benzene complexes containing aroylthiourea ligands

NJC PAPER

Cite this: New J. Chem., 2017, 41, 2672

Synthesis of Ru(II)–benzene complexes containing aroylthiourea ligands, and their binding with biomolecules and in vitro cytotoxicity through apoptosis† Kumaramangalam Jeyalakshmi,a Jebiti Haribabu,a Chandrasekar Balachandran,b Nattamai S. P. Bhuvanesh,c Nobuhiko Emib and Ramasamy Karvembu*a The reaction of [RuCl2(Z6-benzene)]2 with aroylthiourea resulted in the formation of Ru(II) complexes of the type [RuCl2(Z6-benzene)L] (L = monodentate aroylthiourea ligand). The complexes were well characterized using UV-Visible, FT-IR, NMR and mass spectroscopic techniques. Single crystal X-ray diffraction confirmed the monodentate coordination of the ligand through a sulfur atom. The interaction of the Ru(II) complexes with calf thymus DNA (CT DNA) was investigated using UV-Visible and fluorescence spectroscopic methods, and viscosity measurements. The binding ability of the complexes with bovine serum albumin (BSA) was explored using UV-Visible and fluorescence experiments. The results showed that the complexes interact with the biomolecules with appreciable binding constants. The gel electrophoresis technique was used to demonstrate the unwinding of the supercoiled DNA to its nicked form. The cytotoxicity of the Ru(II) complexes was screened for a panel of cancer cell lines like HepG2, A549, MCF7 and SKOV3. Complexes 1, 2 and 3 showed modest activity at the concentration of 31.25 mg mL1 against HepG2 cells. Complexes 1 and 3 displayed moderate cytotoxicity at the

Received 12th October 2016, Accepted 20th February 2017 DOI: 10.1039/c6nj03099h

concentration of 62.5 mg mL1 against A549 and SKOV3 respectively. Low cytotoxicity was observed for all the complexes against MCF7. Advantageously, complexes exhibited only less toxicity against Vero normal cells. Further DNA fragmentation, flow cytometry and fluorescence staining [DAPI (blue), FITC (green) and PI (red)] for the detection of apoptosis in HepG2 cells were carried out. The above methods

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demonstrated that the complexes have a significant ability to induce cell death by apoptosis.

1. Introduction The success of cisplatin as an anticancer drug enticed many researchers to search for potential metallodrugs with enhanced pharmacological activities since cisplatin suffers from severe side effects like neuro-, hepato- and nephrotoxicity.1,2 Ruthenium complexes have attracted attention as anticancer drugs due to similar ligand exchange kinetics to that of platinum. Further, they are quite selective for cancer cells due to their ability to mimic iron in binding to biomolecules. The earlier interest in the anticancer activity of ruthenium compounds was provoked a

Department of Chemistry, National Institute of Technology, Tiruchirappalli 620015, India. E-mail: [email protected] b Department of Hematology, Fujita Health University, 1-98, Dengakugakubo, Kutsukake-cho, Toyoake, Aichi 470-1192, Japan c Department of Chemistry, Texas A & M University, College Station, TX 77842, USA † Electronic supplementary information (ESI) available: Details of NMR spectra, binding studies, DNA cleavage and cytotoxicity of all the complexes. Crystallographic data for the structures reported. CCDC 1477275 and 1477276. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c6nj03099h

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by Clarke’s observation that [RuCl3(NH3)3] is an active anticancer agent.3 Another two important Ru(III) complexes, trans-[RuCl4(DMSO)(Im)]ImH (NAMI-A, Im = imidazole)4 and trans-[RuCl4(Ind)2]IndH (KP1019, Ind = indazole),5 are under clinical trial. It was found that KP1019 is cytotoxic to cancer cells and that NAMI-A is relatively non-toxic but has antimetastatic activity. Several other classical ruthenium complexes have shown promising anticancer activity.6,7 Clarke’s investigation proved that the anticancer activity of Ru(III) complexes is dependent on their in vivo reduction to more labile Ru(II) complexes. Therefore the ‘‘activation by reduction’’ strategy has induced interest towards the anticancer activity of Ru(II) complexes. Though many Ru(II) organometallic compounds were reported as anticancer agents, half-sandwich RuCl2(arene) type complexes received special attention. This was because, the inert hydrophobic arene moiety, the labile chloride ligand and the tunable ligand under investigation might be helpful to enhance the pharmacological properties of the complexes. The nature of the arene could influence cell uptake and interactions with potential targets. The chloride

This journal is © The Royal Society of Chemistry and the Centre National de la Recherche Scientifique 2017

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which occupies the biomolecule binding site on the Ru is expected to control the activation of these complexes. In this line, Ru(II)–arene complexes offered great promise towards the development of anticancer drugs.8–27 The first complex of this kind which was evaluated for cytotoxic property was [Ru(Z6-benzene)Cl2(metronidazole)]

Fig. 1

Structure of (a) [Ru(Z6-benzene)Cl2(metronidazole)] and (b) RAPTA-B.

Scheme 1

(metronidazole = 1-b-hydroxyethyl-2-methyl-5-nitroimidazole) (Fig. 1). Another important class of Ru(II)–arene complexes bearing pta ligands, [RuCl2(Z6-benzene)(pta)] (RAPTA-B) and [RuCl2(Z6-p-cymene)(pta)] (RAPTA-C) (pta = 1,3,5-triaza-7phosphaadamantane), have recently attracted considerable attention due to the promising in vivo activity towards inhibition of metastasis growth, together with high selectivity and low general toxicity (Fig. 1).28 Interestingly, [RuCl2(Z6-benzene)(DMSO)] inhibited the DNA relaxation activity of topoisomerase II.29 The structure of the ligands plays a crucial role in deciding the pharmacological properties of the complexes. We intended to work on aroylthiourea ligands due to their versatile coordination behavior and broad spectrum of biological activities like anticancer, antiviral, antibacterial, antifungal, antitubercular, antiinflammatory, herbicidal, insecticidal, etc.30–34 Herein we describe the synthesis, structure, DNA and protein binding, DNA cleavage,

Synthetic route for the ligands and their Ru(II)–benzene complexes.


