Synthesis of Ru(ii)-benzene complexes containing aroylthiourea ligands, and their binding with biomolecules and in vitro cytotoxicity through apoptosis

Article abstract of DOI:10.1039/c6nj03099h

The reaction of [RuCl₂(η⁶-benzene)]₂ with aroylthiourea resulted in the formation of Ru(ii) complexes of the type [RuCl₂(η⁶-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 μg mL^?1 against HepG2 cells. Complexes 1 and 3 displayed moderate cytotoxicity at the concentration of 62.5 μg mL^?1 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 demonstrated that the complexes have a significant ability to induce cell death by apoptosis.

Full text of DOI:10.1039/c6nj03099h

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Jason B. Benedict et al.
The role of atropisomers on the photo-reactivity and fatigue of
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Synthesis of Ru(II)-benzene complexes containing aroylthiourea ligand, 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
Abstract
The reaction of [RuCl2(η6ꢀbenzene)] with aroylthiourea resulted in the formation of Ru(II)
complexes of the type [RuCl2(η6ꢀbenzene)L] (L = monodentate aroylthiourea ligand). The
complexes have been well characterized using UVꢀVisible, FTꢀIR, NMR and mass spectroscopic
techniques. Single crystal Xꢀray diffraction confirmed the monodentate coordination of the ligand
through sulfur atom. The interaction of the Ru(II) complexes with calf thymus DNA (CT DNA)
was investigated using UVꢀVisible and fluorescence spectroscopic methods, and viscosity
measurements. The binding ability of the complexes with bovine serum albumin (BSA) was
explored using UVꢀVisible and fluorescence experiments. The results showed that the complexes
interact with the biomolecules with appreciable binding constants. Gel electrophoresis technique
has been used to demonstrate the unwinding of the supercoiled DNA to its nicked form. The
cytotoxicity of the Ru(II) complexes has been 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 31.5 µg/mL against HepG2 cells. Complexes 1 and 3 displayed moderate
cytotoxicity at the concentration of 62.5 µg/mL 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 have been carried out. The above methods demonstrated that the
complexes have a significant ability to induce cell death by apoptosis.
______________________________________________________________________________
aDepartment of Chemistry, National Institute of Technology, Tiruchirappalli 620015, India
bDepartment of Hematology, Fujita Health University, 1ꢀ98, Dengakugakubo, Kutsukakeꢀcho,
Toyoake, Aichi 470ꢀ1192, Japan
cDepartment 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

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reported in this paper have been deposited with the Cambridge Crystallographic Data Centre
(CCDC) as supplementary publication numbers CCDC 1477275 and 1477276. Copies of the data
can be obtained free of charge from the CCDC (12 Union Road, Cambridge CB2 1EZ, UK; Tel.:
+ 44ꢀ1223ꢀ336408; Fax: +44ꢀ1223ꢀ336003; eꢀmail: deposit@ccdc.cam.ac.uk; web site:
http://www.ccdc.cam.ac.uk).
1. Introduction
The success of cisplatin as an anticancer drug enticed many researchers to search for the potential
metallodrugs with enhanced pharmacological activities since it suffers from severe side effects
like neuroꢀ, heptoꢀ and nephrotoxicity.1,2 Ruthenium complexes have attracted attention as an
anticancer drug 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 by Clarke’s observation
that [RuCl3(NH3)3] are active anticancer agents.3 Another two important Ru(III) complexes,
transꢀ[RuCl4(DMSO)(Im)]ImH (NAMIꢀA, Im = imidazole)4 and transꢀ[RuCl4(Ind)2]IndH
(KP1019, Ind = indazole),5 are under clinical trial. It was found that KP1019 is cytotoxic to
cancer cells, NAMIꢀA is relatively nonꢀtoxic 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 the in vivo reduction to more labile Ru(II) complexes. Therefore the “activation by
reduction” strategy has induced the interest towards the anticancer activity of Ru(II) complexes.
Though many Ru(II) organometallic compounds were reported as anticancer agents, halfꢀ
sandwich RuCl2(arene) type complexes received a special attention. Because, the inert
hydrophobic arene moiety, 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 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 a great promise towards the development of
anticancer drugs.8ꢀ27 The first complex of this kind, which was evaluated for cytotoxic property
was [Ru(η6ꢀbenzene)Cl2(metronidazole)] (metronidazole
=
1ꢀβꢀhydroxyethylꢀ2ꢀmethylꢀ5ꢀ
nitroimidazole) (Fig. 1). The another important class of Ru(II)ꢀarene complexes bearing pta
ligand, [RuCl2(η6ꢀbenzene)(pta)] (RAPTAꢀB) and [RuCl2(η6ꢀpꢀcymene)(pta)] (RAPTAꢀC) (pta =

