started separating within 1 min; after complete precipitation the ensuing solid was filtered and washed twice with water and ethanol. It was dried in air and used without further purifications. Yield: 95% (0.295 g). Anal. Calcd for C19H21NO2: C, 77.26; H, 7.17; N, 4.74. Found: C, 77.32; H, 7.12; N, 4.69. IR (KBr pellets, cm−1): 568, 695, 769, 847, 1014, 1110, 1277, 1365, 1440, 1624, 1723, 2867, 2955. 1H NMR (CDCl3, δ ppm): 1.35 (s, 9H, C(CH3)3); 3.94 (s, 3H, OCH3); 7.21 (d, 2H, Ph); 7.43 (d, 2H, Ph), 7.96 (d, 2H, Ph), 8.13 (d, 2H, Ph), 8.52 (s, −CHN). 13C NMR (CDCl3, δ ppm): 30.25 (CH3), 33.43 {C(CH3)3}; 51.16 (OCH3); 119.48, 124.97,127.39, 128.80, 130.98, 139.05, 147.65, 148.63, (C6H6); 157.17, (−CHN); 165.47, (−C O). ESI-MS (calcd, found; m/z): 296.1651, 296.1652 [M + H]+. Synthesis of (4-Nitrobenzylidene)(4-tert-butylphenyl)amine (L2). This compound was prepared following the above procedure for L1 starting from 4-nitrobenzaldehyde (1.0 mmol, 0.151 g) and 4-tertbutylaniline (1.0 mmol, 0.149 g).14 The resulting orange precipitate was filtered, washed twice with water and ethanol, dried in air, and used without further purifications. Yield: 94% (0.265 g). Anal. Calcd for C17H18N2O2: C, 72.32; H, 6.43; N, 9.92. Found: C, 72.26; H, 6.51; N, 9.98. IR (KBr pellets, cm−1): 568, 690, 753, 845, 859, 1106, 1341, 1518, 1598, 2872, 2961. 1H NMR (CDCl3, δ ppm): 1.35 (s, 9H, C(CH3)3); 7.22 (d, 2H, Ph); 7.44 (d, 2H, Ph), 8.08 (d, 2H, Ph), 8.32 (d, 2H, Ph), 8.57 (s, −CHN). 13C NMR (CDCl3, δ ppm): 31.35 (CH3), 34.61 {C(CH3)3}; 120.71, 123.94, 120.64, 126.20, 129.25, 141.78, 148.17, 149.13, 150.47 (C6H6); 170.41 (−CHN). ESI-MS (calcd, found; m/z): 283.1447, 283.1441 [M + H]+. Synthesis of (4-Cyanobenzylidene)(4-tert-butylphenyl)amine (L3). This compound was prepared following the above procedure for L1 using 4-cyanobenzaldehyde (1.0 mmol, 0.131 g) and 4-tert-butylaniline (1.0 mmol, 0.149 g). The resulting yellow precipitate was dried in air and used without further purifications. Yield: 93.5% (0.245 g). Anal. Calcd for C18H18N2: C, 82.41; H, 6.92; N, 10.68. Found: C, 82.47; H, 6.85; N, 10.63. IR (KBr pellets, cm−1): 553, 572, 818, 844, 954, 1013, 1171, 1359, 1410, 1504, 1625, 2227, 2867, 2962. 1H NMR (CDCl3, δ ppm): 1.35 (s, 9H, C(CH3)3); 7.20 (d, 2H, Ph); 7.43 (d, 2H, Ph), 7.75 (d, 2H, Ph), 7.99 (d, 2H, phenyl), 8.51 (s, −CHN). 13C NMR (CDCl3, δ ppm): 31.33 (CH3), 34.56 {C(CH3)3}; 114.11, 118.46, 120.64, 126.15, 128.96, 132.44, 140.12, 148.22 (C6H6); 150.24 (−CHN); 156.97 (−CN). ESI-MS (calcd, found; m/z): [M + H]+ 263.1548, 263.1542. Synthesis of [(η5-C5Me5)RhCl(L1)] (1). Sodium acetate (0.25 mmol, 0.02 g) was added to a suspension of [{(η5-C5Me5)Rh(μCl)Cl}2] (0.10 mmol, 0.06 g) in CH2Cl2/MeOH (1/2, 30 mL) and stirred for 1 h at room temperature.15 Subsequently, L1 (0.25 mmol, ∼0.08 g) was added to the above reaction mixture and this mixture was stirred for an additional 12 h. The solvent was removed under reduced pressure and the residue washed twice with water and diethyl ether. Recrystallization from dichloromethane and petroleum ether

damaging the cell. Oxidative stress in cancer cells remains rather high, partially due to anomalous functioning of the mitochondria relative to normal cells.6a,7 The anticancer agents show selectivity toward cells with irregular ROS levels relative to normal cells and destroy tumor cells with irregular redox functioning.7 Further, serum albumin is the most essential protein associated with transportation, distribution, accumulation, and excretion of the drugs in tumor cells.8 In addition, recently it has been revealed that metal complexes not only bind to primary target DNA but also strongly interact with proteins.9 Bovine serum albumin (BSA) is used as a target protein to examine binding behavior and cytotoxicity of the metallodrugs.9 Plasma proteins such as BSA act as a carrier for these drugs by forming protein complexes both in vivo and in vitro.8c,9 Therefore, the development of anticancer agents targeting both DNA and proteins has been highly demanding.10 With the objective of developing anticancer agents based on cyclometalated half-sandwich Rh(III)/Ir(III) complexes, the six new compounds [(η5-C5Me5)RhCl(L1)] (1), [(η5-C5Me5)RhCl(L2)] (2), [(η5-C5Me5)RhCl(L3)] (3), [(η5-C5Me5)IrCl(L1)] (4), [(η5-C5Me5)IrCl(L2)] (5), and [(η5-C5Me5)IrCl(L3)] (6) have been synthesized via C−H bond activation in the Schiff base ligands. Notably, complexes used in this study have diverse substituents (R = ester, nitro, cyano) along with tert-butyl and pentamethylcyclopentadienyl groups, which may provide an electron-rich environment at the metal center. Further, Schiff base ligands L1−L3 have been chosen for the present study, considering their ease of synthesis, stability under physiological conditions, inherent biological activity, and utility against a wide range of microorganisms.11 Through this contribution we describe the synthesis and characterization of some Rh/Ir complexes, their effective binding with DNA and protein, and cytotoxicity against a human lung cancer cell line (A549) supported by MTT assay, Hoechst 33342/PI staining, cell cycle analysis by fluorescence activated cell sorting (FACS), and reactive oxygen species (ROS) generation.

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EXPERIMENTAL SECTION

Reagents. All of the synthetic manipulations have been performed under a nitrogen atmosphere, and solvents were dried and distilled prior to their use following standard literature procedures.12 Hydrated Rh(III)/Ir(III) chloride, pentamethylcyclopentadiene, 4-nitrobenzaldehyde, 4-cyanobenzaldehyde, methyl 4-formylbenzoate, tert-butylaniline, and agarose were procured from the Sigma-Aldrich Chemical Co. (India), and anhydrous sodium acetate was obtained from s. d. finechem. India, Pvt. Ltd. (Mumbai, India), and used as received without further purification. MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide dye) and trypsin were purchased from Hi-media (India), whereas Dulbecco’s modified eagle medium (DMEM), fetal bovine serum (FBS), and antibiotic solution (10000 units of penicillin/mL and 10 mg of streptomycin/mL) were purchased from Cellclone (Genetix Biotech Asia Pvt. Ltd.). DMSO (dimethyl sulfoxide) and RNase were obtained from GeNie, Merck (India), while Hoechst 33342 and 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA) were purchased from Sigma (USA). Propidium iodide (PI) was obtained from EMD Millipore-Calbiochem (USA0, whereas Triton X-100 and other analytical grade chemicals were purchased from Loba chemie Pvt. Ltd. (India). Lung cancer cell line A549 was obtained from the National Centre for Cell Science (NCCS) (Pune, India). The precursor complexes [{(η5-C5Me5)M(μ-Cl)Cl}2] (M = Rh, Ir) were prepared and purified by literature procedures.13 General Methods. Elemental analyses for C, H, and N were obtained on an Elementar Vario EL III Carlo Erba 1108 instrument by the microanalytical laboratory of the Sophisticated Analytical Instrumentation Facility (SAIF), Central Drug Research Institute B