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in vitro cytotoxicity, DNA fragmentation and confocal imaging of Ru–benzene complexes containing aroylthiourea ligands. The complexes we report are similar to the RAPTA-B backbone, with the pta ligand being replaced by aroylthiourea. Moreover there are only a few reports on Ru–benzene complexes with regard to their anticancer activity and this is the first report on Ru–benzene complexes containing aroylthiourea ligands evaluated for their biological applications.

2. Results and discussion 2.1

Synthesis and spectroscopy

The aroylthiourea ligands (L1–L4) used in this work have been synthesized and reported by us earlier.35 The reactions of the ligands with the Ru–benzene dimer produced [RuCl2(Z6-benzene)L] (1–4) (Scheme 1). The Ru(II) complexes were characterized by UV-Visible, FT-IR, mass and NMR spectroscopic techniques. The UV-Visible spectra of the complexes showed new bands around 349–364 and 444–461 nm, which correspond to MLCT and d - d transitions respectively. The FT-IR spectra of the ligands showed bands in the region 3076–3312, 1661–1657 and 1256–1271 cm1 which were assigned to n(N–H), n(CQO) and n(CQS) respectively. The N–H and CQO bands were unaltered in the spectra of the

Table 1

complexes while n(CQS) was decreased (1171–1201 cm1) which confirmed the coordination of only sulfur to the Ru ion. In the 1H NMR spectra of the complexes the carbonyl and thiocarbonyl attached N–H and thiocarbonyl attached N–H protons resonated around 11.49–12.54 and 11.09–11.82 ppm respectively. The signal observed at 5.98 ppm in the spectra of all the complexes confirmed the presence of a benzene moiety (Fig. S1–S4, ESI†). All the other aromatic protons displayed signals in the expected region (8.46–7.27 ppm). In the 13C NMR spectra of the complexes, the signals appeared around 122.7– 138.4 ppm, which corresponded to the aromatic carbons in the complexes. The thiocarbonyl and carbonyl carbons were characterized by the signals around 179.2–181.2 and 162.4– 162.7 ppm respectively. A new resonance appeared in the spectra of the complexes at 88.1 ppm was assigned to benzene (Fig. S5–S8, ESI†). The signals at 18.1 and 48.6 ppm in the spectra of complexes 2 and 3 respectively were attributed to the aliphatic carbons. 2.2

Crystal structure

Tables 1 and 2 summarize the crystallographic data of complexes 1 and 3. Single crystals suitable for X-ray diffraction were obtained by the slow evaporation of chloroform/methanol solution of the complexes. Complexes 1 and 3 crystallized in

Crystal data and structure refinement for complexes 1 and 3

Empirical formula Formula weight Temperature (K) Wavelength (Å) Crystal system Space group Unit cell dimensions a (Å) b (Å) c (Å) a (1) b (1) g (1) Volume (Å3) Z Density (calculated) (Mg m3) Absorption coefficient (mm1) F(000) Crystal size (mm3) Theta range for data collection (1) Index ranges Reflections collected Independent reflections [R(int)] Completeness to theta = 25.242/27.501 Absorption correction Max. and min. transmission Refinement method Data/restraints/parameters Goodness-of-fit on F2 Final R indices [I 4 2sigma(I)] R Indices (all data) Largest diff. peak and hole (e Å3)

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1

3

C18H16Cl2N2ORuS2 512.42 150 0.71073 Monoclinic P21/c

C19H18Cl2N2ORuS2 526.44 110.15 0.71073 Triclinic P1%

11.674(2) 10.818(2) 15.646(3) 90 99.37(3) 90 1949.8(7) 4 1.746 1.302 1024 0.32 0.315 0.106 1.768 to 27.540 15 r h r 15, 14 r k r 13, 20 r l r 20 21 897 4445 [R(int) = 0.0321] 99.7 Semi-empirical from equivalents 0.7456 and 0.6105 Full-matrix least-squares on F2 4445/0/235 1.037 R1 = 0.0204, wR2 = 0.0490 R1 = 0.0240, wR2 = 0.0509 0.430 and 0.368

6.5098(19) 11.428(3) 14.534(4) 101.993(3) 95.432(4) 105.472(3) 1006.3(5) 2 1.737 1.264 528 0.15 0.04 0.03 1.451 to 27.550 8 r h r 8, 14 r k r 14, 18 r l r 18 11 564 4551 [R(int) = 0.0466] 99.7 Semi-empirical from equivalents 0.7456 and 0.5274 Full-matrix least-squares on F2 4551/0/244 1.043 R1 = 0.0399, wR2 = 0.1051 R1 = 0.0441, wR2 = 0.1089 1.867 and 1.652


Paper Table 2

NJC Selected bond lengths (Å) and angles (1) of complexes 1 and 3

Bond length/angle (Å)

1

3

Ru–arene (centroid) Ru(1)–Cl(1) Ru(1)–Cl(2) Ru(1)–S(1) Ru(1)–C(14) Ru(1)–C(15) Ru(1)–C(16) Ru(1)–C(17) Ru(1)–C(18) Ru(1)–C(13)/C(19) Cl(2)–Ru(1)–Cl(1) Cl(2)–Ru(1)–S(1) S(1)–Ru(1)–Cl(1)

1.658 2.4247(8) 2.4227(6) 2.4087(7) 2.1776(19) 2.1821(19) 2.1902(19) 2.1698(19) 2.1805(18) 2.1686(18) 89.251(19) 92.04(3) 88.99(2)

1.665 2.4350(9) 2.4149(10) 2.4139(8) 2.190(3) 2.173(3) 2.173(3) 2.182(3) 2.185(3) 2.171(3) 87.15(3) 88.99(3) 92.63(3)

the monoclinic P21/c and triclinic P1% space group respectively. The complexes featured a half-sandwich three legged piano stool structure about the metal centre (Fig. 2 and 3). It was clear from the crystal structures that the complexes adopted a ‘pianostool’ geometrical configuration with the p-bonded Z6-benzene, monodentate aroylthiourea and two chloride ions. The Ru–C distances were in the range of 2.1686–2.197 Å with an average of 2.1781 (1) and 2.1806 (3) Å.36a–c The Ru–Cl and Ru–S bond lengths were found to be in the range of 2.4149–2.4350 and

