Solvent-assisted formation of ruthenium(II)/copper(I) complexes containing thiourea derivatives: Synthesis, crystal structure, density functional theory, enzyme mimetics and in vitro biological perspectives

Article abstract of DOI:10.1002/aoc.3652

N,N-[(diethylamino)(thiocarbonyl)]-substituted benzamidine ligands have been synthesized from the reaction of N,N-[(diethylamino)(thiocarbonyl)]benzimidoyl chloride with functionalized amines such as 2-aminophenol and 2-picolylamine. The reaction of N,N-[(diethylamino)(thiocarbonyl)]-2-hydroxyphenylbenzamidine (H₂L₁) with ruthenium(II) precursor [RuHCl(CO)(PPh₃)₃] afforded complex 1 of the type [Ru(L₁)(CO)(PPh₃)₂] in which the ligand coordinated in tridentate ONS mode. The reaction of H₂L₁ with copper precursor [Cu(CH₃COO)(PPh₃)₂] induced C═N bond cleavage of the ligand and afforded complex 3 of the type [Cu(1,1-DT)(Cl)(PPh₃)₂] (1,1-DT?=?1,1-diethylthiourea) in which the ligand coordinated in a monodentate fashion. The ligand N,N-[(diethylamino)(thiocarbonyl)]-2-picolylbenzamidine (HL₂) reacted with ruthenium(II) and copper(I) precursors to form complex 2 of the type [Ru(1,1-DT)(Cl₂)(CO)(PPh₃)₂] and complex 3, respectively, in which the ligand underwent C═N cleavage and coordinated in a monodentate fashion via C═S group. In complexes 1 and 2, the two triphenylphosphine co-ligands coordinated in trans position whereas, in complex 3, the two triphenylphosphine co-ligands coordinated in cis position. All the compounds were characterized using infrared, UV–visible, (¹H, ¹³C, ³¹P) NMR, ESI-MS and elemental analyses. The molecular structures of ligand H₂L₁ and complexes 1–3 were determined using X-ray crystallography, which confirmed the coordination mode of the ligands with metals. The crystal structure of complexes 1 and 2 revealed a distorted octahedral geometry around the ruthenium ion and the structure of complex 3 indicated a tetrahedral geometry around the copper ion. With the X-ray structures, density functional theory computations were carried out to determine the electronic structure of the compounds. The interactions of complexes 1–3 with calf thymus DNA and bovine serum albumin protein were investigated using UV–visible and fluorescence spectroscopic and viscometric methods. Catecholase- and phosphatase-like activities promoted by complexes 1–3 under physiological conditions have been studied. In vitro anticancer activities have been demonstrated by MTT assay, acridine orange/ethidium bromide and diamidino-2-phenylindole staining against various cancerous cell lines.

Full text of DOI:10.1002/aoc.3652

Received: 10 June 2016
DOI 10.1002/aoc.3652
Revised: 23 August 2016
Accepted: 23 August 2016
F U L L PA P E R
Solvent‐assisted formation of ruthenium(II)/copper(I) complexes
containing thiourea derivatives: Synthesis, crystal structure,
density functional theory, enzyme mimetics and in vitro biological
perspectives
Paranthaman Vijayan1 | Periasamy Viswanathamurthi1 | Paramasivam Sugumar2 |
Mondikalipudur Nanjappagounder Ponnuswamy2 | Jan Grzegorz Malecki3 |
Krishnaswamy Velmurugan4 | Raju Nandhakumar4 | Manickam
Dakshinamoorthi Balakumaran5 | Pudupalayam Thangavelu Kalaichelvan5
1 Department of Chemistry, Periyar University,
N,N‐[(diethylamino)(thiocarbonyl)]‐substituted benzamidine ligands have been
synthesized from the reaction of N,N‐[(diethylamino)(thiocarbonyl)]benzimidoyl
chloride with functionalized amines such as 2‐aminophenol and 2‐picolylamine.
The reaction of N,N‐[(diethylamino)(thiocarbonyl)]‐2‐hydroxyphenylbenzamidine
Salem 636 011, India
2 Centre of Advanced Studies in Crystallography
and Biophysics, University of Madras, Guindy
Campus, Chennai 600 025, India
3 Department of Crystallography, Silesian
(H2L1) with ruthenium(II) precursor [RuHCl(CO)(PPh3)3] afforded complex 1 of
University, Szkolna 9, 40‐006 Katowice, Poland
the type [Ru(L1)(CO)(PPh3)2] in which the ligand coordinated in tridentate ONS
mode. The reaction of H2L1 with copper precursor [Cu(CH3COO)(PPh3)2] induced
C═N bond cleavage of the ligand and afforded complex 3 of the type [Cu(1,1‐DT)
(Cl)(PPh3)2] (1,1‐DT = 1,1‐diethylthiourea) in which the ligand coordinated in a
monodentate fashion. The ligand N,N‐[(diethylamino)(thiocarbonyl)]‐2‐
picolylbenzamidine (HL2) reacted with ruthenium(II) and copper(I) precursors to
form complex 2 of the type [Ru(1,1‐DT)(Cl2)(CO)(PPh3)2] and complex 3, respec-
tively, in which the ligand underwent C═N cleavage and coordinated in a
monodentate fashion via C═S group. In complexes 1 and 2, the two
triphenylphosphine co‐ligands coordinated in trans position whereas, in complex
3, the two triphenylphosphine co‐ligands coordinated in cis position. All the com-
pounds were characterized using infrared, UV–visible, (1H, 13C, 31P) NMR, ESI‐
MS and elemental analyses. The molecular structures of ligand H2L1 and complexes
1–3 were determined using X‐ray crystallography, which confirmed the coordination
mode of the ligands with metals. The crystal structure of complexes 1 and 2 revealed
a distorted octahedral geometry around the ruthenium ion and the structure of com-
plex 3 indicated a tetrahedral geometry around the copper ion. With the X‐ray struc-
tures, density functional theory computations were carried out to determine the
electronic structure of the compounds. The interactions of complexes 1–3 with calf
thymus DNA and bovine serum albumin protein were investigated using UV–visible
and fluorescence spectroscopic and viscometric methods. Catecholase‐ and phos-
phatase‐like activities promoted by complexes 1–3 under physiological conditions
have been studied. In vitro anticancer activities have been demonstrated by MTT
assay, acridine orange/ethidium bromide and diamidino‐2‐phenylindole staining
against various cancerous cell lines.