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1,3,5ꢀtriazaꢀ7ꢀphosphaadamantane) have recently attracted considerable attention due to the
promising in vivo activity on the inhibition of metastasis growth, together with a high selectivity
and low general toxicity (Fig. 1).28 Interestingly, [RuCl2(η6ꢀbenzene)(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. 30ꢀ34 Herein
we describe the synthesis, structure, DNA and protein binding, DNA cleavage, in vitro
cytotoxicity, DNA fragmentation and confocal imaging of Ruꢀbenzene complexes containing
aroylthiourea ligand. The complexes we report are similar to the RAPTAꢀB backbone, pta ligand
being replaced by aroylthiourea. Moreover there are only a few Ruꢀbenzene complexes reported
for their anticancer activity and this is the first report on Ruꢀbenzene complexes containing
aroylthiourea ligands evaluated for their biological applications.
Fig. 1 Structure of (a) [Ru(η6ꢀbenzene)Cl2(metronidazole)] and (b) RAPTAꢀB.
2. Results and discussion
2.1 Synthesis and spectroscopy
The aroylthiourea ligands (L1ꢀL4) used in the work have been synthesized and reported by us
earlier.35 The reactions of the ligands with Ruꢀbenzene dimer produced [RuCl2(η6ꢀbenzene)L] (1ꢀ
4) (Scheme 1).

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R
Ligand
L1
Complex
1
L2
2
L3
L4
3
4
Scheme 1. Synthetic route for the ligands and their Ru(II)ꢀbenzene complexes
The Ru(II) complexes have been characterized by UVꢀVisible, FTꢀIR, mass and NMR
spectroscopic techniques. The UVꢀVisible spectra of the complexes showed new bands around
349ꢀ364 and 444ꢀ461 nm, which correspond to MLCT and d→d transitions respectively. The FTꢀ
IR spectra of the ligands showed bands in the region 3076ꢀ3312, 1661ꢀ1657 and 1256ꢀ1271 cmꢀ1

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which were assigned to ν(N−H), ν(C=O) and ν(C=S) respectively. The N−H and C=O bands
were unaltered in the spectra of the complexes while ν(C=S) was decreased (1171ꢀ1201 cmꢀ1)
which confirmed the coordination of only sulfur to Ru ion.
1
In H NMR spectra of the complexes the carbonyl and thiocarbonyl attached N−H, and
thiocarbonyl attached N−H protons resonated around 11.49ꢀ12.54 and 11.09ꢀ11.82 ppm
respectively. The signal observed at 5.98 ppm in the spectra of all the complexes confirmed the
presence of benzene moiety (Figs. S1ꢀS4). All the other aromatic protons displayed signals in the
expected region (8.46ꢀ7.27 ppm). In 13C NMR spectra of the complexes, the signals appeared
around 122.7ꢀ138.4 ppm corresponded to the aromatic carbons in the complexes. The
thiocarbonyl and carbonyl carbons were characterized by the signals around 179.2ꢀ181.2 and
162.4ꢀ162.7 ppm respectively. A new resonance appeared in the spectra of the complexes at 88.1
ppm was assigned to benzene (Figs. S5ꢀS8). 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 Xꢀray diffraction were obtained by the slow evaporation of chloroform/methanol solution of
the complexes. Complexes 1 and 3 crystallized in monoclinic P21/c and triclinic Pꢀ1 space group
respectively. The complexes featured a halfꢀsandwich three legged piano stool structure about the
metal centre (Figs. 2 and 3). It was clear from the crystal structures that the complexes adopted a
‘pianoꢀstool’ geometrical configuration with the πꢀbonded η6ꢀbenzene, monodentate aroylthiourea
and two chloride ions. The Ru−C distances were in the range of 2.1686ꢀ2.197 Å with the average
of 2.1781 (1) and 2.1806 (3) Å.36aꢀc The Ru−Cl and Ru−S bond lengths were found to be in the
range of 2.4149ꢀ2.4350 and 2.407ꢀ2.414 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 H···O distance of 2.00 Å in 3.