Fig. 2

Fig. 3

2.407–2.414 Å respectively, which were comparable with the earlier reports.37 The Ru–benzene centroid distances were 1.658 and 1.665 Å for 1 and 3 respectively.38 There were intramolecular hydrogen bonds (N–H Cl and N–H O) in 1 with H Cl and H O distances of 2.22 and 1.95 Å respectively. Two intramolecular hydrogen bonds were detected in 3 between N–H and two chloride ions [N–H Cl(1) and N–H Cl(2)] with H Cl(1) and H Cl(2) distances of 2.49 and 2.51 Å respectively. The hydrogen bonding was also noted between N–H and O with a H O distance of 2.00 Å in 3. 2.3

Lipophilicity

The hydrophobicity of the compounds determines their penetration behavior across the cell membrane, and is investigated in terms of the partition coefficient (log P).39 The log P value depends on the solubility of the compound in aqueous versus organic medium. The log P values of L1–L4 were 1.923, 1.912, 1.610 and 2.232 respectively, which meet the criteria for their complexes to be cytotoxic. 2.4

Binding of the complexes with CT DNA

To explore the interaction mode of the complexes with DNA, the changes in the absorbance of the complexes on adding aliquots of CT DNA were monitored. Usually intercalative mode of binding of the complexes with DNA shows hypochromism with or without red shift.40a,b Non-intercalative/electrostatic interaction results in hyperchromism.41a,b From the absorption spectra, it was clear that upon the addition of CT DNA to the complexes, hypochromism occurred in the intraligand band (23–40%) with 2–3 nm red shift (Fig. 4 and Fig. S9, ESI†), which indicated intercalative mode of binding.42–44 To compare the DNA binding ability of the complexes, the equilibrium binding constant (Kb) was calculated using the equation45 [DNA]/(ea ef) = [DNA]/(eb ef) + 1/Kb (eb ef), where [DNA] is the concentration of DNA in base pairs, ea is the apparent extinction coefficient value

Thermal ellipsoid (50%) plot of 1.

Thermal ellipsoid (50%) plot of 3.

Fig. 4 Absorption spectra of complex 4 in Tris-HCl buffer upon addition of CT DNA. [Complex] = 2.5 105 M, [DNA] = 0–50 mM. The arrow shows that the absorption intensity decreases upon increasing the CT DNA concentration.


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Fig. 5 Plot of [DNA]/(ea ef) versus [DNA] for the titration of the complexes with CT DNA.

Fig. 6 Fluorescence quenching curves of EB bound to DNA in the presence of 4. [DNA] = 5 mM, [EB] = 5 mM and [complex] = 0–50 mM.

Table 3 DNA binding constant (Kb), quenching constant (Kq) and apparent binding constant (Kapp) values

Complex

Kb (M1)

1 2 3 4

1.17 1.41 1.62 7.30

104 104 104 104

Kq (M1)

0.15 0.09 0.13 0.24

3.54 3.65 3.77 3.91

104 104 104 104

Kapp (M1)

0.14 0.11 0.11 0.07

1.77 1.82 1.88 1.95

106 106 106 106

0.09 0.11 0.16 0.13

found by calculating A(observed)/[complex], ef is the extinction coefficient for the free compound and eb is the extinction coefficient for the compound in the fully bound form. A plot of [DNA]/(ea ef) versus [DNA] gave Kb as the ratio of slope to the intercept (Fig. 5). The calculated intrinsic equilibrium binding constants (Kb) for 1–4 were in the range of 1.17 104–7.30 104 M1 respectively (Table 3). The binding affinity of the complexes followed the order 4 4 3 4 2 4 1. The Kb value of 1–4 was comparable with the reported Kb of similar ruthenium–arene complexes.42,46,47 The competitive ethidium bromide (EB) binding studies provided support for the interaction of the complexes with DNA.48–50 Upon addition of the complexes (1–4) (0–50 mM) to CT DNA pretreated with EB in 5% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.2, the emission intensity of DNA-bound EB at 605 nm decreases (Fig. 6 and Fig. S10, ESI†). The percentage of quenching for complexes 1–4 was found to be 63.4, 64.1, 64.6 and 65.3% respectively with 10–12 nm bathochromic shift. The extent of quenching revealed the extent of displacement of the EB molecules from the CT DNA. According to the Stern–Volmer equation, relative fluorescence is directly proportional to the concentration of the quencher. The Stern–Volmer equation is given by F0/F = 1 + Kq[Q], where F0 and F are the fluorescence intensities in the absence and presence of the complex respectively, Kq is a linear Stern–Volmer quenching constant, and [Q] is the concentration of the complex. The slope of the plot of F0/F versus [Q] gave Kq (Fig. 7). From the plot of the observed intensities against the complex concentration, the values of the apparent DNA binding constant (Kapp) were calculated using

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Fig. 7 Stern–Volmer plot of fluorescence titrations of the complexes with CT DNA.

the equation51 KEB[EB] = Kapp[complex], where [complex] is the concentration of the complex at 50% reduction in the fluorescence intensity of EB, KEB = 1.0 107 M1 and [EB] = 5 mM. Kq and Kapp values are listed in Table 3. The interaction of the complexes with CT DNA followed the order 4 4 3 4 2 4 1. The results are in agreement with those found from the electronic absorption studies. Viscosity measurements illustrated the effect of the complexes on the specific relative viscosity of DNA. Since the specific relative viscosity (Z/Z0; Z and Z0 are the specific viscosities of DNA in the presence and absence of the complexes, respectively) will have an effect when the DNA contour length changes, it is helpful in the determination of the mode of interaction of the complexes with DNA. The increase in the contour length associated with the separation of DNA base pairs caused by intercalation of a classical DNA intercalator like EB shows a significant increase in the viscosity


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Fig. 8 Effect of increasing amounts of complexes 1–4 on the relative viscosity of CT DNA.