DOI: 10.1021/acs.organomet.5b00475 Organometallics XXXX, XXX, XXX−XXX

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Organometallics (40−60 °C) afforded 1 as an orange crystalline solid. Yield: 84% (0.07 g). Anal. Calcd for C29H35ClNO2Rh: C, 61.33; H, 6.21; N, 2.47. Found: C, 61.37; H, 6.18; N, 2.43. IR (KBr pellets, cm−1): 767, 1106, 1241, 1271, 1434, 1716, 1725, 2953. 1H NMR (CDCl3, δ ppm): 1.35 (s, 9H, C(CH3)3), 1.48 (s, 15H, C5(CH3)5), 3.94 (s, 3H, OCH3); 7.42, 7.56, 7.72 (6H, Ph); 8.22 (s, 1H, Ph); 8.51 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 8.86 C5(CH3)5); 31.38 (CH3), 34.67 {C(CH3)3}; 52.04 (OCH3); 96.60 (C5(CH3)5), 120.60, 125.46, 128.88, 137.18, 149.09, 151.82 (C6H6); 167.44 (−CHN); 171.12 (−CO). ESI-MS (calcd, found; m/z): 585.1517, 585.1752, [1 + H2O], 532.1723, 532.1662 [1 − Cl]. Synthesis of [(η5-C5Me5)RhCl(L2)] (2). This complex was prepared following the above procedure for 1 using L2 (0.25 mmol, ∼0.07 g) in place of L1. After a routine workup complex 2 was obtained as a brown solid. Yield: 81% (0.06 g). Anal. Calcd for C27H32ClRhN2O2,: C, 50.34; H, 5.01; N, 4.35. Found: C, 50.26; H, 5.10; N, 4.42. IR (KBr pellets, cm−1): 642, 713, 841, 1017, 1201, 1344, 1441, 1512, 2917, 2956. 1H NMR (CDCl3, δ ppm): 1.36 (s, 9H, C(CH3)3), 1.46 (s, 15H, C5(CH3)5), 7.45, 7.55, 7.64, 7.90 (d, 6H, Ph); 8.27 (s, 1H, Ph); 8.66 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 9.06 C5(CH3)5); 31.37 (CH3), 34.73 {C(CH3)3}; 97.04 (C5(CH3)5), 118.18, 121.47, 125.99, 130.36, 147.95, 151.35, 152.35 (C6H6); 170.25 (−CHN). ESI-MS (calcd, found; m/z): 572.1313, 572.1476 [2 + H2O]. Synthesis of [(η5-C5Me5)RhCl(L3)] (3). This complex was prepared following the above procedure for 1 using L3 (0.25 mmol, ∼0.07 g) in place of L1. This complex was obtained as a red solid. Yield: 82% (0.06 g). Anal. Calcd for C27H32ClIrN2O2: C, 53.87; H, 5.17; N, 4.49. Found: C, 53.82; H, 5.22; N, 4.43. IR (KBr pellets, cm−1): 601, 826, 847, 1022, 1201, 1444, 1569, 2221, 2867, 2913, 2959. 1 H NMR (CDCl3, δ ppm): 1.36 (s, 9H, C(CH3)3), 1.43 (s, 15H, C5(CH3)5), 7.34, 7.44, 7.53, 7.58 (d, 6H, Ph); 8.07 (s, 1H, Ph); 8.22 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 8.88 C5(CH3)5); 31.32 (CH3), 34.71 {C(CH3)3}; 96.83 (C5(CH3)5), 113.57, 120.51, 126.11, 128.35, 139.38, 148.33, 151.44 (C6H6); 170.75 (−CHN); 184.36 (−CN). ESI-MS (calcd, found; m/z): 552.1415, 552.1611 [3 + H2O]. Synthesis of [(η5-C5Me5)IrCl(L1)] (4). This complex was prepared following the above procedure for 1 using L1 (0.25 mmol, 0.08 g) and [{(η5-C5Me5)Ir(μ-Cl)Cl}2] (0.10 mmol, ∼0.08 g). After a routine workup it was isolated as a brownish solid. Yield: 82% (0.065 g). Anal. Calcd for C29H35ClIrNO2: C, 52.99; H, 5.37; N, 2.13. Found: C, 53.08; H, 5.43; N, 2.20. IR (KBr pellets, cm−1): 419, 769, 1100, 1241, 1271, 1447, 1724, 2923, 2970. 1H NMR (CDCl3, δ ppm): 1.34 (s, 9H, C(CH3)3), 1.48 (s, 15H, C5(CH3)5), 3.93 (s, 3H, OCH3); 7.40, 7.42, 7.48, 7.66 (d, 6H, Ph); 8.38 (s, 1H, Ph); 8.51 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 8.69 C5(CH3)5); 31.34 (CH3), 34.63 {C(CH3)3}; 52.05 (OCH3); 89.62 (C5(CH3)5), 121.87, 123.32, 125.71, 128.58, 131.91, 136.03, 149.06, 148.92 (C6H6); 169.04 (−CHN); 174.07 (−CO). ESI-MS (calcd, found; m/z): 675.2091, 675.2285 [4+H2O]. Synthesis of [(η5-C5Me5)IrCl(L2)] (5). This complex was prepared following the above procedure for 1 using L2 (0.25 mmol, ∼0.07 g) and the iridium complex [{(η5-C5Me5)Ir(μ-Cl)Cl}2] (0.10 mmol, 0.08 g). It was isolated as a yellow-brown solid. Yield: 80% (0.06 g). Anal. Calcd for C27H32ClIrN2O2: C, 50.31; H, 5.04; N, 4.41. Found: C, 50.39; H, 5.08; N, 4.45. IR (KBr pellets, cm−1): 716, 841, 1028, 1083, 1106, 1202, 1343, 1442, 1512, 2913, 2961. 1H NMR (CDCl3, δ ppm): 1.36 (s, 9H, C(CH3)3), 1.50 (s, 15H, C5(CH3)5), 7.44, 7.49, 7.73, 7.85 (d, 6H, Ph); 8.44 (s, 1H, Ph); 8.62 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 8.69 C5(CH3)5); 31.33 (CH3), 34.70 {C(CH3)3}; 90.18 (C5(CH3)5), 117.45, 121.80, 125.86, 129.12, 148.85, 151.54, 152.35 (C6H6); 173.16 (−CHN). ESI-MS (calcd, found; m/z): 662.1887, 662.2093 [5 + H2O]. Synthesis of [(η5-C5Me5)IrCl(L3)] (6). The complex 6 was prepared using L3 (0.25 mmol, 0.07 g) and the iridium complex [{(η5-C5Me5)Ir(μ-Cl)Cl}2] (0.10 mmol, 0.08 g) following the above procedure for 1. It was isolated as a red solid. Yield: 78% (0.06 g). Anal. Calcd for C27H32Cl-IrN2O2: C, 53.87; H, 5.17; N, 4.49. Found: C, 53.81; H, 5.22; N, 4.54. IR (KBr pellets, cm−1): 606, 822, 847,