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Table 1 Crystal data and structure refinement for complexes 1 and 3
1
3
Empirical formula
Formula weight
Temperature (K)
Wavelength (Å)
Crystal system
Space group
Unit cell dimensions
a (Å)
C18H16Cl2N2ORuS2
512.42
C19H18Cl2N2ORuS2
526.44
150
110.15
0.71073
0.71073
Monoclinic
P21/c
Triclinic
Pꢀ1
11.674(2)
10.818(2
15.646(3)
90
6.5098(19)
11.428(3)
14.534(4)
101.993(3)
95.432(4)
105.472(3)
1006.3(5)
2
b (Å)
c (Å)
α
β
γ
(°)
(°)
(°)
99.37(3)
90
Volume (Å3)
1949.8(7)
4
Z
Density (calculated) (Mg/m3)
Absorption coefficient (mmꢀ1)
F(000)
1.746
1.737
1.302
1.264
1024
528
Crystal size (mm3)
0.32×0.315×0.106
0.15×0.04×0.03
Theta range for data collection (°) 1.768 to 27.540
ꢀ15<=h<=15,
1.451 to 27.550
ꢀ8<=h<=8,
ꢀ14<=k<=13,
ꢀ20<=l<=20
ꢀ14<=k<=14,
ꢀ18<=l<=18
11564
Index ranges
Reflections collected
21897
4445 [R(int) = 0.0321]
4551 [R(int) = 0.0466]
Independent reflections [R(int)]
Completeness to theta =
99.7
99.7

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25.242/27.50°
Semiꢀempirical from
equivalents
Semiꢀempirical from
Absorption correction
equivalents
0.7456 and 0.5274
Max. and min. transmission
Refinement method
0.7456 and 0.6105
Fullꢀmatrix leastꢀsquares
on F2
Fullꢀmatrix leastꢀsquares
on F2
Data / restraints / parameters
Goodnessꢀofꢀfit on F2
4445 / 0 / 235
1.037
4551 / 0 / 244
1.043
R1 = 0.0204,
wR2 = 0.0490
R1 = 0.0240,
wR2 = 0.0509
R1 = 0.0399,
wR2 = 0.1051
R1 = 0.0441,
wR2 = 0.1089
1.867 and ꢀ1.652
Final R indices [I>2sigma(I)]
R indices (all data)
Largest diff. peak and hole (e.Åꢀ3) 0.430 and ꢀ0.368
Table 2 Selected bond lengths (Å) and angles (°) of complexes 1 and 3
Bond length/angle(Å)
Ru−arene (centroid)a
Ru(1)−Cl(1)
1
3
1.658
1.665
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)
2.4350(9)
2.4149(10)
2.4139(8)
2.190(3)
2.173(3)
2.197(3)
2.182(3)
2.185(3)
2.171(3)
87.15(3)
88.99(3)
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)

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S(1)−Ru(1)−Cl(1)
88.99(2)
92.63(3)
Fig. 2 Thermal ellipsoid (50 %) plot of 1.
Fig. 3 Thermal ellipsoid (50 %) plot of 3.

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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
depend on the solubility of the compound in aqueous versus organic medium. The log P values of
L1ꢀL4 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 absorbance of the
complexes on adding aliquots of CT DNA have been monitored. Usually intercalative mode of
binding of the complexes with DNA shows hypochromism with or without red shift40a,b. Nonꢀ
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 occured in the
intraligand band (23ꢀ40 %) with 2ꢀ3 nm red shift (Figs. 4 and S9), which indicated intercalative
mode of binding.42ꢀ44 To compare the DNA binding ability of the complexes, equilibrium binding
constant (Kb) was calculated using the equation45 [DNA]/(εa−εf) = [DNA]/(εb−εf) + 1/Kb (εb−εf)
where [DNA] is the concentration of DNA in base pairs, εa is the apparent extinction coefficient
value found by calculating A(observed)/[complex], εf is the extinction coefficient for the free
compound and εb is the extinction coefficient for the compound in the fully bound form. A plot of
[DNA]/(εa−εf) versus [DNA] gave Kb as the ratio of slope to the intercept (Fig. 5). The calculated
intrinsic equilibrium binding constants (Kb) for 1ꢀ4 were in the range of 1.17×104ꢀ7.30×104 Mꢀ1
respectively (Table 3).The binding affinity of the complexes followed the order 4 > 3 > 2 > 1. The
Kb value of 1ꢀ4 was comparable with the reported Kb of similar rutheniumꢀarene complexes.
42,46,47

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Fig. 4 Absorption spectra of complex 4 in TrisꢀHCl buffer upon addition of CT DNA. [Complex]
= 2.5 × 10−5 M, [DNA] = 0ꢀ50 ꢁM. The arrow shows that the absorption intensity decreases upon
increasing the CT DNA concentration.
Fig. 5 Plot of [DNA]/(εa−εf ) versus [DNA] for the titration of the complexes with CT DNA.