Fig. 10 Absorption spectra of BSA (10 mM) and BSA with 1–4 (4 mM).

of the DNA solutions. In contrast, partial and/or non-intercalation of the complex could result in a less pronounced effect on the viscosity.52 The viscosity of DNA increased on incremental addition of the complexes, which showed the intercalative mode of binding (Fig. 8). Complex 4 showed a comparatively increased effect on the viscosity among the complexes investigated. Complex 1 did not cause a steady increase in the viscosity of DNA. The effect of the complexes on the viscosity of DNA followed the order 4 4 3 4 2 4 1, which was in good agreement with the results obtained from the spectroscopic studies.

(SC) DNA (Form I) into nicked circular (NC) DNA (Form II), and the DNA cleaving ability of the complexes followed the order 3 (48.45%) 4 4 (44.53%) 4 1 (34.46%) E 2 (33.32%). In order to investigate the mechanism of DNA cleavage, experiments were performed by incubating 3 with DNA for 4 h in the presence of singlet oxygen scavenger (NaN3) and hydroxyl radical scavenger (DMSO).53a There was no reduction in the percentage of cleavage even in the presence of scavengers (Fig. S11, ESI†), which ruled out the oxidative and photo-induced cleavage mechanisms. Therefore the DNA cleavage might occur through a hydrolytic pathway as observed in Ru–polypyridyl complexes.53b

2.5

Unwinding of supercoiled DNA

A change in the electrophoretic mobility of plasmid DNA on agarose gel is commonly taken as evidence for DNA–drug interactions. The change in the DNA structure from supercoiled form to its nicked or linear form triggers a change in the extent of migration in the gel. In order to explore the cleaving ability of the Ru(II)–benzene complexes, supercoiled (SC) pBR322 DNA (40 mM in base pairs) was incubated at 37 1C with the complexes (50 mM) in a 5% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.2 for 4 h in the absence of any external agent. Fig. 9 illustrates the gel electrophoretic separation showing the cleavage of plasmid pBR322 DNA by the Ru(II) complexes under identical conditions. All the complexes investigated cleaved supercoiled

Fig. 9 Cleavage of supercoiled pBR322 DNA (40 mM) by complexes 1–4 in a buffer containing 5% DMF/5 mM Tris HCl/50 mM NaCl at pH = 7.2 and 37 1C with an incubation time of 4 h. Lane 1, DNA; lane 2, DNA + 1 (50 mM); lane 3, DNA + 2 (50 mM); lane 4, DNA + 3 (50 mM); lane 5, DNA + 4 (50 mM). Forms SC and NC are supercoiled and nicked circular DNA, respectively.

2.6

Binding of the complexes with protein

The electronic spectra of BSA in the absence and presence of the complexes are depicted in Fig. 10. The enhancement in the absorption band of BSA at 280 nm on addition of the complexes indicated a static type of quenching mechanism as reported.54,55 The quenching experiments were carried out to investigate the interaction of the complexes with BSA. Upon addition of the complexes (1–4), the emission intensity of BSA was decreased [77.8 (1), 84.9 (2), 70.9 (3), 82.2% (4)] with 2–3 nm hypsochromic shift (Fig. 11 and Fig. S12, ESI†).56 The extent of quenching of fluorescence intensity is expressed by Stern–Volmer constant (Kq) which is a measure of protein binding affinity of the complexes.57 The value of Kq obtained as the slope of the linear plot of F0/F versus [complex] followed the order 2 4 4 4 1 4 3 (Fig. 12). Further, the equilibrium binding constant was evaluated using the Scatchard equation, log[(F0 F)/F] = log KBSA + n log[Q], where F and F0 are the fluorescence intensity in the presence and absence of the complexes, KBSA is the binding constant and n is the number of binding sites. The KBSA values were derived from the plot between log[(F0 F)/F] and log[Q] (Fig. 13). The values of KBSA and Kq suggested that complex 2 exhibited good binding ability compared to the other complexes under investigation (Table 4). Vekshin suggested a useful method known as synchronous fluorescence spectroscopy which gives valuable information on


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Fig. 11 Fluorescence quenching curves of BSA in the absence and presence of 2. [BSA] = 1 mM and [complex] = 0–20 mM.

Fig. 13 Scatchard plot of the fluorescence titrations of the complexes with BSA.

Table 4 Protein binding constant (KBSA), quenching constant (Kq) and number of binding sites (n) values

Complex 1 2 3 4

KBSA (M1) 1.66 8.63 1.16 1.74

5

10 105 105 105

Kq (M1)

0.18 0.14 0.11 0.15

1.67 2.44 1.20 2.26

n 5

10 105 105 105

0.08 0.10 0.13 0.11

1.03 1.33 0.97 0.99

Fig. 12 Stern–Volmer plot of the fluorescence titrations of the complexes with BSA.

the molecular microenvironment, particularly in the vicinity of the fluorophore functional groups.58 To get an insight into the structural changes in BSA on addition of the complexes, synchronous fluorescence spectra were recorded.59 The difference between the excitation and emission wavelengths (Dl = lem lex) indicates the types of chromophores. A higher Dl value, such as 60 nm, is indicative of the tryptophan residue, whereas a lower Dl value, such as 15 nm, is characteristic of the tyrosine residue.60 In the synchronous fluorescence spectra of BSA at Dl = 15 nm, the incremental addition of the complexes to BSA resulted in quenching of the fluorescence intensity at 304 nm with 61.6, 74.8, 57.0 and 66.8% for complexes 1–4 respectively (Fig. 14 and Fig. S13, ESI†). Similarly quenching was also observed in the synchronous fluorescence spectra of BSA (348 nm) at Dl = 60 nm, with 74.4, 84.6, 71.0 and 79.7% for complexes 1–4 respectively (Fig. 15 and Fig. S14, ESI†). The

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Fig. 14 Synchronous spectra of BSA (1 mM) as a function of concentration of 2 (0–20 mM) with Dl = 15 nm.

results showed that all the complexes affected the microenvironments of both tyrosine and tryptophan residues and the effect was more pronounced towards tryptophan than tyrosine. 2.7

Cytotoxicity

The cytotoxicity of Ru(II)–benzene complexes was studied by MTT assay against HepG2, A549, MCF7, SKOV3 and Vero cell lines and the results were compared with those obtained with


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Fig. 15 Synchronous spectra of BSA (1 mM) as a function of concentration of 2 (0–20 mM) with Dl = 60 nm.