1027, 1202, 1443, 1564, 1746, 2220, 2854, 2923, 2959. 1H NMR (CDCl3, δ ppm): 1.35 (s, 9H, C(CH3)3), 1.47 (s, 15H, C5(CH3)5), 7.30, 7.43, 7.49, 7.64 (d, 6H, Ph); 8.05 (s, 1H, Ph); 8.39 (s, 1H, −CHN). 13C NMR (CDCl3, δ ppm): 8.69 C5(CH3)5); 31.34 (CH3), 34.70 {C(CH3)3}; 54.93 (OCH3); 90.01 (C5(CH3)5), 121.81, 125.21, 128.68, 138.30, 151.41, 156.99 (C6H6); 173.73 (−CHN); 182.16 (−CN). ESI-MS (calcd, found; m/z): 642.1989, 642.1353 [6 + H2O]. X-ray Structure Determination. Crystals suitable for X-ray single crystal analyses for 1−3 were obtained by slow diffusion of methanol over solutions of the respective complexes in dichloromethane. X-ray data for 1−3 were collected on a Bruker Kappa Apex-II diffractometer at room temperature with Mo Kα radiation (λ = 0.71073 Å) at the Department of Chemistry, Center for Advanced Studies, Guru Nanak Dev University, Amritsar, India. Structures were solved by direct methods (SHELXS 97) and refined by full-matrix least squares on F2 (SHELX-97).16 All non-H atoms were treated anisotropically. The H atoms attached to carbon were included as fixed contributions and geometrically calculated and refined using the SHELX riding model. The computer program PLATON was used for analyzing the interaction and stacking distances.17 Additional crystallographic information is available in the Supporting Information. Partition Coefficient Determination. The lipophilicity of complexes 1−6 was determined by the “shake flask” method between octanol/water phase partitions.18 Octanol-saturated water (OSW) and water-saturated octanol (WSO) were prepared using analytical grade octanol (Merck) and doubly distilled water. Complexes 1−6 (1 mg/ mL; ethanol/water 1/6) were diluted to 2, 4, 6, 8, and 10 μg/mL in water; alternatively these (1 mg/mL) were diluted to 2, 4, 6, 8, and 10 μg/mL in octanol, respectively. Appropriate amounts of the complexes (4 mg/mL) were shaken for 24 h at room temperature in equal volume (50/50). After the attainment of equilibrium, the organic and aqueous phases were separated and centrifuged. Finally, the concentration of the drug in each phase was determined by UV/ visible spectroscopy. The sample solution concentration was used to calculate log P. Partition coefficients for 1−6 were calculated using the equation log P = log[(1−6)oct/(1−6)aq].19 Molecular Docking. Molecular docking studies on 1−6 have been made using HEX 6.1 software and Q-site finder, which is an interactive molecular graphics program for interaction and docking calculations, in order to find out possible binding sites for biomolecules. DFT calculations were carried out using GAUSSIAN 09 by B3LYP methods. The geometries of 1−6 were optimized with the standard 6-31G** basis set for C, H, N, O, and Cl, while LANL2DZ was used for Rh and Ir with effective core pseudopotential for the metal.20 The coordinates for metal complexes were taken from their optimized structures as a .mol file and transformed to PDB format using CHIMERA 1.5.1 software. The crystal structure of B-DNA (PDB ID: 1BNA) was taken from the Protein Data Bank (http://www.rcsb.org./ pdb). Visualization of the docked molecules has been made with Discovery Studio 3.5 software. By default, the parameters used for docking calculations were correlation type shape only, FFT mode at the 3D level, and grid dimension 6 with receptor range 180 and ligand range 180 with twist range 360 and distance range 40. Preparation of the Drugs. The complexes were dissolved in DMSO (c = 100 mM) and then further diluted in DMEM to 10 mM. The maximum concentration of DMSO even at the highest concentration of the drugs was less than 0.1% v/v. Cytotoxicity and Proliferation Assay by MTT Assay. The MTT assay is a quantitative, sensitive, and reliable colorimetric technique generally used for assessing viability and proliferation of the cells. In this assay mitochondrial dehydrogenase enzymes of the live cells convert the yellow water-soluble substrate 3-(4,5-dimethylthiazol2-yl)-2,5 diphenyltetrazolium bromide (MTT) into a purple formazan crystal which is insoluble in water or the medium.21 Lung cancer cells (A549) with a density of 10000 cells per well were seeded and treated with different concentrations of 1−6 and cisplatin for positive control in 96-well tissue culture plates and incubated for 24 h. The working concentration of MTT (5 mg/mL) was prepared in PBS, and 10 μL was added into each well; after incubation for 2 h, 100 μL of DMSO C

DOI: 10.1021/acs.organomet.5b00475 Organometallics XXXX, XXX, XXX−XXX

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Organometallics Scheme 1. Synthesis of the Ligands L1−L3 and Complexes 1−6

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was used to dissolve formazan crystals. This solution was incubated for 30 min, and the absorbance was measured spectrophotometrically in an ELISA plate reader at a wavelength of 570 nm. All experiments were performed in three replicates and IC50 values for each compound estimated. Morphological Analysis of Nucleus with Hoechst 33342/PI Staining. Hoechst 33342 and propidium iodide (PI) double staining provides a fast and suitable assay for nuclear morphology of the apoptotic cells. Hoechst 33342 binds to the minor groove of doublestranded DNA preferably at the AT-rich region, while PI binds to DNA by intercalating between bases without any sequence preference. PI is usually used to identify dead cells. It is cell membrane impermeable and gets excluded from the viable cells, while Hoechst 33342 dye is permeable and binds to DNA in live or fixed cells. For this experiment 10000 A549 cells were seeded in a 96-well culture plate in DMEM and incubated for 24 h. Cells were treated at two different concentrations, including the IC50 value for the drugs (8 and 10 μM, 1; 6 and 8 μM, 2; 20 and 25 μM, 4) and incubated for 24 h in a CO2 incubator. Cells were stained with Hoechst and PI followed by images acquired with a fluorescent microscope at particular wavelengths (Hoechest 33342, λex 378 nm and λem 457 nm; PI, λex 535 nm and λem 617 nm). Cell Cycle Analysis by Fluorescence-Activated Cell Sorting (FACS). Fluorescence activated cell sorting (FACS) is a specialized type of flow cytometry that provides a method for sorting of the cells from their heterogeneous mixture by suspending them in a stream of fluid and passing through an electronic detector.22 It is based on the fact that each cell has specific light -scattering and fluorescent properties. Cell cycle analyses by FACS with PI staining have been performed to check the cell cycle arrest or alteration in cell cycle phases. In this direction 0.5 × 106 cells were seeded in a 6-well cell culture plate and treated with three different concentrations of the complexes close to their IC50 value. Cells were incubated for 24 h in a CO2 incubator. After incubation these were washed and suspended in PBS (500 μL) and thereafter fixed with 70% ethanol and left for 24 h at −20 °C. After incubation the fixed cells were washed with ice-cold PBS and a single-cell suspension was prepared for each sample. Staining of the cells was carried out with PI-RNase solution (1 mg/mL PI, 0.1% v/v Triton X-100, and 10 mg/mL RNase), and the cells were incubated at 37 °C for 30 min and analyzed in FACScan using CellQuest software (Becton Dickinson). ROS Detection. Reactive oxygen species include various molecules (such as H2O2, NO, O2−, OH−, ONOO−, and HOCl) which oxidize protein and lipids and damage DNA/RNA. 2′,7′-Dichlorofluorescine diacetate (DCFH-DA) is an oxidation-sensitive fluorescence probe, and by intracellular esterase hydrolysis it forms nonfluorescent DCFH. In the presence of ROS, DCF-H becomes oxidized and forms highly fluorescent DCF.23 A portion of 1 × 106 cells/mL was seeded in 6-well culture plates, treated with two different concentrations of the complexes, and incubated at 37 °C with 5% CO2 for 24 h. After treatment with these drugs the cells were stained with 10 μM DCFHDA solution (stock concentration 10 mM in DMSO) and incubated at 37 °C for 30 min and images were taken with a fluorescence microscope after incubation.