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Table 3 DNA binding constant (Kb), quenching constant (Kq) and apparent binding constant
(Kapp) values
Complex Kb (M-1)
Kq (M-1)
Kapp (M-1)
1.17×104±0.15
3.54×104±0.14
3.65×104±0.11
3.77×104±0.11
3.91×104±0.07
1.77×106±0.09
1.82×106±0.11
1.88×106±0.16
1.95×106±0.13
1
2
3
4
1.41×104 ±0.09
1.62×104±0.13
7.30×104±0.24
The competitive ethidium bromide (EB) binding studies provided support for DNA
interaction of the complexes.48ꢀ50 Upon addition of the complexes (1-4) (0ꢀ50 ꢁM) to CT DNA
pretreated with EB in 5 % DMF/5 mM TrisꢀHCl/50 mM NaCl buffer at pH 7.2, the emission
intensity of DNAꢀbound EB at 605 nm decreases (Figs. 6 and S10). The percentage of quenching
for complexes 1ꢀ4 was found to be 63.4, 64.1, 64.6 and 65.3 % respectively with 10ꢀ12 nm
bathochromic shift. The extent of quenching revealed the extent of displacement of the EB
molecules from the CT DNA. According to SternꢀVolmer equation, relative fluorescence is
directly proportional to the concentration of the quencher. The SternꢀVolmer equation is given by
Fo/F = 1 + Kq [Q] where Fο and F are the fluorescence intensities in the absence and presence of
complex respectively, Kq is a linear SternꢀVolmer quenching constant, and [Q] is the
concentration of complex. The slope of the plot of Fo/F versus [Q] gave Kq (Fig. 7). From a plot
of the observed intensities against the complex concentration, the values of the apparent DNA
binding constant (Kapp) was calculated using the equation51 KEB [EB] = Kapp [complex] where
[complex] is the complex concentration at 50 % reduction in the fluorescence intensity of EB, KEB
= 1.0×107 Mꢀ1 and [EB] = 5 µM. Kq and Kapp values are listed in Table 3. The interaction of the
complexes with CT DNA followed the order 4 > 3 > 2 > 1. The results are in agreement with
those found from the electronic absorption studies.

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Fig. 6 Fluorescence quenching curves of EB bound to DNA in the presence of 4. [DNA] = 5 µM,
[EB] = 5 µM and [complex] = 0ꢀ50 µM.
Fig. 7 SternꢀVolmer plot of fluorescence titrations of the complexes with CT DNA.

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Viscosity measurements illustrated the effect of the complexes on the specific relative
viscosity of DNA. Since the specific relative viscosity (η/η0; η and η0 are the specific viscosities
of DNA in the presence and absence of the complexes, respectively) will have an effect when
DNA contour length changes, it is helpful in the determination of the mode of interaction of the
complexes with DNA. The increase in contour length associated with the separation of DNA base
pairs caused by intercalation, a classical DNA intercalator like EB shows a significant increase in
the viscosity of the DNA solutions. In contrast, a partial and/or nonꢀintercalation of the complex
could result in a less pronounced effect on the viscosity.52 The viscosity of DNA was increased on
incremental addition of the complexes, which showed the intercalative mode of binding (Fig. 8).
Complex 4 showed comparatively increased effect in the viscosity among the complexes
investigated. Complex 1 did not show a steady increase in viscosity on the addition of DNA. The
effect of viscosity on addition of the complexes followed the order 4 > 3 > 2 > 1, which was in
good agreement with the results obtained from the spectroscopic studies.
Fig. 8 Effect of increasing amounts of complexes 1ꢀ4 on the relative viscosity of CT DNA.
2.5 Unwinding of supercoiled DNA
Change in the electrophoretic mobility of plasmid DNA on agarose gel is commonly taken as
evidence for DNAꢀdrug interactions. The change in the DNA structure from supercoiled form to
its nicked or linear form triggers 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 ꢁM in