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the well-known anticancer drug cisplatin. Complex solutions of all concentrations used in the experiment decreased the cell viability significantly in a concentration-dependent manner. The graph of percentage of cell death versus concentration is shown in Fig. 16, 17 and Fig. S15, S16 (ESI†). The half minimum inhibitory concentration (IC50) calculated for the complexes is tabulated (Table 5). From the results it was inferred that complexes 1, 2 and 3 showed modest activity towards HepG2 at lower concentration (31.25 mg mL1) with IC50 ranging from 40.80 to 53.14 mM. Complex 4 showed only least activity towards HepG2 cell line with an IC50 value of 162.20 mM. Among the investigated complexes, complexes 1 and 3 showed moderate toxicity against A549 (IC50, 95.60 mM) and SKOV3 cell lines (IC50, 87.52 mM) respectively. The complexes were less toxic towards A549, MCF7 and SKOV3 cancer cell lines and the Vero normal cell line. The cytotoxicity of the investigated complexes was reasonably low compared to that of similar p-cymene complexes.35 The cytotoxicity of 1–4 was compared with that of other reported ruthenium–benzene complexes. The cytotoxicity of 1–4 is superior to that exhibited

Fig. 16 Comparison of anticancer activity of the complexes (1–4) against HepG2 cancer cells. Data are mean SD of three independent experiments with each experiment conducted in triplicate.

Fig. 17 Comparison of anticancer activity of the complexes (1–4) against A549 cancer cells. Data are mean SD of three independent experiments with each experiment conducted in triplicate.


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Table 5 In vitro cytotoxicity of the complexes in HepG2, A549, MCF7 and SKOV3 cancer cell lines

IC50 (mM) Complex

HepG2

A549

MCF7

SKOV3

1 2 3 4 Cisplatin

53.14 50.44 40.80 162.20 21.5

95.60 4250 4250 4250 18.0

151.29 153.93 162.99 154.40 12.0

4250 165.90 87.52 4250 8.3

Fig. 19 Detection of DNA fragmentation by agarose gel electrophoresis (2%). The apoptotic DNA fragmentation was detected by agarose gel electrophoresis in HepG2 cells treated with complexes 1–3 at IC50 concentration (31.25 mg mL1) for 24 h. Lane 1, 1 kb DNA ladder; lane 2, control; lane 3, 1; lane 4, 2; lane 5, 3. The arrow marks indicate the DNA fragmentation.

Fig. 18

IC50 value of reported ruthenium–benzene complexes.

by ruthenium(II)(Z6-N-benzylmaleimide) containing 1,3,5-triaza-7phosphaadamantane (PTA) (IC50, 640 mM). The triphenylphosphine complex of Ru–benzene showed a comparable activity (IC50, 116 mM) against the A549 cancer cell line (Fig. 18).61 The ruthenium–benzene complex bearing the 4-(biphenyl-4-carbonyl)-3-methyl-1-phenyl-5pyrazolonate ligand showed similar half-inhibitory concentration (35.0 mM) against the HepG2 cancer cell line.62 The comparison with the potential drug cisplatin showed that our complexes were less effective. Compounds 1–4 showed varied activity in different cell lines, which exposed the specificity of the complexes against various cancer targets. Further studies on structure–activity relationship and gene profiling, in future, will provide more insight into the specific activity of the complexes against various cancer cell lines. 2.8

Apoptosis mechanism

There are different pathways by which cell death occurs in response to a variety of external and physiological stimuli. Apoptosis (programmed cell death) is a normal component of the development and maintenance of health of multicellular organisms. During apoptosis, cell death occurs in a controlled and regulated fashion whereas other cell death pathways like

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necrosis (unprogrammed cell death) lead to serious health issues.63–65 This makes apoptosis distinct from the other form of cell death. Therefore, fluorescence staining [DAPI (blue), FITC (green) and PI (red) fluorescence] to assess the morphological changes and gel electrophoresis for detecting DNA fragmentation were carried out. DNA fragmentation was demonstrated by incubating HepG2 cells with IC50 concentrations (31.25 mg mL1) of 1, 2 and 3 for 24 h. DNA fragmentation became apparent and the response was dose dependent (Fig. 19). The DNA ladders were visible and complex 3 showed effective DNA fragmentation compared to 1 and 2, which was in agreement with the results obtained from cytotoxicity. The fragmentation pattern observed clearly demonstrates that the cell death mechanism is through apoptosis. Flow cytometry is one of the suitable methods for cell death or apoptosis analysis. The fractions of cell populations in different quadrants were analyzed using quadrant statistics. The lower left quadrant (R2), the lower right quadrant (R3), the upper right quadrant (R4) and the upper left quadrant (R5) contained the living cells, early apoptosis cells, late apoptosis cells and dead cells respectively. To analyze apoptosis, HepG2 cell lines were treated with IC50 concentrations of the complexes (1–3) for 24 h. There was a dramatic increase in the apoptosis effect on HepG2 cells when compared to the control cells (Fig. 20). Complexes 1–3 showed 23.4, 31.6 and 62.3% of apoptosis in HepG2 cells, respectively. It is to be noted that complex 3 exposed more % of late apoptotic cells (46.9%) compared to the other two complexes at 24 h incubation.66


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Fig. 20 Flow cytometry analysis carried out in HepG2 cells (mean SD of three independent experiments with each experiment conducted in triplicate). R2 = live cells; R3 = early apoptotic cells; R4 = late apoptotic cells; R5 = dead cells.

On the basis of overall cell morphology and cell membrane integrity, necrotic and apoptotic cells could be distinguished from one another using confocal microscopy.67,68a,b HepG2 (50 mg mL1) cells treated with the Ru(II)–benzene complexes (1–3) were subjected to confocal microscopic analysis. In this study, DAPI, FITC and PI fluorescence stains were used. DAPI binds strongly to A-T rich regions in DNA and passes through an intact cell membrane. FITC acts as a phosphatidyl serine tracer and suggests the presence of apoptosis. PI can only penetrate cells where the cell membrane has been compromised. Confocal microscopic images showed that significant morphological changes were found in HepG2 cells after treatment with the complexes and apoptotic cells are indicated with arrows in Fig. 21.