RESULTS AND DISCUSSION Synthesis and Characterization. The chloro-bridged dimeric complexes [{(η5-C5Me5)M(μ-Cl)Cl}2] (M = Rh, Ir) reacted with the Schiff base ligands benzylidene(4-tertbutylphenyl)amine 4-methyl ester (L1), (4-nitrobenzylidene)(4-tert-butylphenyl)amine (L2), and (4-cyanobenzylidene)(4tert-butylphenyl)amine (L3) in a 1/2.5 molar ratio in CH2Cl2/ MeOH (1/2, 30 mL) in the presence of sodium acetate with stirring at room temperature to afford complexes 1−6 in reasonably good yield (75−85%). A simple synthetic strategy adopted for the preparation of the ligands and complexes is shown in Scheme 1. The complexes under investigation are nonhygroscopic, airstable crystalline solids that are highly soluble in common organic solvents such as dichloromethane, chloroform, acetone, dimethyl sulfoxide, and acetonitrile, partially soluble in methanol, and ethanol, and insoluble in diethyl ether, benzene, hexane, and petroleum ether. These complexes have been thoroughly characterized by satisfactory elemental analyses, spectroscopic studies (ESI-MS, IR, 1H, 13C NMR, UV−vis), and the structures of 1−3 have been authenticated by X-ray single-crystal analyses. The ν(−CO), ν(−NO2), and ν(−CN) bands in the IR spectra of ligands L1−L3 vibrated at 1723, 1341, and 2226 cm−1, while these appeared at 1725 (1), 1344 (2), 2221 (3), 1724 (4), 1343 (5), and 2221 (6) cm−1 in the respective complexes (Figures S1−S3 in the Supporting Information). The presence of the bands due to ν(−CO), ν(−NO2), and ν(−CN) in the IR spectra of the complexes strongly suggested coordination of the ligands with the metal center, which has further been supported by structural studies on 1−3 (vide infra). NMR Spectral Studies. 1H and 13C NMR spectra of the ligands and complexes 1−6 have been acquired in CDCl3, and the resulting data are gathered in the Experimental Section along with other characterization data (Figures S4−S12 in the Supporting Information). The aldimine protons of L1 upon coordination with the metal center exhibited an insignificant shift (8.52 to 8.51 ppm) in the complexes 1 and 4, while L2 aldimine protons displayed a moderate downfield shift of 0.05 ppm (8.57 to 8.66 ppm, 2) and 0.09 ppm (8.57 to 8.62 ppm, 5). The shifts in 1, 3, 4, and 6 are inappreciable, as the ester and cyano groups are poorly electron withdrawing in comparison to the −NO2 group. On the other hand, L3 aldimine protons exhibited a significant upfield shift of 0.12 (8.51 to 8.22 ppm, 3) and 0.29 ppm (8.51 to 8.39 ppm, 6). The downfield shifts in 2 and 5 may be attributed to the greater electron-withdrawing effect of the −NO2 group. Due to M−C bond formation via C−H activation, the number of integrated protons became lower by 1 relative to the respective ligands in the complexes. D

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2.39 (2), 2.39 Å (3)) are normal and comparable to those in other closely related systems. The bond angles C−Rh−Cl and N−Rh−Cl are close to 90° (C11−Rh1−Cl1 88.32° and N1− Rh1−Cl1 86.45°, 1; C3−Rh1−Cl1 86.55° and N2−Rh1−Cl1 87.49°, 2; C1−Rh1−Cl1 86.68° and N1−Rh1−Cl1 86.81°, 3) (Table 2). The angles N1−Rh1−Cg, Cl1−Rh1−Cg, C11− Rh1−Cg are 122.69, 131.76, and 133.25° for 1, 123.40, 133.29, and 131.12° for 2, and 123.39, 133.47, and 132.11° for 3, respectively, suggesting a piano-stool geometry about the metal center (Table 3). The Cp* rings in 1−3 are symmetrically bonded to the metal center with average Rh−C distances of 2.21 Å (range 2.15−2.26 Å). The carbon atoms in the Cp* ring are planar, and rhodium is displaced from the centroid of the ring by 1.84 Å, which is normal and comparable to values in other closely related rhodium complexes.24,25 Electronic Absorption Spectroscopy. UV−vis absorption spectra (Figure 2) of 1−6 have been acquired in EtOH/ PBS (c = 10 μM; 1/1 v/v; pH ∼7.3). The low-energy weak bands at ∼345 nm in the spectra of the complexes have been assigned to mixing of the n−π* metal to ligand (MLCT) or ligand to metal charge transfer (LMCT), while strong highenergy bands at ∼240 nm have been attributed to spin-allowed π−π* transitions.26 UV−Vis Titration Studies. Electronic absorption titration studies offer an effective method to follow binding of the complexes with DNA.27 Intercalation or interaction of the metal complexes with base pairs of DNA is usually followed by a hypochromic shift with a small red/blue shift.28 On the other hand, nonintercalative, electrostatic interactions or damage of the DNA double helix results in a hyperchromic shift.29 UV−vis titration plots for 1−6 vs CT-DNA are depicted in Figure 3 (Figures S16 and S17 in the Supporting Information). Addition of CT DNA (0−20 μM) to solutions of 1−6 resulted in hyperchromism with a small red shift. Shifting in the position of absorption bands and changes in molar extinction coefficients are summarized in Table S1 in the Supporting Information. Compounds 4−6 showed the emergence of a new band (256 (4), 249 (5), 252 (6) nm) assignable to covalent interaction of the iridium complexes with DNA. From the UV−vis titration studies it has been inferred that complexes do interact with CT DNA via electrostatic interaction or covalent bond formation. From available data it has been concluded that rhodium complexes 1−3 interact with CT DNA in a dissimilar manner relative to the iridium complexes 4−6. To compare the binding affinity of the complexes, equilibrium binding constants (Kb) and binding site sizes (s, per base pair) have been calculated using the equation of Bard and co-workers based on the McGhee−von Hippel (MvH) model.30 Calculated values of intrinsic equilibrium binding constant (Kb) and binding site size (s) (7.3 × 105, 0.12, 1; 7.4 × 105, 0.11, 2; 5.8 × 105, 0.13, 3; 7.2 × 105, 0.15, 4; 4.2 × 105, 0.14, 5; 3.9 × 105, 0.15, 6) are comparable to those in earlier reports.31 Absorption titration studies clearly suggested that, among the complexes under investigation, 1, 2, and 4 bind with CT DNA rather strongly relative to 3, 5, and 6 and their binding affinity lies in the order 2 > 1 > 4 > 3 > 5 > 6. This may be ascribed to the presence of different functional groups in the complexes and the lower value of s attested groove and/or surface binding. Ethidium Bromide (EtBr) Displacement Studies. UV− vis titration studies clearly indicated effective binding of 1−6 with CT DNA. To get a better understanding about the nature and mechanism of binding, ethidium bromide displacement studies have been performed. EtBr is an intercalating agent

The aromatic proton adjacent to the M−C bond in these complexes (1−6) exhibited a downfield shift due to the electropositive metal center.24,25 The position of −COOMe protons of L1 exhibited an insignificant change (3.94 ppm for 1 to 3.93 ppm for 4) in 1 and 4. The Cp* protons in 1−6 displayed a small upfield shift and resonated at almost the same position (δ ∼1.48 (1), 1.46 (2), 1.43 (3), 1.48 (4), 1.50 (5), and 1.47 ppm (6)) with respect to the precursor complexes, indicating a rather small change in electronic environment about the metal center in these complexes. The tert-butyl protons in all three ligands and the six complexes resonated at their usual position (1.34−1.36 ppm).25c 13C NMR spectral data of the ligands and complexes 1−6 further supported their formation and proposed formulations. Crystal Structure. Molecular structures of 1−3 have been determined by single-crystal X-ray diffraction analyses. Details about data collection, solution, and refinement are given in Table 1, and selected geometrical parameters are gathered in Table 1. Crystal Data and Structure Refinement Parameters for 1−3 param

1

2

3

empirical formula formula wt cryst syst space group a (Å) b (Å) c (Å) α (deg) β (deg) γ (deg) V (Å3) color, habit