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base pairs) was incubated at 37 °C with the complexes (50 µM) in a 5 % DMF/5 mM TrisꢀHCl/50
mM NaCl buffer at pH 7.2 for 4 h in the absence of any external agent. Fig. 9 illustrates the gel
electrophoretic separations showing the cleavage of plasmid pBR322 DNA by the Ru(II)
complexes under identical conditions. All the complexes investigated cleaved supercoiled (SC)
DNA (Form I) into nicked circular (NC) DNA (Form II), and the DNA cleavage followed the
order 3 (48.45 %) > 4 (44.53 %) > 1 (34.46 %) ≈ 2 (33.32 %).
Fig. 9 Cleavage of supercoiled pBR322 DNA (40 ꢁM) by complexes 1ꢀ4 in a buffer
containing 5 % DMF/5 mM Tris HCl/50 mM NaCl at pH = 7.2 and 37 °C with an
incubation time of 4 h. Lane 1, DNA; lane 2, DNA + 1 (50 ꢁM); lane 3, DNA + 2 (50 ꢁM);
lane 4, DNA + 3 (50 ꢁM); lane 5, DNA + 4 (50 ꢁM). Forms SC and NC are supercoiled and
nicked circular DNA, respectively.
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 is no reduction in the percentage of cleavage even in the
presence of scavengers (Fig. S11), which ruled out the oxidative and photoꢀinduced cleavage
mechanisms. Therefore the DNA cleavage might occurred through hydrolytic pathway as
observed in Ruꢀpolypyridyl complexes.53b
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

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Fig. 10 Absorption spectra of BSA (10 µM) and BSA with 1ꢀ4 (4 µM).
The quenching experiments have been 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 2ꢀ3 nm hypsochromic shift (Figs. 11 and
S12).56 The extent of quenching of fluorescence intensity is expressed by SternꢀVolmer constant
(Kq) which is a measure of protein binding affinity of the complexes.57 The value of Kq obtained
as slope of the linear plot of Fo/F versus [complex] followed the order 2 > 4 > 1 > 3 (Fig. 12).
Further, the equilibrium binding constant was evaluated using the Scatchard equation,
log[(Fo−F)/F] = log KBSA + n log[Q] where F and Fo are the fluorescence intensity in the presence
and absence of the complexes, KBSA is the binding constant and n is the number of binding site.
The KBSA values were derived from the plot between log[(Fo−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).

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Fig. 11 Fluorescence quenching curves of BSA in the absence and presence of 2. [BSA] = 1 µM
and [complex] = 0ꢀ20 µM.
Fig. 12 SternꢀVolmer plot of the fluorescence titrations of the complexes with BSA.

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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
KBSA (M-1)
Kq (M-1)
n
1.66×105±0.18
8.63×105±0.14
1.16×105±0.11
1.74×105±0.15
1.67×105±0.08
2.44×105±0.10
1.20×105±0.13
2.26×105±0.11
1.03
1.33
0.97
0.99
1
2
3
4
Vekshin suggested an useful method known as synchronous fluorescence spectroscopy
which gives valuable information on 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 (ꢂλ = λem−λex) indicates the types of
chromophores. A higher ꢂλ value, such as 60 nm, is indicative of the tryptophan residue, whereas
a lower ꢂλ value, such as 15 nm, is characteristic of the tyrosine residue.60 In the synchronous
fluorescence spectra of BSA at ꢂλ = 15 nm, the incremental addition of the complexes to BSA
resulted in the quenching of the fluorescence intensity at 304 nm with 61.6, 74.8, 57.0 and 66.8 %
for complexes 1ꢀ4 respectively (Figs. 14 and S13). Similarly there was also quenching observed

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in the synchronous fluorescence spectra of BSA (348 nm) at ꢂλ = 60 nm, with 74.4, 84.6, 71.0
and 79.7 % for complexes 1ꢀ4 respectively (Figs. 15 and S14). The 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.
Fig. 14 Synchronous spectra of BSA (1 µM) as a function of concentration of 2 (0ꢀ20 µM) with
ꢂλ = 15 nm.
Fig. 15 Synchronous spectra of BSA (1 µM) as a function of concentration of 2 (0ꢀ20 µM) with
ꢂλ = 60 nm.