4 Department of Chemistry, Karunya University,
Karunya Nagar, Coimbatore 641 114, India
5 Centre for Advanced Studies in Botany, School of
Life Sciences, University of Madras, Guindy
Campus, Chennai 600 025, Tamil Nadu, India
Correspondence
Periasamy Viswanathamurthi, Department of
Chemistry, Periyar University, Salem – 636 011,
India.
Email: viswanathamurthi72@gmail.com
Appl Organometal Chem 2016; 1–23
wileyonlinelibrary.com/journal/aoc
Copyright © 2016 John Wiley & Sons, Ltd.
1

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VIJAYAN ET AL.
KEYWORDS
anticancer activities in vitro, DFT, DNA/BSA binding, enzyme catalysis, ruthenium
(II)/copper(I) complex
1
|
INTRODUCTION
their binding capability with DNA/nuclear proteins. Many
important applications of these complexes require that they
In recent years, there has been a surge in the development of
thiourea‐ based ligands that support highly reactive transition
metal complexes. N,N ‐[(dialkylamino) (thiocarbonyl)]
benzamidine has been observed as a diverseselection of
multifunctional thiourea scaffolds that can support metal ions
by providing a wide variety of different steric and electronic
environments.[1–3] The benzamidine ligands can readily be
prepared by the simple reaction between the corresponding
benzimidoyl chlorides and functionalized primary amines.[4]
Due to the simultaneous presence of O, N, S electron donors,
the substituted benzamidine ligands exhibit diverse coordina-
tion modes (Scheme 1) and form stable complexes with a
large number of transition metal ions.[5,6]
Among these, the dibasic tridentate (ONS, SSS, SOS,
SNS or SNN) mode of coordination is very common and
many examples have been reported previously.[2,7] But coor-
dination of benzamidine ligands to the metal ion in
monodentate fashion through the sulfur atom is rare. How-
ever, recently there have been a few reports of ruthenium(II)
arene complexes with thiourea ligands forming a six‐mem-
bered ring, in which ruthenium is coordinated by
monodentate sulfur atom without any internal cleavage in
ligands due to intramolecular hydrogen bonding between
the thiourea N─H and the amidic oxygen donor atom.[8,9]
In the present paper, hydrolytic cleavage via C═N bond in
the benzamidine ligands during the formation of new ruthe-
nium(II) and copper(I) complexes is reported.
can bind to DNA via an intercalative mode that could induce
cellular degradation. The planarity of the ligand and the pres-
ence of aromatic systems could favour the intercalation.
Moreover, the binding of drugs to serum albumin in vitro,
considered as a model in protein chemistry to study the bind-
ing behaviour of proteins, has been an interesting research
field in bioinorganic chemistry. The binding of protein folded
drugs (metal complexes) has made an important contribution
to the stabilization of extracellular fluid volume by contribut-
ing to oncotic pressure (known also as colloid osmotic pres-
sure), and assistance in the transport and distribution of
body fluids between blood vessels and body tissues.[11,12]
Therefore, the development of anticancer agents targeting
both DNA and proteins has been much in demand.
A number of catalysts having bio‐mimicking activity for
various enzymes have been designed by chemists. Such arti-
ficial enzymes have the same catalytic function but they are
more stable and structurally less complex than enzymes. In
particular, biochemically important enzymatic processes like
catalytic oxidation of 3,5‐di‐tert‐butylcatechol to quinone
(catecholase activity) and hydrolytic reactions, i.e. hydrolysis
of phosphodiester bond (phosphodiester cleavage), are of
considerable importance.[13] Synthetic enzyme models are
helpful in understanding the mechanistic aspects of enzyme
action. Studies of metal complexes which mimic the activities
of catacholase and phosphatase are very useful and promising
for the development of bioinspired environmentally friendly
catalysts.
Cisplatin, carboplatin and oxaliplatin are proven antican-
cer drugs used worldwide for the treatment of a range of can-
cers. Nevertheless, there is limited effectiveness due to severe
side effects and acquired resistance due to prolonged treat-
ment which have spurred investigators to find alternative
metal‐based drugs.[10] Among them, ruthenium(II)/copper(I)
incorporating benzamidine ligands have attracted much atten-
tion due to their interesting biological properties, particularly
Based on the above considerations, in the work reported
here,
the
reactions
between
N,N‐[(diethylamino)
(thiocarbonyl)]‐substituted benzamidine ligands with ruthe-
nium(II)/copper(I) precursors were carried out and the coordi-
nation flexibility of ligands was studied (Schemes 2 and 3).
Moreover, complexes 1–3 were also investigated using den-
sity functional theory (DFT) calculation, and their DNA/
bovine serum albumin (BSA) binding, enzyme kinetico‐cata-
lytic properties and in vitro anticancer activities studied.