3. Conclusion The present work describes the synthesis and characterization of half-sandwich Ru(II)–benzene complexes and their biological applications. Various physicochemical methods validated the interaction of the complexes with DNA via intercalation. The binding constants

were in the order of 104 M1. The complexes also cleaved DNA through a hydrolytic pathway. The protein binding efficacy of the complexes was also demonstrated using UV-Visible and fluorescence spectroscopic techniques. In vitro cytotoxic assay revealed that the complexes (1–3) were active only against HepG2 among the four tested cancer cell lines and the results were compared with those for cisplatin. The DNA fragmentation, flow cytometry and staining methods disclosed that the possible way of cell death was through apoptosis. Unfortunately complex 4 which showed a better binding ability to DNA/protein failed to exhibit cytotoxicity against the four cancer cell lines. Therefore further mechanistic aspects have to be dealt in detail to develop an anticancer drug.

4. Experimental section 4.1

Methods

The melting points were determined on a Lab India instrument and were uncorrected. FT-IR spectra were obtained as KBr pellets using a Nicolet-iS5 spectrophotometer. UV-Visible spectra were recorded using a Shimadzu-2600 spectrophotometer. Emission spectra were


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Fig. 21 DAPI (blue), FITC (green) and PI (red) fluorescence staining for the detection of apoptosis in HepG2 cells. Control: (a1 – DAPI, a2 – FITC, a3 – PI and a4 – merged) and treated (1: b1 – DAPI, b2 – FITC, b3 – PI and b4 – merged); (2: c1 – DAPI, c2 – FITC, c3 – PI and c4 – merged); (3: d1 – DAPI, d2 – FITC, d3 – PI and d4 – merged). The arrows indicate apoptotic cancer cells.

measured on a Jasco V-630 spectrophotometer using 5% DMF in buffer as the solvent. NMR spectra were recorded in DMSO-d6 solvent by using TMS as an internal standard on a Bruker 500 MHz spectrometer. ESI-MS spectra were recorded using a Thermo Exactive Orbitrap mass spectrometer. The lipophilicity of the ligands was measured by the ‘‘shake flask’’ method in octanol– water phase partitions as reported earlier.39 4.2

Synthesis of the ligands

The ligands (L1–L4) were prepared from thiophene-2-carbonylchloride, potassium thiocyanate and primary amine (aniline,

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o-toluidine, benzylamine or 1-naphthylamine) by following a reported procedure.32

4.3

Synthesis of the Ru(II)–benzene complexes

The ruthenium–benzene [RuCl2(Z6-benzene)]2 precursor was synthesized using an earlier reported procedure.69 [RuCl2(Z6-benzene)]2 (0.1000 g, 0.2 mmol) and the ligand (L) were dissolved in 15 mL of toluene and stirred for 14–20 h at 27 1C. The resultant solution was concentrated to 2 mL under reduced pressure, and addition of petroleum ether (60–80 1C) (10 mL) gave an orange solid.


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The product was collected by filtration, washed with petroleum ether and dried in vacuo. 4.3.1 [RuCl2(g6-benzene)L1] (1). L1 (0.1049 g, 0.4 mmol) was used. Yield: 79%. Orange solid. M.p.: 215 1C. Anal. calcd for C18H16Cl2N2ORuS2: C, 42.19; H, 3.15; N, 5.47; S, 12.51. Found: C, 42.07; H, 3.21; N, 5.40; S, 12.41. ESI-MS (m/z): (M 2H+ 2Cl + H+)+ found (calcd) 440.9666 (440.9669). UV-Vis (DMF): lmax, nm (e, dm3 mol1 cm1) 265 (22 533), 350 (9666), 444 (1666). FT-IR (KBr, cm1): 3213 (m; n(amide N–H)), 3115 (s; n(thiourea N–H)), 1655 (s; n(CQO)), 1176 (s; n(CQS)). 1H NMR (500 MHz, DMSO-d6) d, ppm 12.46 (s, 1H), 11.62 (s, 1H), 8.40 (d, J = 2.8 Hz, 1H), 8.07 (d, J = 4.7 Hz, 1H), 7.68 (d, J = 7.7 Hz, 2H), 7.42 (t, J = 7.6 Hz, 2H), 7.28 (t, J = 6.3 Hz, 2H), 5.98 (s, 6H, benzene). 13C NMR (125 MHz, DMSO-d6) d, ppm 179.2 (CQS), 162.5 (CQO), 138.4, 137.1, 135.8, 133.2, 129.2, 129.1, 128.8, 126.8, 124.8 (aromatic carbons of the thiourea ligand), 88.1 (aromatic carbons of benzene). 4.3.2 [RuCl2(g6-benzene)L2] (2). L2 (0.1105 g, 0.4 mmol) was used. Yield: 81%. Orange solid. M.p.: 203 1C. Anal. calcd for C19H18Cl2N2ORuS2: C, 43.35; H, 3.45; N, 5.32; S, 12.18. Found: C, 43.11; H, 3.31; N, 5.26; S, 12.03. ESI-MS (m/z): (M 2H+ 2Cl + H+)+ found (calcd) 454.9820 (454.9826). UV-Vis (DMF): lmax, nm (e, dm3 mol1 cm1) 265 (31 600), 290 (34 066), 349 (15 800), 461 (1966). FT-IR (KBr, cm1): 3220 (m; n(amide N–H)), 3162 (s; n(thiourea N–H)), 1654 (s; n(CQO)), 1171 (s; n(CQS)). 1 H NMR (500 MHz, DMSO-d6) d, ppm 12.15 (s, 1H), 11.68 (s, 1H), 8.42 (d, J = 3.1 Hz, 1H), 8.07 (d, J = 4.6 Hz, 1H), 7.56 (d, J = 7.2 Hz, 1H), 7.32 (d, J = 6.7 Hz, 1H), 7.28–7.22 (m, 3H), 5.98 (s, 6H, benzene), 2.26 (s, 3H). 13C NMR (125 MHz, DMSOd6) d, ppm 180.2 (CQS), 162.6 (CQO), 137.4, 137.1, 135.8, 134.0, 133.2, 130.8, 129.2, 128.8, 127.5, 127.2, 126.6 (aromatic carbons of the thiourea ligand), 88.1 (aromatic carbons of benzene), 18.1 (aliphatic carbon). 4.3.3 [RuCl2(g6-benzene)L3] (3). L3 (0.1105 g, 0.4 mmol) was used. Yield: 82%. Orange solid. M.p.: 230 1C. Anal. calcd for C19H18Cl2N2ORuS2: C, 43.35; H, 3.45; N, 5.32; S, 12.18. Found: C, 43.23; H, 3.19; N, 5.53; S, 12.11. ESI-MS (m/z): (M 2H+ 2Cl + H+)+ found (calcd) 454.9821 (454.9826). UV-Vis (DMF): lmax, nm (e, dm3 mol1 cm1) 265 (17 100), 286 (15 900), 349 (4166), 448 (933). FT-IR (KBr, cm1): 3245 (m; n(amide N–H)), 3161 (s; n(thiourea N–H)), 1651 (s; n(CQO)), 1191 (s; n(CQS)). 1H NMR (500 MHz, DMSO-d6) d, ppm 11.49 (s, 1H), 11.09 (s, 1H), 8.36 (d, J = 2.9 Hz, 1H), 8.02 (d, J = 4.6 Hz, 1H), 7.37 (d, J = 13.3 Hz, 4H), 7.31 (d, J = 5.4 Hz, 1H), 7.24 (d, J = 3.9 Hz, 1H), 5.98 (s, 6H, benzene), 4.87 (d, J = 5.3 Hz, 2H). 13C NMR (125 MHz, DMSO-d6) d, ppm 180.6 (CQS), 162.4 (CQO), 137.7, 137.2, 135.5, 132.9, 129.1, 128.9, 128.0, 127.7 (aromatic carbons of the thiourea ligand), 88.1 (aromatic carbons of benzene), 48.6 (aliphatic carbon). 4.3.4 [RuCl2(g6-benzene)L4] (4). L4 (0.1249 g, 0.4 mmol) was used. Yield: 80%. Orange solid. M.p.: 223 1C. Anal. calcd for C22H18Cl2N2ORuS2: C, 46.98; H, 3.23; N, 4.98; S, 11.40. Found: C, 46.84; H, 3.37; N, 4.81; S, 11.55. ESI-MS (m/z): (M 2H+ 2Cl + H+)+ found (calcd) 490.9819 (490.9826). UV-Vis (DMF): lmax, nm (e, dm3 mol1 cm1) 265 (21 500), 293 (21 433), 364 (7766), 454 (1466). FT-IR (KBr, cm1): 3209 (m; n(amide N–H)), 3143 (s; n(thiourea N–H)), 1656 (s; n(CQO)), 1201 (s; n(CQS)).