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2.7 Cytotoxicity
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 wellꢀknown anticancer
drug cisplatin. Complex solutions of all concentrations used in the experiment decreased the cell
viability significantly in a concentrationꢀdependent manner. The graph of percentage of cell death
versus concentration is shown in Figs. 16, 17, S15 and S16. The half minimum inhibitory
concentration (IC50) calculated for the complexes is tabulated (Table 5). From the results it was
inferred that the complexes 1, 2 and 3 showed modest activity towards HepG2 at lower
concentration (31.5 ꢁg/mL) with IC50 ranging from 40.80 to 53.12 ꢁM. Complex 4 showed only
least activity towards HepG2 cell growth with IC50 value of 162.20 ꢁM. Among the investigated
complexes, only complex 1 showed moderate toxicity towards A549 and SKOV3 cell lines with
IC50 values of 95.60 and 87.52 ꢁM respectively. The complexes were less toxic towards A549,
MCF7 and SKOV3 cancer cell lines and Vero normal cell line. The cytotoxicity of the
investigated complexes was reasonably low compared to that of similar pꢀcymene complexes.35
Cytotoxicity of 1ꢀ4 was compared with that of other reported rutheniumꢀbenzene complexes. The
cytotoxicity of 1ꢀ4 is superior to that exhibited by ruthenium(II)(η6ꢀNꢀbenzylmaleimide)
containing 1,3,5ꢀtriazaꢀ7ꢀphosphaadamantane (PTA) (IC50, 640 µM). Triphenylphosphine
complex of Ruꢀbenzene showed a comparable activity (IC50, 116 µM) against A549 cancer cell
line (Fig. 18).61 Rutheniumꢀbenzene complex bearing 4ꢀ(biphenylꢀ4ꢀcarbonyl)ꢀ3ꢀmethylꢀ1ꢀ
phenylꢀ5ꢀpyrazolonate ligand showed similar halfꢀinhibitory concentration (35.0 µM) against
HepG2 cancer cell line.62 The comparison with the potential drug cisplatin showed that our
complexes were less effective. Compounds 1ꢀ4 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 on specific
activity of the complexes against various cancer cell lines.

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Fig. 16 Comparison of anticancer activity of the complexes (1ꢀ4) 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 (1ꢀ4) against A549 cancer cells. Data
are mean ± SD of three independent experiments with each experiment conducted in triplicate.

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Fig. 18 IC50 value of reported rutheniumꢀbenzene complexes.
Table 5 In vitro cytotoxicity of the complexes in HepG2, A549, MCF7 and SKOV3 cancer cell
lines
IC50 (µM)
Complex
SKOV3
>250
HepG2
53.14
50.44
40.80
162.20
21.5
A549
95.60
>250
>250
>250
18.0
MCF7
151.29
153.93
162.99
154.40
12.0
1
165.90
87.52
>250
2
3
4
8.3
Cisplatin

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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 necrosis
(unprogrammed cell death) leads to serious health issues.63ꢀ65 This makes apoptosis distinct from
the other form of cell death. Therefore, fluorescent staining [DAPI (blue), FITC (green) and PI
(red) fluorescence] to assess the morphological changes and gel electrophoresis for detecting
DNA fragmentation have been carried out. DNA fragmentation was demonstrated by incubating
HepG2 cells with IC50 concentrations (31.25 µg/mL) of 1, 2 and 3 for 24 h. DNA fragmentation
became apparent and 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
demonstrate that the cell death mechanism is through apoptosis.
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 1ꢀ3 in IC50 concentration (31.25 µg/mL) for 24 h. Lane 1, 1 kb DNA ladder; lane 2,
control; lane 3, 1; lane 4, 2; lane 5, 3. Arrow marks indicate the DNA fragmentation.

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The 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), lower right quadrant (R3), upper right quadrant (R4) and upper left
quadrant (R5) contained the living cells, early apoptosis cells, late apoptosis cells and dead cells
respectively. To analyze the apoptosis, HepG2 cell lines were treated with IC50 concentration of
the complexes (1ꢀ3) for 24 h. There was a dramatic increase in the apoptosis effect on HepG2
cells when compared to the control cells (Fig. 20). Complexes 1ꢀ3 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
Fig. 20 Flow cytometry analysis carried out in HepGꢀ2 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.

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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 ꢁg/mL) 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 AꢀT rich regions in DNA and pass 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 image showed that
significant morphological changes were found in HepG2 cells after treatment with the complexes
and apoptotic cells were indicated with arrows in Fig. 21.

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a1
b1
c1
d1
a2
b2
c2
d2
a3
a4
b4
c4
d4
b3
c3
d3
Fig. 21 DAPI (blue), FITC (green) and PI (red) fluorescence staining for the detection of
apoptosis in HepG2 cells. Control: (a1ꢀDAPI, a2ꢀFITC, a3ꢀPI and a4ꢀmerged) and treated (1: b1ꢀ
DAPI, b2ꢀFITC, b3ꢀPI and b4ꢀmerged); (2: c1ꢀDAPI, c2ꢀFITC, c3ꢀPI and c4ꢀmerged); (3: d1ꢀ
DAPI, d2ꢀFITC, d3ꢀPI and d4ꢀmerged). Arrows indicate apoptotic cancer cells.