2
| EXPERIMENTAL
2.1 | Materials
All chemicals used in this study were of reagent grade and used
withoutfurtherpurification.Syntheticmanipulationswererou-
tinely performed under oxygen atmosphere. Doubly distilled
water was used to prepare buffers. Calf thymus DNA (CT‐
SCHEME 1 Diverse coordination modes of benzamidine ligands

VIJAYAN ET AL.
3
spectra were obtained with a JASCO V‐570 spectrophotome-
ter. Geometry optimization with the DFT method was
performed using the B3LYP (LANL2DZ/6‐311G) package.
Fluorescence spectral data were obtained with a JASCO
FP‐8200 fluorescence spectrophotometer at room tempera-
ture. Single‐crystal X‐ray diffraction data collections were
carried out at 293(2)–296(2) K with a Bruker Apex‐II CCD
area detector (for H2L1, 1 and 2) and Gemini A Ultra (for
3). The melting points were checked on a Technico micro
heating apparatus and are uncorrected. Stock solutions of
compounds (1.0 × 10−3 M in DMSO) were stored at 4 °C
and required concentrations were prepared for all experi-
ments. All the stock solutions were used after no more than
four days. Solutions of compounds were freshly prepared
1 h prior to biochemical evaluation. Data were expressed as
the mean the standard deviation from three independent
experiments.
SCHEME 2 Synthetic route of complexes 1 and 2
2.3 | Synthesis of [Ru(L1)(CO)(PPh3)2] (1)
A solution of H2L1 (0.1 mmol) in 20 ml of MeOH–CHCl3
(1:1 v/v) was added dropwise to a boiling solution of
[RuHCl(CO)(PPh3)3] (0.100 g, 0.1 mmol) in a MeOH–
CHCl3 solvent mixture (1:1 v/v) (20 ml). The mixture was
gently heated under reflux for 12 h, which resulted in a rapid
change of colour from light yellow to maroon. The reaction
was monitored by TLC using a silica gel on aluminium sheets
with a 90/10 mixture of petroleum ether–ethyl acetate as the
mobile phase. After completion of the reaction, the resulting
solution was filtered and the filtrate was left standing for slow
evaporation of the solvent. After three days reddish‐brown
coloured small size crystals suitable for X‐ray diffraction
were obtained. Yield 85%; brown. Micro analytical data for
C55H49N3O2P2RuS required (%): C, 67.47; H, 5.04; N,
4.29; S, 3.28. Found (%): C, 67.32; H, 5.01; N, 4.11; S,
3.09. IR (KBr pellet, cm−1): 1527 (C═N), 820 (C═S),
1951 (C ≡ O). 1H NMR (DMSO‐d6, ppm): 1.15 (t,
J = 7.0 Hz, 3H, CH3), 1.19 (t, J = 7.0 Hz, 3H, CH3),
3.64 (q, J = 7.0 Hz, 2H, CH2), 3.81 (q, J = 7.0 Hz, 2H,
CH2), 6.65 (d, J = 7.2 Hz, 1H, H2), 6.67 (d, J = 7.2 Hz,
1H, H5), 6.72 (dd, J(4–3) = 7.2 Hz, J(4–5) = 7.2 Hz 1H,
H4), 6.81 (dd, J(3–2) = 7.2 Hz, J(3–4) = 7.2 Hz 1H, H3),
6.85 (dd, J(10–9) = 7.4 Hz, J(10–11) = 7.4 Hz, 1H, H10),
6.85 (dd, J(12–13) = 7.4 Hz, J(12–11) = 7.4 Hz, 1H, H12),
6.89 (s, 1H, H11), 7.25 (d, J = 7.7 Hz, 2H, H9,13), 7.41–
7.61 (m, PPh3). 13C NMR (CDCl3, ppm): 203.01 (C ≡ O),
169.19 (C═S), 165.85 (C─O, C1), 158.08 (C═N, C7),
141.99 (C─N, C7), 132.65 (Ar C6),132.30 (Ar C2), 132.15
(Ar C5), 132.10 (Ar C4), 132.04 (Ar C3), 131.89 (Ar C8),
131.75 (Ar C9), 131.60 (Ar C13), 131.44 (Ar C10), 126.42
(Ar C12), 126.35 (Ar C11), 126.10–117.62 (PPh3), 13.21,
13.31 (CH3, C), 47.61, 48.74 (CH2, C). 31P NMR
(DMSO‐d6, ppm): 36.98. UV–visible (CHCl3, λmax (nm)
(ε, dm3 mol−1 cm−1)): 238 (46 400), 268 (24 800), 307
(16 740), 474 (6580). ESI‐MS (calcd, found, m/z): 979.08,
SCHEME 3 Synthetic route of complex 3
DNA), BSA and 4‐nitrophenyl phosphate disodium salt hexa-
hydrate (4‐NPP) were obtained from Genei, Bangalore and
Himedia, India, respectively. Ethidium bromide (EB), 3,5‐di‐
tert‐butylcatechol (3,5‐DTBC) and tris(hydroxymethyl)
aminomethane were purchased from Sigma‐Aldrich and used
as received.
2.2 | General methods
Elemental analyses (C, H, N and S) were carried out with a
Vario EL III CHNS analyser. Infrared (IR) spectra were
recorded as KBr pellets using a PerkinElmer FT‐IR spectro-
photometer in the range 400–4000 cm−1. (1H, 13C) NMR
spectra were measured with a Bruker Ultra Shield at
300 MHz using DMSO‐d6 or CDCl3 as solvent and
tetramethylsilane as an internal reference. 31P NMR spectra
were measured with a Bruker Ultra Shield at 300 MHz using
DMSO‐d6 or CDCl3 as solvent and o‐phosphoric acid as an
external standard. Mass spectra for the complexes were
obtained with an advanced Q‐TOF micro™ mass spectrome-
ter using an electrospray ionization probe. All mass spectral
data are given in the form: m/z, assignment. Electronic

4
VIJAYAN ET AL.