C29H35ClNO2Rh

C27H32ClN2O2Rh

C28H32ClN2Rh

567.94 triclinic P1̅ 8.577(12) 11.815(16) 14.119(2) 98.924(6) 99.556(6) 101.077(6) 1358.5(3) brown, rectangular

534.92 triclinic P1̅ 8.365(2) 11.763(3) 13.777(4) 100.123(10) 99.718(11) 98.578(10) 1292.8(6) brown, prism

Z dcalcd (g/cm3) temp (K) wavelength (Å) μ (mm−1) GOF on F2 R indices (all data) R1 wR2 final R indices (I > 2σ(I)) R1 wR2

2 1.388 296(2) 0.71073

554.91 triclinic P1̅ 8.337(2) 12.012(4) 13.615(4) 100.444(12) 98.346(12) 98.460(12) 1305.4(7) colorless, rectangular 2 1.412 296(2) 0.71073

2 1.374 296(2) 0.71073

0.752 1.056

0.782 0.836

0.781 1.014

0.0405 0.0916

0.0329 0.0876

0.0359 0.0359

0.0331 0.0860

0.0289 0.0825

0.0306 0.0746

Tables 2 and 3, respectively. Pertinent views of 1−3 along with a partial atomic numbering scheme are depicted in Figure 1. All of these crystallize in the triclinic system with the P1̅ space group. The metal center in these complexes adopted a typical “piano-stool” geometry with coordination sites about the rhodium center occupied by one nitrogen from the aldimine core, a carbon from the adjacent phenyl ring of the respective ligand, a chloro group, and the pentamethylcyclopentadienyl ring (Cp*) in an η5 manner. The Rh−N and Rh−Cl bond distances in the complexes (Rh1−N1 2.09 (1), 2.09 (2), 2.09 Å (3); Rh1−Cl1 2.40 (1), E

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Organometallics Table 2. Selected Bond Lengths (Å) of 1−3 1 C11−Rh1 C17−N1 C18−N1 N1−Rh1 Cl1−Rh1 C28−O2 C28−O1 C29−O1 Rh−Cg Rh−C (av)

2 2.037(2) 1.295(4) 1.423(3) 2.089(2) 2.397(8) 1.205(5) 1.322(5) 1.446(5) 1.84 2.21

C3−Rh1 C7−N2 C8−N2 N2−Rh1 N1−O2 N1−O1 Cl1−Rh1 Rh−Cg Rh−C (av)

3 2.019(2) 1.297(3) 1.430(3) 2.094(19) 1.101(7) 1.197(6) 2.393(9) 1.84 2.23

C1−Rh1 C8−N1 C9−N1 C4−N2 N1−Rh1 Cl1−Rh1 Rh−Cg Rh−C (av)

2.018(3) 1.299(4) 1.430(3) 1.121(6) 2.091(2) 2.390(7) 1.84 2.25

Table 3. Selected Bond Angles (deg) for 1−3 1 C16−C11−Rh1 C17−N1−Rh1 C11−Rh1−N1 C11−Rh1−Cl1 N1−Rh1−Cl1

2 114.51(19) 115.58(18) 78.41(10) 86.45(7) 88.32(6)

C4−C3−Rh1 C3−Rh1−N2 C7−N2−Rh1 C3−Rh1−Cl1 N2−Rh1−Cl1

3 114.93(17) 78.75(9) 115.31(15) 86.65(7) 87.49(6)

C7−C1−Rh1 C8−N1−Rh1 C1−Rh1−N1 C1−Rh1−Cl1 N1−Rh1−Cl1

114.86(19) 115.39(18) 78.66(10) 85.68(8) 86.81(6)

Figure 2. Electronic absorption spectra of 1−6 in EtOH/PBS (c = 10 μM; 1/1 v/v; pH ∼7.3).

The fluorescence spectral pattern for EtBr−DNA and quenching in the presence of 1−6 is shown in Figure 4 (Figure S18 in the Supporting Information). Addition of a 50 μM solution of the aforementioned complexes to a saturated solution of EtBr−DNA causes quenching relative to the initial intensity (90% (1), 86% (2), 76% (3), 97% (4), 87% (5), and 88% (6)). From the above results it has been concluded that EtBr is being displaced quite efficiently from the binding sites of CT DNA by the complexes under study.33 Quenching parameters have been analyzed by the Stern− Volmer equation (Figure S19 in the Supporting Information)

Figure 1. ORTEP views of 1 (a), 2 (b), and 3 (c) at the 30% thermal ellipsoid probability level (H atoms omitted for clarity).

I0/I = Kq[Q] + 1

frequently used as a fluorescent tag for staining nucleic acids and other biomolecules. It itself is weakly fluorescent, but after binding with DNA it fluoresces rather strongly (almost 20-fold enhancement) with an orange color.32 Therefore, if the compounds of interest displace EtBr from the EtBr−DNA complex, significant quenching occurs due to accumulation of free EtBr in the system and lowers the number of binding sites available for EtBr−DNA binding.

(1)

where I0 is the emission intensity in absence of quencher, I is the emission intensity in the presence of quencher, Kq is the quenching constant, and [Q] is the concentration of the complex (Q = quencher). Kq values have been derived from the slope of I0/I vs [Q] plots (2.88 × 104 (1), 2.90 × 104 (2), 2.7 × 104 (3), 2.85 × 104 (4), 2.5 × 104 (5), and 2.2 × 104 (6)). Further, apparent DNA binding constants (Kapp) have been calculated using the equation F

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Figure 3. UV−vis titration spectra of 1 (a) and 4 (b) in EtOH/PBS (1/1) with an increasing concentration of CT DNA in a solution of the complex (0−20 μM) at room temperature. The arrows show absorbance changes upon increasing CT DNA concentration.

Figure 4. Fluorescence spectra of EtBr bound to CT DNA in the absence and presence of (a) 1 and (b) 4 (EtBr, 10 μM; DNA, 10 μM; [1] and [4], 0−50 μM). The arrows show changes in emission intensity upon increasing the amounts of 1 and 4.

Figure 5. Emission spectra of BSA (0.5 μM; λex 280 nm; λem 343 nm) in the presence of increasing amounts of the complexes (a) 1 and (b) 4.

KEB[EtBr] = K app[complex]

conclude that all of the complexes may bind DNA via intercalation. Protein Binding Studies. Due to its remarkable binding properties and highest abundance in blood plasma, serum albumin plays a crucial role in the drug delivery system.34 Bovine serum albumin (BSA) offers a cost-effective model for protein binding because of its abundance, ease of purification, stability, medical significance, ligand binding properties, and resemblance to human serum albumin (HSA).35 The highly fluorescent nature of BSA is related to the presence of tryptophan and tyrosine residues. Quenching and shifts in the

(2)

where [complex] is the value at 50% lowering in the fluorescence intensity for EtBr, KEtBr (1.0 × 107 M−1) is the DNA binding constant for EtBr, and [EtBr] is the concentration of EtBr (10 μM). Kapp values have been found to be 2.46 × 105 (1), 2.53 × 105 (2), 2.16 × 105 (3), 2.22 × 105 (4), 2.13 × 105 (5), and 1.93 × 105 (6) M−1. From the available data it is clear that all the complexes effectively replace EtBr from the EtBr−DNA complex, which is in accord with the results obtained from electronic absorption studies, and we G