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3. Conclusion
The present work describes the synthesis and characterization of halfꢀsandwich 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 Mꢀ1. The complexes also cleaved DNA through hydrolytic pathway. The protein binding
efficacy of the complexes has also been demonstrated using UVꢀVisible and fluorescence
spectroscopic techniques. In vitro cytotoxic assay revealed that the complexes (1ꢀ3) were active
only against HepG2 among the four tested cancer cell lines and the results were compared with
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 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 Lab India instrument and are uncorrected. FTꢀIR spectra
were obtained as KBr pellets using a NicoletꢀiS5 spectrophotometer. UVꢀVisible spectra were
recorded using a Shimadzuꢀ2600 spectrophotometer. Emission spectra were measured on a Jasco
Vꢀ630 spectrophotometer using 5 % DMF in buffer as the solvent. NMR spectra were recorded in
DMSOꢀd6 solvent by using TMS as an internal standard on a Bruker 500 MHz spectrometer. ESIꢀ
MS spectra were recorded using a Thermo Exactive Orbitrap mass spectrometer. The lipophilicity
of the ligands was measured by the “shake flask” method in octanolꢀwater phase partitions as
reported earlier.39
4.2 Synthesis of the ligands
Ligands (L1ꢀL4) were prepared from thiopheneꢀ2ꢀcarbonylchloride, potassium thiocyanate and
primary amine (aniline, oꢀtoluidine, benzylamine or 1ꢀnaphthylamine) by following a reported
procedure.32
4.3 Synthesis of the Ru(II)-benzene complexes
The rutheniumꢀbenzene [RuCl2(η6ꢀbenzene)]2 precursor was synthesized using earlier reported
procedure.69 [RuCl2(η6ꢀbenzene)]2 (0.1000 g, 0.2 mmol) and ligand (L) were dissolved in 15 mL
of toluene and stirred for 14ꢀ20 h at 27 °C. The resultant solution was concentrated to 2 mL under

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reduced pressure, and addition of petroleum ether (60ꢀ80 °C) (10 mL) gave an orange solid. The
product was collected by filtration, washed with petroleum ether and dried in vacuo.
4.3.1 [RuCl2(η6-benzene)L1] (1)
L1 (0.1049 g, 0.4 mmol) was used. Yield: 79 %. Orange solid. M.p.: 215 °C. 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. ESIꢀMS (m/z): (M−2H+−2Clꢀ+H+)+ found (calcd) 440.9666 (440.9669). UVꢀVis (DMF):
λ
max, nm (ε, dm3molꢀ1 cmꢀ1) 265 (22533), 350 (9666), 444 (1666). FTꢀIR (KBr, cm−1): 3213 (m;
1
ν(amide N−H)), 3115 (s; ν(thiourea N−H)), 1655 (s; ν(C=O)), 1176 (s; ν(C=S)). H NMR (500
MHz, DMSOꢀd6) δ, 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, DMSOꢀd6) δ, ppm 179.2 (C=S), 162.5 (C=O), 138.4, 137.1,
135.8, 133.2, 129.2, 129.1, 128.8, 126.8, 124.8 (aromatic carbons of thiourea ligand), 88.1
(aromatic carbons of benzene).
4.3.2 [RuCl2(η6-benzene)L2] (2)
L2 (0.1105 g, 0.4 mmol) was used. Yield: 81 %. Orange solid. M.p.: 203 °C. 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. ESIꢀMS (m/z): (M−2H+−2Clꢀ+H+)+ found (calcd) 454.9820 (454.9826). UVꢀVis (DMF):
λ
max, nm (ε, dm3molꢀ1cmꢀ1) 265 (31600), 290 (34066), 349 (15800), 461 (1966). FTꢀIR (KBr,
cm−1): 3220 (m; ν(amide N−H)), 3162 (s; ν(thiourea N−H)), 1654 (s; ν(C=O)), 1171 (s; ν(C=S)).
1H NMR (500 MHz, DMSOꢀd6) δ, 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), 7.28ꢀ7.22 (m, 3H),
13
5.98 (s, 6H, benzene), 2.26 (s, 3H). C NMR (125 MHz, DMSOꢀd6) δ, ppm 180.2 (C=S), 162.6
(C=O), 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 thiourea ligand), 88.1 (aromatic carbons of benzene), 18.1 (aliphatic carbon).
4.3.3 [RuCl2(η6-benzene)L3] (3)
L3 (0.1105 g, 0.4 mmol) was used. Yield: 82 %. Orange solid. M.p.: 230 °C. 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. ESIꢀMS (m/z): (M−2H+−2Clꢀ+H+)+ found (calcd) 454.9821 (454.9826). UVꢀVis (DMF):
λ
max, nm (ε, dm3molꢀ1 cmꢀ1) 265 (17100), 286 (15900), 349 (4166), 448 (933). FTꢀIR (KBr, cm−1):
1
3245 (m; ν(amide N−H)), 3161 (s; ν(thiourea N−H)), 1651 (s; ν(C=O)), 1191 (s; ν(C=S)). H
NMR (500 MHz, DMSOꢀd6) δ, ppm 11.49 (s, 1H), 11.09 (s, 1H), 8.36 (d, J = 2.9 Hz, 1H), 8.02