980.4 (M)+. Single crystals suitable for X‐ray determination
were grown by slow evaporation of methanol–chloroform
(1:1 v/v) solution of 1 at room temperature.
performed with a Bruker SMART APEX 2 at 293(2)–296
(2) K and a Gemini Ultra Oxford Diffraction automatic dif-
fractometer equipped with a CCD detector. Suitable single
crystals of compounds were mounted on a glass fibre with
epoxy cement. The crystals were cut into a fitting size (less
than collimator cross‐sectional diameter). The X‐ray radia-
tion employed was generated from Mo sealed X‐ray tube
(Kα, λ = 0.71073 Å) fitted with a graphite monochromator
for all compounds. The data were corrected for Lorentz and
polarization effects with the SMART suite of programs and
for absorption effects with SADABS[14] and SCALE3
ABSPACK (CrysAlis RED, Oxford Diffraction Ltd, Version
1.171.29.2) scaling algorithm. A data collection strategy
using ω and ϕ scans at 0.5° scan technique yielded full hemi-
spherical data with excellent intensity statistics. Structure
solutions and refinements were done using the programs
SHELXS‐97/14. The structures were solved by direct
methods to locate the heavy atoms, followed by difference
maps for the light non‐hydrogen atoms. Anisotropic thermal
parameters were refined for the rest of the non‐hydrogen
atoms. Hydrogen atoms were placed geometrically and
refined isotropically. Details of the data collection and refine-
ment are gathered in Table 1 and important bond lengths and
angles for the compounds are summarized in Table 2.
2.4 | Synthesis of [Ru(1,1‐DT)(Cl2)(CO)(PPh3)2] (2)
Complex 2 was prepared using the above procedure by
reacting a suspension of [RuHCl(CO)(PPh3)3] with HL2.
Yield 76%; yellow. Micro analytical data for
C42H42Cl2N2O2P2RuS required (%): C, 58.88; H, 4.94; N,
3.27; S, 3.74. Found (%): C, 58.54; H, 4.79; N, 3.04; S,
3.65. IR (KBr pellet, cm−1): 3381 (NH2), 829 (C═S), 1952
1
(C ≡ O). H NMR (CDCl3, ppm): 1.22 (t, J = 7.2 Hz, 3H,
CH3), 1.26 (t, J = 7.2 Hz, 3H, CH3), 3.85 (q, J = 7.2 Hz,
2H, CH2), 3.94 (q, J = 7.2 Hz, 2H, CH2), 7.19–7.89 (m,
PPh3), 8.17 (s, NH2). 13C NMR (CDCl3, ppm): 204.12
(C ≡ O), 177.69 (C═S), 140.66 (C─N), 118.41–125.44 (m,
PPh3) 14.42, 14.65 (CH3), 46.50, 47.71 (CH2). 31P NMR
(DMSO‐d6, ppm): 34.83. UV–vis (CHCl3, λmax (nm) (ε,
dm3 mol−1 cm−1)): 239 (45 400), 273 (26 800), 330 (11
200), 420 (1800). ESI‐MS (calcd, found, m/z): 856.78,
822.3 (M − Cl)+. Single crystals suitable for X‐ray determi-
nation were grown by slow evaporation of methanol–chloro-
form (1:1 v/v) solution of 2 at room temperature.
2.5 | Synthesis of [Cu(1,1‐DT)(Cl)(PPh3)2] (3)
2.7 | Theoretical calculation (DFT)
Complex 3 was prepared by a procedure similar to that used
for complex 1 by reacting equimolar quantity of the ligands
(H2L1 or HL2) (0.1 mmol) in 10 ml of methanol and a sus-
pension of [Cu(CH3COO)(PPh3)2] (0.1 mmol) in 10 ml of
chloroform. After completion of the reaction, the resulting
solution was filtered and the filtrate was left standing for
the slow evaporation of the solvent. After four days orange
coloured crystals suitable for X‐ray diffraction were obtained.
Yield 60%; orange. Micro analytical data for
C41H42ClCuN2P2S required (%): C, 65.15; H, 5.60; N,
3.71; S, 4.24. Found (%): C, 65.01; H, 5.51; N, 3.61; S,
The calculations were carried out using the Gaussian 09
program.[15] The DFT/B3LYP method was used for the
geometry optimization and electronic structure determination.
The calculations were performed using the DZVP basis set on
ruthenium atom and 6‐311 g (d, p) basis set was used to
describe the copper atom. The PCM solvent model was used
to calculate the electronic structures with trichloromethane
as the solvent. GaussSum 3.0 was used to calculate group
(ligands and metal centre) contribution to the molecular
orbitals.[16] The molecular orbital plots were generated using
the Chemcraft program package (http://www.chemcraftprog.
com).
1
4.11. IR (KBr pellet, cm−1): 3379 (NH2), 814 (C═S). H
NMR (CDCl3, ppm): 1.37 (t, J = 7.2 Hz, 3H, CH3), 1.62 (t,
J = 7.2 Hz, 3H, CH3), 3.68 (q, J = 7.2 Hz, 2H, CH2), 3.52
(q, J = 7.2 Hz, 2H, CH2), 6.43–7.65 (m, PPh3), 8.21 (s,
NH2). 13C NMR (CDCl3, ppm): 171.44 (C═S), 140.66
(C─N), 121.41–131.53 (m, PPh3), 13.91, 14.09 (CH3),
48.17, 49.61 (CH2). 31P NMR (DMSO‐d6, ppm): −3.80.