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H NMR (500 MHz, DMSO-d6) d, ppm 12.54 (s, 1H), 11.82 (s, 1H), 8.46 (s, 1H), 8.10 (d, J = 4.6 Hz, 1H), 8.02 (d, J = 7.7 Hz, 1H), 7.98 (d, J = 7.8 Hz, 1H), 7.94 (d, J = 8.2 Hz, 1H), 7.81 (d, J = 7.2 Hz, 1H), 7.60 (q, J = 8.6 Hz, 3H), 7.30 (d, J = 4.1 Hz, 1H), 5.98 (s, 6H, benzene). 13C NMR (125 MHz, DMSO-d6) d, ppm 181.2 (CQS), 162.7 (CQO), 137.2, 135.8, 134.7, 134.2, 133.2, 129.2, 129.1, 128.8, 127.8, 127.2, 126.8, 125.9, 125.1, 122.7 (aromatic carbons of the thiourea ligand), 88.1 (aromatic carbons of benzene). 4.4

X-ray crystallography

A BRUKER APEX 2 X-ray (three-circle) diffractometer was employed for crystal screening, unit cell determination, and data collection. Integrated intensity information for each reflection was obtained by reduction of data frames with APEX2.70 SADABS was employed to correct the data for absorption effects.71 A solution was obtained readily using SHELXT (XT).72a,b Hydrogen atoms were placed in idealized positions and were set riding on the respective parent atoms. All the non-hydrogen atoms were refined with anisotropic thermal parameters. Absence of additional symmetry and/or voids was confirmed using PLATON. Olex2 was employed for final data presentation and structure plots.73 4.5

DNA binding studies

For absorption spectral studies, DNA was dissolved in a 50 mM NaCl/5 mM Tris-HCl (pH 7.2) solution. The CT DNA solution displayed a UV absorbance ratio (A260/A280) of 1.9 : 1, indicating that the CT DNA was sufficiently in protein free form. The DNA was further diluted in such a way that its maximum absorbance should be 260 nm with the absorption coefficient value of 6600 cm1 M1 per nucleotide.74a,b Solutions of the Ru(II)– benzene complexes were prepared using 5% DMF/Tris-HCl/ NaCl buffer. The absorption titrations were performed by varying the CT DNA concentration (0–50 mM) against the fixed concentration of the complexes (25 mM). As the complexes did not have fluorescence property, the interaction studies were performed indirectly using EB. Fluorescence spectra were recorded at an excitation wavelength of 510 nm in the range of 520–800 nm. The Ru(II)–benzene complexes were added in aliquots (5 mM) to evaluate the changes in the fluorescence spectrum of EB–DNA.75 Viscosity changes in DNA were measured using a MicroUbbelohde viscometer whose temperature was controlled by an external thermostat (25 0.1 1C). The DNA concentration was maintained at 100 mM, while the complex concentration was varied from 0 to 60 mM. Flow time was measured three times for each addition and the average flow time was calculated. The values of relative specific viscosity (Z/Z0)1/3 (Z is the relative viscosity of DNA in the presence of the complex and Z0 is the relative viscosity of DNA alone) were plotted against 1/R (1/R = [complex]/[DNA]). Relative viscosity (Z0) values were calculated from the observed flow time of the DNA solution (t) corrected for the flow time of the buffer alone (t0), using the expression Z0 = (t t0)/t0.76,77 4.6

DNA mobility shift assay

DNA cleavage experiments to illustrate the unwinding of supercoiled DNA were performed using agarose gel electrophoresis.