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Further, to get an idea about the quenching sequence, emission quenching data have been analyzed using the Stern− Volmer and Scatchard equations. The quenching constants (Kq) have been calculated using I0/I vs [Q] plots (Figure S21 in the Supporting Information). In addition, equilibrium binding constants have been derived using the Scatchard equation:

emission spectra depend on binding with the tryptophan residue followed by conformational changes, subunit associations, substrate binding, or denaturation and provide an idea about the reaction dynamics and protein folding. To understand the nature of binding with complexes, quenching of the emission intensity of BSA has been investigated in the presence of the complexes under study. In this direction, fluorescence titration studies have been performed at room temperature using BSA (0.5 μM) and with varying concentrations of 1−6 (0−50 μM) in the range 290−500 nm (λex 280 nm; Figure 5 and Figure S20 in the Supporting Information). It has been observed that the fluorescence intensity of BSA at ∼343 nm was quenched with a small red shift (Δλ (nm), ΔΦ (%): 12, 89.18, 1; 10, 86.40, 2; 12, 85.50, 3; 12, 82.50, 4; 10, 72.46, 5; 10, 70.12, 6). Observed red shifts may be due to the presence of active sites of the protein in a hydrophobic environment. From these observations we conclude that some interaction is definitely taking place between the complexes and BSA.35 The quenching mechanism may follow a static or dynamic mode. Static quenching typically arises from complexation between the quencher and fluorophore in the ground state, while dynamic quenching results from interaction between the fluorophore and quencher in the short-lived excited state.34,35 Temperature-dependent emission quenching experiments permit us to make a distinction between the dynamic and static quenching. In dynamic quenching a higher temperature results in rapid diffusion and therefore the quenching rate constant increases with an increase in temperature. On the other hand, in static quenching an increase in temperature lowers the stability of the complex and the value of quenching constant (Figure S21 and Table S2 in the Supporting Information). In the present study it has been observed that KSV decreases with an increase in temperature, which indicated that BSA fluorescence quenching in the presence of 1−6 occurs via a static mechanism.34,35 One can have an idea about the type of quenching from the UV−vis absorption spectrum of BSA (fluorophore). In a dynamic quenching significant changes in absorption spectra of the fluorophore are not expected, in contrast, in static quenching perturbations occur.34,35 From Figure 6 it is clear that the absorption intensity increases with a small blue shift (∼7 nm) in the presence of 1−6, indicating a static interaction between BSA and the complexes.

log[(I0 − I )/I ]=log Kbin + n log [Q]

(3)

where Kbin is the binding constant of the compound with BSA and n is the number of binding sites. From the plot of log[(I0 − I)/I vs log [Q], the number of binding sites n and binding constant (Kbin) have been calculated. The resulting Kq, Kbin, and n values are gathered in Table S2 in the Supporting Information. The value of n for all compounds came out to be close to 1 and strongly suggested the existence of a single binding site in BSA for 1−6. Further, Kq and Kbin values for these compounds suggested that 1, 2, and 4 interact with BSA rather strongly relative to the other complexes under investigation. Conformational Investigation. To gain deep insight into the conformational changes in BSA in the presence of 1−6, synchronous fluorescence experiments have been performed under analogous conditions. This method provides detailed information about the molecular microenvironment, particularly in the vicinity of the fluorophore in BSA.36 The use of Δλ = 15 nm gives a synchronous emission spectrum characteristic of tyrosine, while the use of Δλ = 60 nm highlights tryptophan residues.37 Emission maxima of the tryptophan and tyrosine residues in proteins are related to the polarity of their surroundings. The synchronous fluorescence spectra (Δλ, 15 nm) for BSA with increasing concentrations of 1−6 displayed a decrease in fluorescence intensity at 288 nm (65.59% (1), 68.52% (2), 66.18% (3), 68.60% (4), 55.56% (5), and 50.15% (6)) with an inappreciable shift of 1 or 2 nm (Figure 7 and Figure S22 in the Supporting Information). Simultaneously a new band characteristic for the complexes emerged at 324 nm. However, synchronous fluorescence spectra of BSA at Δλ = 60 nm exhibited a decrease in fluorescence intensity at 280 nm (69.18% (1), 69.23% (2), 58.40% (3), 71.99% (4), 56.60% (5), and 40.16% (6)) with significant red shifts of 4, 7, 5, 6, 8, and 11 nm, respectively. Isoemissive points at ∼290−330 nm for 1−6 indicated the existence of an equilibrium between the complexes and BSA. The spectra displayed a decrease in fluorescence intensity with a small red shift for both tryptophan and tyrosine residues with increasing concentrations of the complexes, as cyclometalated rhodium and iridium complexes are nonemissive. The results clearly suggested that 1−6 effectively interact with BSA and affect the conformatiosn of the tyrosine and tryptophan microregions.37 In addition, the results suggested an increase in hydrophobicity around both tyrosine and tryptophan residues along with a decrease in polarity. The synchronous measurements indicated that 1−6 effectively bind with BSA and can be utilized for anticancer activity. Three-Dimensional (3D) Fluorescence Spectroscopy. Excitation−emission matrix spectroscopy (EEMS) or threedimensional fluorescence spectroscopy (3D) provides overall information about the emission characteristics of the fluorophores by changing excitation and emission wavelengths simultaneously.38 It is well-known that 3D emission spectral analysis presents conformational and microenvironmental changes for BSA. The 3D fluorescence spectra of BSA in the

Figure 6. UV−vis spectra of BSA (10 μM) in the presence of the complexes 1−6 (5 μM). H

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Figure 7. Synchronous spectra of BSA (0.5 μM) in the presence of increasing amounts of 1 and 4 (0−50 μM) with a wavelength difference of Δλ = 15 nm for (a) 1 and (b) 4 and Δλ = 60 nm for (c) 1 and (d) 4. The arrows show changes in emission intensity accompanied by red shifts upon increasing concentrations of the complexes.

Figure 8. 3D fluorescence spectra of (a) BSA and (b) BSA + 4. Conditions: c(BSA) = 2 × 10−6 mol L−1, c(2) = 2 × 10−6 mol L−1.

absence and presence of 1−6 are depicted in Figure 8 (Figure S23 in the Supporting Information), and the resulting data are gathered in Table S3 in the Supporting Information. From Figure 8 it can be seen that the 3D fluorescence spectrum of BSA exhibits four characteristic peaks: the peak on the extreme left is assigned as the first-order Rayleigh scattering peak 1, whose emission wavelength equals the excitation wavelength (λex = λem) ,whereas that on the extreme right is assigned to the second-order Rayleigh scattering peak 4, for which λem equals 2λex. In addition, two more peaks, namely peaks 2 and 3, is also displayed. Peak 2 mainly reflects the spectral behavior of the tyrosine and tryptophan residues.39 The results demonstrated that peak 2 exhibited a decrease in the emission intensity by 70%, 71%, 68%, 82%, 72%, and 69%, respectively, in the

presence of 1−6 with insignificant changes in emission wavelength. This suggested that the polarity of both residues decreases and more amino acid residues of BSA are buried in the hydrophobic pocket (Table S3). Furthermore, peak 3, assigned to the polypeptide backbone, showed a decrease in fluorescence intensity by 77% (1), 73% (2), 76% (3), 85% (4), 69% (5), and 58% (6), suggesting extension of the polypeptides of BSA to a greater extent and conformational changes in BSA. From the above data it can be concluded that the interaction of 1−6 with BSA leads to changes in tyrosine and tryptophan residues as well as in microenvironment polarities for the amino acid residues and conformation of BSA. I

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Figure 9. Molecular docked model of complex 2 with DNA (PDB ID: 1BNA).