UV–visible (CHCl3, λmax (nm) (ε, dm3 mol−1 cm−1)): 238
(45 800), 263 (35 200), 333 (19 400). ESI‐MS (calcd, found,
m/z): 755.8, 721.3 (M − Cl)+. Single crystals suitable for X‐
ray determination were grown by slow evaporation of metha-
nol–chloroform (1:1 v/v) solution of 3 at room temperature.
2.8 | DNA binding studies
2.8.1
| Emissive titration
Fluorescence spectra titrations were performed at room tem-
perature in 95% Tris–HCl/NaCl buffer (5 mM Tris–HCl/
50 mM NaCl buffer, pH = 7.2) to investigate the binding
affinity between CT‐DNA and complexes 1–3. Mixing of
such solutions with the aqueous buffer DNA solutions used
in the studies never exceeded 5% (v/v) DMSO in the final
solution, which was needed due to low aqueous solubility
of most compounds. The CT‐DNA concentration per nucleo-
tide was determined by fluorescence spectrometry using the
complex excitation wavelength. During titration, an equal
quantity of CT‐DNA was added to both the complex solution
and reference solution to eliminate the intensity of CT‐DNA
2.6 | X‐ray structure determination
Crystals of H2L1 and 1–3 suitable for single‐crystal X‐ray
analysis were obtained from slow evaporation of methanol–
chloroform solutions. The crystal data collections were

VIJAYAN ET AL.
5

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VIJAYAN ET AL.

VIJAYAN ET AL.
7
itself and Tris–HCl/NaCl buffer was subtracted through base-
line correction. Emissive titration experiments were
performed with a fixed concentration of complexes 1–3
(25 μM). While gradually increasing the concentration
(0–10 μM) of DNA, the emission intensities were recorded
for complexes in the range 350–650 nm. Titrations were man-
ually done by a micropipette for the addition of CT‐DNA.
2.10 | Enzyme kinetic studies
2.10.1
| Catechol oxidation
These kinetic experiments were carried out using fluores-
cence quenching spectra under pseudo‐first‐order condi-
tions.[17] Complexes 1–3 were preferred as representative
catalysts to study the catalytic oxidation of catechol. The
emission intensity of 3,5‐DTBC at λemis = 435 nm
(λex = 401 nm) was monitored using selected compounds.
Complexes 1–3 (1 × 10−4 M) in DMF were added to 100
equivalents of 1 × 10−3 M solutions of 3,5‐DTBC under
aerobic conditions. Emissive intensity of the resultant
reaction mixture was plotted with respect to wavelength at a
regular interval of 15 min using a fluorescence spectropho-
tometer in the range 410–700 nm. The dependence of the rate
on substrate concentration and different kinetic parameters
were obtained by treatment of complexes with 3,5‐DTBC
and monitoring the increase in emission intensity at 435 nm
(the peak corresponding to the quinone band maxima) as a
function of time.
2.8.2
| EB displacement assay
The relative binding propensity of complexes to CT‐DNA
was determined with an EB‐bound CT‐DNA solution in
Tris–HCl/NaCl buffer (5 mM Tris–HCl/50 mM NaCl buffer,
pH = 7.2). The fluorescence spectra were recorded at excita-
tion wavelength of 510 nm for an emission range of 604 nm.
In the fluorescence quenching spectra, the reduction in emis-
sion intensity measured the binding mode of complexes to
CT‐DNA. The addition of increasing amounts of the com-
plexes to the DNA‐EB adduct quenched the fluorescence.
Before the emission spectra were recorded, CT‐DNA was
pretreated with EB in the ratio [DNA]/[EB] = 1 for 30 min
at room temperature in order to fully react. Then the titration
compounds were added to this mixture of EB–DNA and the
changes in the fluorescence intensity were recorded.
2.10.2
| Phosphate ester hydrolysis
The hydrolysis of 4‐NPP was determined using fluorescence
quenching spectroscopy. Its hydrolytic tendency was measur-
able spectrophotometrically by monitoring the time evolution
of p‐nitrophenolate in DMF through a wavelength scan from
410 to 700 nm over 2 h. The emission intensity of 4‐NPP at
485 nm (λex = 401 nm) was monitored using complexes
1–3. The hydrolase activity involved the preparation of stock
solutions of complexes (0.05 × 10−3 M) and the substrate
4‐NPP (1 × 10−3 M), at higher concentrations in DMF. The
dependence of the rate on various concentrations and differ-
ent kinetic parameters were obtained by treatment of
0.05 × 10−3 M solution of complexes 1–3 with 40 equiva-
lents of 4‐NPP (the peak corresponding to the phenolate band
maxima) as a function of time.
2.8.3
| Viscosity experiment.
Viscosity measurements were carried out using an
Ubbelodhe viscometer maintained at a constant temperature
of 30.0 °C ( 0.1 °C) in a thermostatic bath. The flow time
was measured three times, after 5 min of incubation with each
addition of the complexes 1–3 and the average flow time was
taken for calculation of relative viscosity. Relative viscosities
for CT‐DNA in the presence and absence of the complexes
were calculated from the relation η = (t − to)/to, where t is
the observed flow time of DNA‐containing solution and to
is the flow time of Tris–HCl/NaCl buffer alone. Data were
presented as (η/ηη0)1/3 versus binding ratio (r = [com-
pounds]/[DNA] = 0.0–0.1), where η is the viscosity of
CT‐DNA in the presence of the compound and η0 is the
viscosity of CT‐DNA alone.