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(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),
13
5.98 (s, 6H, benzene), 4.87 (d, J = 5.3 Hz, 2H). C NMR (125 MHz, DMSOꢀd6) δ, ppm 180.6
(C=S), 162.4 (C=O), 137.7, 137.2, 135.5, 132.9, 129.1, 128.9, 128.0, 127.7 (aromatic carbons of
thiourea ligand), 88.1 (aromatic carbons of benzene), 48.6 (aliphatic carbon).
4.3.4 [RuCl2(η6-benzene)L4] (4)
L4 (0.1249 g, 0.4 mmol) was used. Yield: 80 %. Orange solid. M.p.: 223 °C. 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. ESIꢀMS (m/z): (M−2H+−2Clꢀ+H+)+ found (calcd) 490.9819 (490.982 6). UVꢀVis (DMF):
λ
max, nm (ε, dm3molꢀ1 cmꢀ1) 265 (21500), 293 (21433), 364 (7766), 454 (1466). FTꢀIR (KBr,
cm−1): 3209 (m; ν(amide N−H)), 3143 (s; ν(thiourea N−H)), 1656 (s; ν(C=O)), 1201 (s; ν(C=S)).
1H NMR (500 MHz, DMSOꢀd6) δ, 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 =
13
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). C NMR
(125 MHz, DMSOꢀd6) δ, ppm 181.2 (C=S), 162.7 (C=O), 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 thiourea
ligand), 88.1 (aromatic carbons of benzene).
4.4 X-ray crystallography
A BRUKER APEX 2 Xꢀray (threeꢀcircle) 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
72a,b
for absorption effects.71 A solution was obtained readily using SHELXT (XT).
Hydrogen
atoms were placed in idealized positions and were set riding on the respective parent atoms. All
the nonꢀhydrogen atoms were refined with anisotropic thermal parameters. Absence of additional
symmetry and / or voids were 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 TrisꢀHCl (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 cmꢀ1Mꢀ1 per nucleotide.74a,b Solutions of the Ru(II)ꢀbenzene complexes were prepared

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using 5 % DMF/TrisꢀHCl/NaCl buffer. The absorption titrations were performed by varying the
CT DNA concentration (0ꢀ50 µM) against the fixed concentration of the complexes (25 µM).
As the complexes did not have fluorescence property, the interaction studies have been
performed indirectly using EB. Fluorescence spectra were recorded at the excitation wavelength
of 510 nm in the range 520ꢀ800 nm. The Ru(II)ꢀbenzene complexes were added in aliquots (5
µM) to evaluate the changes in the fluorescence spectrum of EBꢀDNA.75
Viscosity changes in DNA was measured using MicroꢀUbbelohde viscometer whose
temperature was controlled by an external thermostat (25 ± 0.1 °C). The DNA concentration was
maintained at 100 ꢁM, while the complex concentration was varied from 0 to 60 ꢁM. Flow time
was measured three times for each addition and the average flow time was calculated. The values
of relative specific viscosity (η/η0)1/3 (η is the relative viscosity of DNA in the presence of the
complex and η0 is the relative viscosity of DNA alone) were plotted against 1/R (1/R =
[complex]/[DNA]). Relative viscosity (η0) 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 η0 =
(t − t0)/t0.76,77
4.6 DNA mobility shift assay
DNA cleavage experiments to illustrate the unwinding of supercoiled DNA was performed using
agarose gel electrophoresis. To explore the DNA cleavage ability of complexes 1ꢀ4, supercoiled
(SC) pBR322 DNA (40 ꢁM in base pairs) was incubated at 37 °C with the complexes (50 µM) in
a 5% DMF/5 mM TrisꢀHCl/50 mM NaCl buffer at pH 7.2 for 4 h in the absence of any external
agent. To predict DNA cleavage mechanism, pBR322 DNA and 3 were also incubated with
reactive oxygen species quenchers (NaN3, 200 ꢁM and DMSO, 2 ꢁL) under identical conditions.
Subsequently, 6X 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 ꢁg mL−1 of EB.
Agarose gel electrophoresis of plasmid DNA was performed at 60 mV for 1 h in TBE buffer
(Trisꢀboric 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 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