complexes.41 Hydrolyses of compounds 1 and 4 in 5% MeODd4/95% D2O (v/v) was monitored by 1H NMR spectroscopy at 278−293 K (Figure S25 in the Supporting Information). The presence of methanol affirmed the solubility of complexes 1 and 4 (c = 1 mM), which underwent rapid hydrolysis in MeOD-d4/95% D2O solution within 1 min. Integration of the 1 H NMR peaks for complexes was enhanced with the addition of NaCl solution (4 equiv) in deuterated water instantaneously, suggesting that the equilibrium is rapidly shifting toward chloride from the aqua complex and the rate of aquation is very fast. Again, DNA is a possible target site for transition-metal anticancer complexes;42 the binding of 9-ethylguanine (9-EtG) to 1 and 4 was studied by 1H NMR titration. Interesting results were observed upon addition of 9-EtG (1.0 equiv) to the aforementioned stock solution of 1 and 4 (Figure S26 in the Supporting Information). Complex 1 does not show any remarkable shift in the spectrum except for the clear separation of 1H NMR peaks, while 4 showed a substantial shift of the high-field proton from 11.39 to 10.04 ppm along with the peak separation. The result clearly suggested that 1 does not bind with 9-EtG and only a weak interaction is occurring between them, whereas 4 certainly binds with the N atom of 9-EtG, attesting to different modes of action for these two complexes.43 1 interacts electrostatically with the minor groove of DNA, whereas 4 binds covalently. Partition Coefficient Determination. The lipophilicity of a biologically active compound plays an central role in absorption, distribution, metabolism, and elimination processes.44 log P is usually used for classification of the lipophilicity in n-octanol/water owing to similar environments in biological membranes.15 Partition coefficients, P, have been estimated to interpret the permeability of 1−6 through the cell membrane.46,47 The log P values determined (Table S3 in the Supporting Information) are comparable to those in earlier reports.48 The partition coefficient values (log P) of the complexes suggested that lipophilicity can be arranged in the order 2 > 1 > 4 > 3 > 5 > 6. These results are consistent with MTT assay; the cytotoxicity of the complexes increases with increasing lipophilicity.

Reversibility of BSA−Metallodrug Interaction. BSA− drug interactions are crucial in determining the utility of drugs. If the interaction is irreversible, then the drugs will not be released at the cellular target and the drug will have limited utility.35e To affirm the reversibility of interactions between the drugs and BSA, fluorescence experiments have been performed. The reversibility is achieved in a low pH environment as in the tumor tissues or by chelators present in the cytosol, which may displace amino acids from BSA−metallodrug complexes. It was observed that the emission intensity of BSA was quenched upon addition of 50 μL (1 mM) of 1 and 4. Addition of 200 μL (1 mM) of citric acid after saturation showed no change immediately or after 1 h; however, after 3 h the emission intensity was enhanced markedly for both 1 and 4 and after 12 h the enhancement was recorded to be over 50% (Figure S24 in the Supporting Information). Again, similar results were obtained when the acidities of the solutions of 1 and 4 were lowered to pH 4. Emission intensity enhancement at 337 nm clearly indicated release of the interacting BSA from the complexes with the addition of chelating agent or to lowering of the pH. From the above discussion it is clear that binding and unbinding of BSA with the drugs is reversible, which may be relevant for further in vivo implications of these drugs. Mass Spectral Studies. Information about the composition and stability of the ligands and complexes has also been acquired from ESI-MS spectral studies (Figures S13−S15 in the Supporting Information). In their ESI-MS the complexes displayed a common feature: strong peaks due to M − Cl and M + H2O (m/z: 532.1662, 585.1752, 1; 519.1448, 572.1476, 2; 499.1561, 552.1415, 3; 622.2264, 675.2285, 4; 609.2026, 662.1887, 5; 589.2148, 642.1921, 6). Notably, 1−6 do not show a molecular ion peak; rather, they exhibited peaks due to M − Cl and M + H2O, suggesting that the chloro group is highly labile and the ensuing species easily interacts with H2O to form aqua complexes. Hydrolysis Study and Interaction of Nucleobase. Hydrolysis of the M−Cl bond symbolizes an activation step for transition-metal anticancer complexes.40 M−OH2 aqua complexes are habitually more reactive than the related chloro J

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Figure 10. (right) Docked model of 2 located within the hydrophobic pocket of HSA (PDB ID: 1h9z) and (left) the interaction mode between 2 (stick) and HSA (cartoon).

Figure 11. Images of Hoechst 33342/PI stained A549 cells after 24 h of treatment with different concentrations of 1, 2, and 4. White, green, yellow, and red arrows show live, early apoptotic, late apoptotic, and necrotic cells, respectively.

Molecular Docking with DNA. To understand the mode of interaction and binding affinity, 1−6 have been subjected for docking with B-DNA (PDB ID: 1BNA) (Figure S27 in the Supporting Information). The results revealed that 1−6 interact with DNA via an electrostatic mode involving outside edge-stacking interactions with oxygen of the phosphate backbone from DNA. The planarity of the imine core is suitable for strong π−π stacking interactions, and the complexes fit well with the DNA strands. From the resulting docked structures it is clear that 1−6 fit well into the minor groove of the targeted DNA and G−C rich region, stabilized by van der Waals interactions and hydrophobic contacts (Figure 9 and Figures S28−S32 in the Supporting Information).49 Relative binding energies for the resulting structures have been found to be −266.69 (1), −264.55 (2), −267.03 (3), −264.26 (4), −267.09 (5), and −271.82 (6) eV. The overall results corroborated well with spectroscopic results and molecular modeling. Molecular Docking with HSA. Further, docking studies have been carried out to establish the binding mode of 1−6 with the most probable binding sites of human serum albumin (HSA). HSA has been chosen over BSA, as it gives a clear idea about human protein binding interactions. The crystal structure of HSA has revealed that it consists of three structurally

homologous domains: I (residues 1−195), II (196−383), and III (384−585). The principal region for binding of HSA with complexes is the hydrophobic environment of the subdomains IIA and IIIA, corresponding to sites I and II, respectively, and the tryptophan residue (Trp-214) in subdomain IIA. The large hydrophobic cavity in subdomain IIA can accommodate these complexes. The molecular docking pattern for 1−6 with HSA indicated that complexes are localized within the subdomain IIA hydrophobic cavity. The binding result indicated that 2 is in close proximity with hydrophobic residues such as VAL 462, GLN 459, ARG 197, LEU 198, ALA 194, VAL 455, ALA 191, ASN 458, VAL 456, LYS 190, ASN 429, and ASP 187 (within 4 Å) and establishment of a hydrophobic interaction between them (Figure 10 and Figures S33−S37 and Table S4 in the Supporting Information). In addition, a number of electrostatic and hydrogen-bonding interactions between the complexes and HSA are also operative.50 The results suggested that the occurrence of hydrogen bonding lowered the hydrophilicity and improved the hydrophobicity to keep the 2·HSA system stable. They also revealed that the interaction between 2 and HSA is dominated by hydrophobic forces. Similar results have been obtained with 1 and 3−6. Our results also offer good structural evidence to explain the efficient fluorescence quenching of BSA in the presence of 1−6. K

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Figure 12. Percentage cell distribution in different phases of cell cycle after treatment with 1, 2, and 4 in a concentration-dependent manner, including the IC50 values along with control.

Figure 13. Increased ROS generation after treatment with 1, 2, and 4, as analyzed by dichlorodihydrofluorescein diacetate (DCFH-DA), an ROSdetecting dye.