2.11 | In Vitro anticancer activities
2.11.1
| Cell culture
Human cervical cancer (HeLa), human hepatocellular carci-
noma (HepG2) and human breast cancer (MCF‐7) cell lines
were obtained from the National Centre for Cell Sciences
Repository, University of Pune, India. The normal Vero and
cancer cells were maintained in a humidified atmosphere
containing 5% CO2 at 37 °C in Dulbecco's modified Eagle's
medium supplemented with 100 units of penicillin,
100 μg ml−1 streptomycin and 10% foetal bovine serum.
Briefly, normal Vero and cancer cells, HeLa, HepG2,
MCF‐7, were precultured in 96‐well microtiter plates for
48 h under 5% CO2. Complexes 1–3 were dissolved in 0.1%
DMSO (the concentration of DMSO did not exceed 0.1% v/
v) to obtain a solution of 1 mM each. The samples were then
diluted to 100 μM in phosphate‐buffered saline (PBS) solu-
tion and filter‐sterilized using a 0.22 μm syringe filter. This
2.9 | Protein binding studies
The binding mode of complexes 1–3 with BSA was investi-
gated using fluorescence spectra at room temperature with
an excitation wavelength for BSA at 280 nm and monitoring
the emission at 344 nm by keeping the concentration of BSA
constant (1 μM) while increasing the complex concentration
(0–50 μM) regularly. The excitation and emission slit widths
(each 5 nm) remained constant for all the experiments. A scan
rate of 200 nm min−1 was used. In addition, absorption titra-
tion experiments were carried out by keeping the concentra-
tion of the complexes (20 μM) and BSA (1 μM) as
constant. Furthermore, the type of quenching mechanism of
complexes was determined from UV–visible absorption
spectra in the range 200–600 nm.

8
VIJAYAN ET AL.
100 μM solution in PBS was further used in cell cytotoxicity
studies. The cells (1 × 106 cells ml−1 per well) were seeded in
a 96‐well plate. One day after seeding, the cells were treated
with or without different concentrations of test complexes
and re‐incubated at 37 °C in a CO2 incubator for 24 h. After
the incubation, the cells were visualized using an inverted
Olympus microscope.
counted, and the percentage of apoptotic cells was calculated
on the basis of cellular morphological features. The results
were recorded as the mean of three independent experiments.
3
| RESULTS AND DISCUSSION
3.1 | Synthesis and characterization
2.11.2
| Protocol for MTT assay
Surprisingly little is known about the coordination behaviour
of benzamidine ligands H2L1 and HL2, which can be synthe-
sized from the reactions of N,N‐[(diethylamino)
(thiocarbonyl)]benzimidoyl chloride with functionalized
amines such as 2‐aminophenol and 2‐picolylamine in dry
acetone.[21] In the presence of supporting base NEt3, such
reactions proceed quickly under mild conditions. The prog-
ress of the reaction can readily be checked by TLC on silica
gel and is indicated by the formation of a colourless precipi-
tate of NEt3⋅HCl, which is almost insoluble in acetone. We
have relied on the use of the stable starting materials
[RuHCl(CO)(PPh3)3][22] and [Cu(CH3COO)(PPh3)2][23] for
the synthesis of ruthenium(II)/copper(I) complexes. The reac-
tion of H2L1 with ruthenium(II) precursor [RuHCl(CO)
(PPh3)3] affords complex 1 of the type [Ru(L1)(CO)(PPh3)2]
in which the ligand coordinates in tridentate ONS mode.
While the reaction of H2L1 with copper precursor [Cu
(CH3COO)(PPh3)2] induces C═N bond cleavage of the
ligand and affords complex 3 of the type [Cu(1,1‐DT)(Cl)
(PPh3)2] (1,1‐DT = 1,1‐diethylthiourea) in which the ligand
coordinates in monodentate fashion.
The MTT (3‐(4,5‐dimethylthiazol‐2‐yl)‐2,5‐diphenyltetrazolium
bromide) assay is a quantitative, sensitive and reliable colori-
metric technique generally used for assessing viability and pro-
liferation of cells. Cell viability with complexes 1–3 against
normal Vero and HeLa, HepG2 and MCF‐7 cancer cell lines
was determined by MTT assay.[18] Complexes 1–3 were added
to micro‐wells containing the cell culture at final concentrations
in the range 10–100 μg ml−1. Then each well was loaded with
10 μl of MTT solution (5 mg ml−1 in PBS, pH = 7.4) for 4 h
at 37 °C. The formed purple formazan crystals were dissolved
in 200 μl of DMSO and the cell viability was determined by cal-
culating the absorbance of each well at 540 nm using a Bio‐Rad
ELISA plate reader.[19] All data were representative of three
independent experiments, and the percentage of cell viability
was calculated according to the following equation:
Inhibition rate (IR %) = [OD (control) − OD (drug‐
treated cells)/OD (control)] × 100.
<NI > The corresponding IC50 (concentration of drug
that inhibits cell growth by 50%) value was determined by
nonlinear regression analysis using Origin 6.0 software.