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To explore the DNA cleavage ability of complexes 1–4, supercoiled (SC) pBR322 DNA (40 mM in base pairs) was incubated at 37 1C with the complexes (50 mM) in a 5% DMF/5 mM Tris-HCl/ 50 mM NaCl buffer at pH 7.2 for 4 h in the absence of any external agent. To predict the DNA cleavage mechanism, pBR322 DNA and 3 were also incubated with reactive oxygen species quenchers (NaN3, 200 mM and DMSO, 2 mL) under

plate. The cells were cultured for 4 h and then the solution in the medium was removed. An aliquot of 100 mL of DMSO was added to the plate which was shaken until the crystals were dissolved.78 The cytotoxicity against cancer cells was determined by measuring the absorbance of the converted dye at 570 nm in an ELISA reader. The cytotoxicity of each sample was expressed as the IC50 value. The percentage of growth inhibition was calculated using the formula

Mean OD of the untreated cells ðcontrolÞ Mean OD of the treated cells 100 Mean OD of the untreated cells ðcontrolÞ identical conditions. Subsequently, 6 DNA loading buffer containing 0.25% bromophenol blue, 0.25% xylene cyanol and 60% glycerol was loaded onto 1.5% agarose gel containing 1.0 mg mL1 of EB. Agarose gel electrophoresis of plasmid DNA was performed at 60 mV for 1 h in TBE buffer (Tris-boric acid/ethylenediaminetetraacetic acid, pH 8) and the bands were visualized under UV light and photographed. The extent of cleavage from supercoiled to its nicked form was assessed by measuring the intensities of the bands using an Alpha Imager instrument. 4.7

Protein binding studies

The binding of Ru(II)–benzene complexes with BSA was studied using fluorescence spectra recorded at a fixed excitation wavelength corresponding to BSA at 280 nm and monitoring the emission at 345 nm. The excitation and emission slit widths and scan rates were constantly maintained for all the experiments. A stock solution of BSA was prepared in 50 mM phosphate buffer (pH = 7.2) and stored in the dark at 4 1C for further use. Concentrated stock solutions of each test compound were prepared by dissolving them in DMF–phosphate buffer (5 : 95) and then diluted with phosphate buffer to get required concentrations. 2.5 ml of BSA solution was titrated by successive additions of a 106 M stock solution of the complexes using a micropipette. For synchronous fluorescence spectra measurements, the same concentrations of BSA and the complexes were used and the spectra were measured at two different Dl (difference between the excitation and emission wavelengths of BSA) values of 15 and 60 nm. 4.8

MTT assay

The cytotoxic property of the Ru(II)–benzene complexes was evaluated against HepG2, A549, MCF7, SKOV3 and Vero cell lines. The cancer cells were maintained in DMEM (Dulbecco’s Modified Eagle’s Medium) with 10% fetal bovine serum and 2 mM L-glutamine, along with antibiotics (100 international units per mL of penicillin, 100 mg mL1 of streptomycin) with the pH adjusted to 7.2. For screening experiments, 100 mL of medium containing 5000 cells/well was incubated at 37 1C for 24 h prior to addition of the complexes. Complexes of different concentrations (3.9 mg mL1 to 250 mg mL1) were seeded in 96 well plates. Triplicate analysis was done and the medium without the complexes served as the control. After 24 h, an aliquot of 100 mL of medium containing 1 mg mL1 of 3-(4,5-dimethylthiazol2-yl)-2,5-diphenyl-tetrazolium bromide (MTT) was loaded into the

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4.9

DNA fragmentation

For DNA fragmentation assay, HepG2 cancer cells (1 106 cells) (control and treated with the Ru(II)–benzene complexes) were collected by centrifugation at 1000 rpm for 5 min and washed twice with phosphate-buffered saline (PBS). The cell pellet was suspended in 100 mL of cell lysis buffer (10 mM Tris-HCl buffer, pH 7.4 containing 10 mM EDTA and 0.5% Triton X-100), kept at 4 1C for 15 min and the cell lysate was centrifuged at 12 000 rpm for 20 min. The supernatants were incubated with proteinase K (0.4 mg mL1) at 58 1C for 60 min, and then incubated with RNase A (0.4 mg mL1) at 37 1C for 60 min. The supernatants were mixed with 20 mL of 5 M NaCl and 120 mL of isopropyl alcohol overnight at 20 1C. The supernatants were then collected by centrifugation at 10 000 rpm for 10 min. DNA samples were dissolved in TE buffer (10 mM Tris-HCl, pH 7.4 and 1 mM EDTA, pH 8.0), and separated by 2% agarose gel electrophoresis.79 4.10

Apoptosis determination by flow cytometry

The ability of the complexes to induce apoptosis was analyzed using an annexin V-FITC/PI detection kit (Biolegend, San Diego).80 HepG2 cells were harvested after treatment and resuspended in binding buffer. Aliquots of 105 cells were mixed with 5 mL each of annexin V-FITC and PI solution for 15 min at room temperature in the dark. After incubation, 400 mL of binding buffer was added, and the number of apoptotic cells was analyzed using a FACS Calibur flow cytometer (Becton Dickinson). 4.11

Confocal microscopy

HepG2 cancer cells with and without the complexes at their IC50 concentration (31.25 mg mL1) were used for confocal imaging. After 24 h of treatment, cells were washed twice with 0.01 M PBS and suspended in binding buffer (ice-cold 1 : 1 methanol : acetone). Cells were incubated with DAPI (40 ,6-diamidino-2-phenylindole), FITC (fluorescein isothiocyanate) and PI (propidium iodide) for 30 min at 4 1C in the dark. Cells were then centrifuged and pellets were smeared. DAPI, FITC and PI fluorescence was immediately observed under a confocal laser scanning microscope (ZEISS, LSM710, Germany).81

Acknowledgements K. J. thanks the Department of Science and Technology, Ministry of Science and Technology, Government of India, for


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doctoral fellowship under the DST-INSPIRE programme. R. K. gratefully acknowledges DST for the financial support.

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