Determination of IC50 Value. The complexes under investigation have been screened toward cytotoxicity, proliferation, and IC50 by MTT assay at different concentrations against A549 cells. These induce cytotoxicity in a concentration-dependent manner. An analysis of the MTT assay for 1−3 displayed their IC50 values at 8 (1), 6 (2) and >20 (3) μM, while for 4−6 the values were 30 (6) μM (Figure S37 in the Supporting Information). The IC50 value of cisplatin has also been determined under analogous conditions and was found to be 6 μg/mL (20 μM) toward a lung cancer cell line (A549). Here we conclude that cytotoxicity

toward A549 cells increases with increasing concentrations of the drugs from lower to higher values. The data obtained from MTT assay clearly indicated that 1 and 2 are more cytotoxic and antiproliferative toward A549 cells relative to cisplatin, which was taken as a positive control. Estimated IC50 values for these complexes are superior to those reported by Kandioller et al.51 against A549 cells. On the basis of their IC50 values the cytotoxicity of these compounds has been arranged in the order 2 > 1 > 4 > 3 > 5 > 6 (Figure S38 in the Supporting Information). L

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Organometallics Changes in Nuclear Morphology of Cells. Anticancer agents exhibit their activity via apoptosis, which is taken as a dependable marker for the assessment of potential of anticancer drugs.52 Changes in the nuclear morphology of the cells during apoptosis can be followed in either fixed tissue or live cells using fluorescent dyes.53 DNA binding dyes Hoechst 33342/PI have been used to investigate the morphological changes in the nucleus and inhibition of cell growth by apoptosis. During morphological analysis using the Hoechst/PI double-staining method it was observed that cell death increased due to cell membrane disruption with an increase in the concentration of each drug, as nuclei appeared light red with a bluish background (Figure 11). Condensation of chromatin is one of the major properties of apoptosis. It was also observed with bright blue fluorescence during morphological analysis in A549 cells treated with 1, 2, and 4. Increase in necrotic cell death in a dose-dependent manner with an increase in concentration of the compounds has also been observed (nucleus with high-red and low-blue fluorescence). However, untreated A549 cells exhibited almost uniform blue fluorescence (low level) that indicates healthy cells and very few showed high-red fluorescence (i.e. dead cells). Increasing the intensity of red fluorescence in a dosedependent manner indicated an increase in cell death and antiproliferative activity against A549 cells, which has been previously shown in MTT assay for these compounds. A similar study has been reported by Pawlowska et al., where the nuclear morphology of CHO cells after treatment with the anticancer agent imidazoacridinone C-1311 changes in a time-dependent manner.54 The cells with condensed, intensely stained fragmented chromatin are defined as apoptotic cells, and those stained with PI are defined as dead. Likewise, Cao et al. has reported that an Ir(III) complex induces mitochondriamediated apoptosis in HeLa cells.55 Cell Cycle Analysis. To investigate the role of these complexes in cell cycle arrest in a dose-dependent manner, FACS analysis for cell cycle distribution has been performed on asynchronous A549 cells. The effect of 1, 2, and 4 on the cell cycle distribution of A549 cells and percentage cell distribution is depicted in Figure S39 in the Supporting Information and Figure 12, respectively. The results showed that as the concentration of 2 increases from 4 to 8 μM, including IC50 (6 μM), the population of the cells in the sub G1 phase increases relative to control (untreated A549 cells), which indicates that apoptotic cell death is increasing in a dose-dependent manner. For 1 there is an increase in cell population in sub G1 in comparison to control (untreated cells) and cell population in the sub G1 phase is at a maximum at the IC50 value. Cell population also increases in the S phase in comparison to control, which clearly indicated that 1 is capable of arresting the cells in the S phase of the cell cycle. A similar study has been reported by Yellol et al. in HT29 cells treated with cyclometalated benzimidazole Ru(II), Ir(III), and Rh(III) complexes.53 There was an increase in cell population in the G2/M phase after treatment of A549 cells with 4; the number of cells arrested in the G2/M phase was enhanced from 15% (control cells) to 20.3% (cell treated with 4). This suggested that 4 selectively arrested the cell cycle in the G2/M phase after 24 h of treatment relative to the control. Similar results have been reported on H446 and H1688 human small-cell lung cancer cells, where treatment with evodiamine induces G2/M arrest and apoptosis.56

Generation of Reactive Oxygen Species (ROS). To investigate the efficacy of 1, 2, and 4 in inducing ROS generation, A549 cells were treated with these compounds close to their IC50 values (8 and 10 μM, 1; 6 and 8 μM, 2; 20 and 25 μM, 4; Figure 13). All three compounds generated ROS in a concentration-dependent manner in comparison to control. It has already been reported that the ability of the drugs to induce apoptosis in cancer cells depends upon the ability of the cancer cells to generate ROS. From the available data we conclude that cells treated with 1, 2, and 4 increase ROS generation in A549 cells and consequently induce apoptosis. Notably, untreated A549 cells show low levels of ROS in comparison to treated cells. An increase in the generation of ROS with an increase in the concentration of 1, 2, and 4 with respect to control clearly indicated enhancement of the oxidative stress, which leads to cellular damage. On the basis of the oxidative stress, an elevated level of ROS can induce apoptosis in cancer cells.57 A similar study has been reported by Song et al. with iridium complexes, where the level of ROS increases after adding the drugs, while Kandioller et al. have shown ROS generation by a rhodium complex in a concentration- dependent manner in HL60 cells.58 Overall results from various studies clearly illustrated that all rhodium complexes exhibit better anticancer activity relative to their iridium counterparts. Ir(III) is a third-row transition metal having a relativistic destabilization due to the presence of a 5d shell and displays a different mechanism of action (MOA) in comparison to its lighter equivalent Rh(III). Hence, analogous Rh(III) and Ir(III) compounds should not only display different kinetic responses but also may perhaps favor binding with dissimilar target molecules in biological systems.59,60 This is why the responses of Rh(III) and Ir(III) complexes toward CT DNA, BSA, and the A549 cancer cell line are different.

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CONCLUSIONS In conclusion, through this work six new C−H activated cyclometalated Rh(III)/Ir(III) complexes have been synthesized and characterized by various physicochemical techniques. The structures of 1−3 have been validated by single-crystal Xray analyses. The properties of the complexes have been finetuned by changing the functional groups, viz. −COOMe, −NO2, and −CN, in the ligand. It has been established that complexes effectively bind with DNA through intercalative/ electrostatic interactions or display covalent bonding. Further, molecular docking studies revealed that these complexes bind with the minor groove of DNA by insertion of the imine core through electrostatic interaction, whereas with protein they bind with the hydrophobic residues such as VAL 462, GLN 459, ARG 197, LEU 198, ALA 194, VAL 455, ALA 191, ASN 458, VAL 456, LYS 190, ASN 429, ASP 187, etc. and are situated within the subdomain IIA cavity. Moreover, complexes 1−6 exhibit significant cytotoxicity toward the A549 cell line. Among these, 1 and 2 displayed the lowest IC50 value (lower than that of cisplatin), the highest ROS generation, and prominent blebbing at low concentration (c = 6−10 μM). The results shown by these complexes can be informative in understanding the MOA for interaction with serum albumin, DNA, and cancer cells. This new approach for the lucid design of organorhodium and -iridium drugs may become useful in the future for treating platinum-resistant cancers. M

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.organomet.5b00475. CCDC Nos. 1050647−1050649 (1−3) also contain supplementary crystallographic data for this paper. They can be had free of charge via https://summary.ccdc.cam. ac.uk/structure-summary-form (or from the CCDC, 12 Union Road, Cambridge CB2 1EZ, U.K.; fax, +44-1223-336033; email, [email protected]). 1 H and 13C NMR spectra, ESI-MS, UV−vis titration curves, fluorescence spectra, and Tables S1−S5 (PDF) Crystallographic data for 1−3 (CIF)

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AUTHOR INFORMATION

Corresponding Author

*D.S.P.: tel, + 91 542 6702480; fax, + 91 542 2368174; e-mail, [email protected]. Notes

The authors declare no competing financial interest. ∥ Deceased May 8th, 2015. Included as posthumous coauthor.

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ACKNOWLEDGMENTS We gratefully acknowledge financial support from the Department of Science and Technology (DST), New Delhi, India, for providing financial assistance through the scheme SR/S1/IC25/2011 and also to the UGC, New Delhi, India, for the award of the Senior Research Fellowship 19-6/2011(i)EU-IV to S.M. We are thankful to the Head, Department of Chemistry, Faculty of Science, Banaras Hindu University, Varanasi (U.P.), India, for extending use of the laboratory facilities. We are deeply obliged to the late Prof. M. S. Hundal, Department of Chemistry, Guru Nanak Dev University, Amritsar (Punjab), India, for providing single-crystal X-ray data and helpful suggestions.

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DOI: 10.1021/acs.organomet.5b00475 Organometallics XXXX, XXX, XXX−XXX

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DOI: 10.1021/acs.organomet.5b00475 Organometallics XXXX, XXX, XXX−XXX