The ligand HL2 reacts with ruthenium(II) and copper(I)
precursors to form complex 2 of the type [Ru(1,1‐DT)(Cl2)
(CO)(PPh3)2] and complex 3, respectively, in which the
ligand undergoes C═N cleavage and coordinates in
monodentate fashion via C═S group. The proposed mecha-
nism for hydrolytic cleavage and solvent‐assisted formation
of complex 2 is shown in Scheme 4. The formation complex
3 follows the same mechanistic pathway. To the best of our
2.11.3
| Nuclear staining assay
Nuclear staining using acridine orange (AO)/EB and
diamidino‐2‐phenylindole (DAPI) was measured by fluores-
cence microscopy according to the method previously
described.[20] HeLa and HepG2 cancer cells (1 × 106 in
number) were cultured in six‐well plates at 37 °C in an incu-
bator with fixed concentration of complexes 1–3 (50 μM) for
24 h. The cells were harvested and washed with ice‐cold PBS
and 40 μl of AO/EB solution (1 part of 100 μg ml−1 AO in
PBS; 1 part of 100 μg ml−1 EB in PBS) was added. After
staining, the cells were washed with PBS twice, suspended
in 200 μl of PBS, and the nuclear morphology was observed
under a fluorescence microscope in less than 20 min. For
DAPI staining, the treated cells were fixed with 80% ethanol
at room temperature for 30 min. The fixative was removed
and the cells were washed three times with PBS, and then
incubated with DAPI (1 μg ml−1) for 45 min at room temper-
ature in the dark. Both techniques were used to determine
viable cells, early apoptotic cells with blebbing, and necrotic
cells. The functions of dye have been reported beforehand.[31]
The HeLa and HepG2 cancer cells were mounted on slides,
and the images were observed under a fluorescent micro-
scope in green/blue filter with excitation at 350 nm and emis-
sion at 460 nm. At least 200 cells from each slide were
SCHEME 4 Proposed mechanism for the formation of complex 2

VIJAYAN ET AL.
9
knowledge, there are no other reports of hydrolytic cleavage
in benzamidine ligands (Scheme 2) with ruthenium(II) and
copper(I) complexes (1–3).
deprotonation of the ligand which is also confirmed by the
IR data. In the free ligand HL2, the signal due to Car─NH
group appears as a broad singlet at 6.93 ppm. But for 2 and
3, Car─NH group signal completely disappears and new
─NH2 peak appears in the range 8.17–8.21 ppm due to
hydrolytic cleavage via C═N bond. Moreover, the proton
signals of the two methylene groups for ligands and com-
plexes, which should consequently be two quartets, appear
The new compounds are crystalline, non‐hygroscopic
solids, air‐stable at room temperature, soluble in common
organic solvents such as dichloromethane, chloroform,
benzene, acetonitrile, ethanol, methanol, dimethylsulfoxide
and dimethylformamide and insoluble in hexane, petroleum
ether and diethyl ether. The analytical data for the compounds
agree well with the proposed molecular formulae. These
compounds have been systematically characterized using
UV–visible, IR, NMR (1H, 13C, 31P) and ESI‐MS spectral
studies. In addition, the molecular structures of ligand H2L1
and complexes 1–3 have been authenticated using single‐
crystal X‐ray crystallography.
as four
well‐separated
multiplet
resonances
at
3.52–3.94 ppm. This pattern of the methylene signals has
previously been rationalized by the rigid structure of the
tertiary amine group, which makes the methylene protons
magnetically non‐equivalent with respect to their axial and
equatorial positions.[9] Furthermore two triplet signals at
1.15–1.62 ppm for the methyl groups in N(CH2CH3)2 are
observed in the spectra of ligands and complexes. In addi-
tion, the spectra of ligands and complexes show a series of
overlapping multiplets for aromatic protons at
6.43–7.89 ppm.
3.2 | Spectroscopic studies
The characteristic IR bands of ligands H2L1 and HL2 and
their complexes 1–3 provide substantial information about
metal–ligand bonding and all the bands are listed in Section
2. No attempt has been made to assign each individual band
to a specific vibration. However, the IR spectra of ligands
exhibit νNH bands in the region 3217–3421 cm−1. Upon com-
plexation, νNH peaks disappear from the spectra of complexes
1–3 indicating the deprotonation of these groups in the
ligand.[24] However for complexes 2 and 3 a new broad band
appears in the region 3379–3381 cm−1 indicating the forma-
tion of new NH2 group due to hydrolytic cleavage via C═N
in the benzamidine ligands. In addition, the broad signal
appearing at 3060 cm−1 due to phenolic ─OH group in ligand
H2L1 completely disappears during formation of complex 1.
Furthermore, a strong vibration observed at 1608–1620 cm−1
for the ligands corresponding to νC═N group is shifted to
lower frequency in the spectrum of complex 1 and
completely disappears in those of complexes 2 and 3 due to
hydrolytic cleavage via C═N bond. A sharp band is observed
in the range 820–843 cm−1 ascribed to νC═S group in the
ligands, which is shifted to lower frequency upon complex
1 formation. However, the νC═S stretching frequency of com-
plexes 2 and 3 decreases by 20–30 cm−1, which suggests that
the ligands are bound to the ruthenium(II)/copper(I) centre
via the sulfur atom only. Parallel properties were witnessed
for previously reported thiourea complexes.[25] These results
clearly confirm that the coordination binding mode occurs
through ONS donor (for 1) and S donor (for 2 and 3) atoms.
Proof of chelation can also be obtained from the NMR
(1H, 13C, 31P) spectra of ligands and complexes. The data
are summarized in Section 2 and the spectra depicted in
The 13C NMR spectra show the expected signals in the
appropriate regions. For the uncoordinated ligands, the C═N
and C═S signals of H2L1 and HL2 appear in the regions
around 149.36–155.41 and 187.08–189.61 ppm, respec-
tively. Upon the coordination and formation of new ruthe-
nium(II)/copper(I) complexes,
a
downfield shift is
observed for the signals of C═N (around 6–10 ppm), while
C═S carbon signals appear in the upfield region between
169.19 and 177.69 ppm.[26,27] These are consistent with
ONS (for 1) or S (for 2 and 3) coordination and deproton-
ation as well as C═N cleavage of benzamidine ligands. The
C ≡ O carbon resonating region at 203.01–204.12 ppm is
comparable with that of reported ruthenium carbonyl com-
plexes.[28] In addition, a couple of signals appear at 13.21–
49.61 ppm for the CH2 and CH3 carbons. Furthermore, the
aromatic carbons appear in the region 117.62–132.83 ppm
for all the complexes.
The coordination of the triphenylphosphine as co‐ligands
and their configuration in the new complexes 1–3 are con-
firmed from analysis of 31P NMR spectra (Figures S5–S7).
The appearance of only one sharp singlet at 36.98 and
34.83 ppm for ruthenium(II) complexes 1 and 2 suggests
the
presence
of
two
magnetically
equivalent
triphenylphosphines trans to each other.[29] The appearance
of one sharp singlet observed at −3.80 ppm for complex 3
suggests the presence of two magnetically equivalent
triphenylphosphines cis to each other.[30]
The UV–visible spectra of the complexes (Figure S8)
were recorded in DMSO solution for different time intervals
(0–12 h). The obtained spectra are similar, suggesting that
complexes 1–3 retain their structure in solution within the
timeframe used for the biological experiments. Electronic
spectra of the complexes display intense absorptions in the
visible and UV regions. The absorptions in the UV region
are assignable to transitions within the ligand orbitals. The
bands around 238–273 nm are assigned to ligand‐centred
1
Figures S1–S4. In the H NMR spectra, the signal due to
the phenolic Car─OH and Car─NH groups of free ligand
H2L1 appear at 9.95 and 9.42 ppm, respectively. These
groups are absent in the spectrum of 1, supporting the depro-
tonation and coordination via oxygen and nitrogen atom to
the ruthenium(II) ion.[2] This fact confirms the double

10
VIJAYAN ET AL.
π → π* and n → π* transitions. The absorption maxima
located in the range 307–333 nm are assigned to S
(pπ) → M(dπ) (M = Ru2+) ligand‐to‐metal charge transfer
(LMCT) transition for 1 and 2 and M(dπ) → S(pπ)
(M = Cu+) metal‐to‐ligand charge transfer (MLCT) transi-
tion for 3.[31] Furthermore, the complexes show broad bands
at 420–474 nm, attributed to the d–d transition bands of
spin‐paired d[6] (complexes 1 and 2) species with a distorted
octahedral structure. The geometry of the complexes was
further confirmed using single‐crystal X‐ray diffraction
analyses.
Information about the composition and stability of the
complexes was also acquired from ESI‐MS (in positive‐
ion mode) spectral studies. The ESI‐MS data for com-
plexes 1–3 are listed in Section 2 and spectra depicted
in Figures S9–S11. Out of these, 1–3 display their
molecular‐ion peaks at m/z 980.4, 822.3 and 721.3,
respectively, assigned to [M]+ for 1 and [M − Cl]+ for
2 and 3 which reveal that the identity of the complexes
is retained in solution. The observed isotopic distributions
and their simulation patterns are in good agreement with
the assigned formulations.
FIGURE 2 Perspective view (30% probability ellipsoids) of complex 1 with
atom numbering scheme
3.3 | Single‐Crystal X‐ray diffraction studies
two independent molecules within the unit cell. The C─N
and C═N bond distances (Table 2) found in H2L agree well
with that reported for other similar ligands containing C─N/
C═N bonds. The thione form is confirmed by the bond
length of C(14)─S(1) (1.696 Å) and C(32)─S(2)
(1.693 Å) which are very close to a formal C═S bond
length (1.60 Å). While C(7)─N(1) bond (1.289 Å) has only
partial double bond character. This is found in all similar
ligands and suggests that the proton is mainly located at
the N1 atom. The configuration of H2L1 is described as
E, dZ configuration, which has also been found in some
related ligands.[2,32] The E, dZ configuration is obtained
due to intramolecular hydrogen bond between O1 and S1
atoms. In addition, H2L1 contains one intramolecular
hydrogen bond which creates a pseudo‐hydroxo bridged
binuclear structure (Figure S12).
Single‐crystal X‐ray diffraction studies of H2L1 and 1–3
confirm the conclusions drawn from the spectroscopic
studies. To gain insight into the coordination chemistry
and structural parameters of these compounds, good‐quality
single crystals were isolated by slow evaporation of a
concentrated methanol–chloroform solution and character-
ized by X‐ray diffraction. Details of the data collection,
solution and refinement are gathered in Section 2 and
Table 1. ORTEP views of H2L1 and the complexes (30%
probability ellipsoids) along with a partial atom numbering
scheme are shown in Figures 1–4 and important bond
lengths and angles for the complexes are summarized in
Table 2.
Figure 1 depicts the molecular structure of H2L1. It
crystallizes in the triclinic Pі space group. H2L1 shows
Perspective views of complexes 1–3 with atomic num-
bering scheme are depicted in Figures 2–4, while selected
bond lengths and angles are given in Table 2. Complexes
1–3 crystallize in the orthorhombic, monoclinic and triclinic
P212121, P21/c and P‐1 space groups, respectively. In 1, the
coordination geometry around the ruthenium ion is hexa‐
coordination with an octahedral geometry, where the basal
plane is constructed with nitrogen, phenolic oxygen and
thiolate sulfur atoms of the ligand in a binegative tridentate
fashion. The remaining apical coordination sites are filled
by a carbonyl group and two triphenylphosphines trans to
each other. The ONS chelated benzamidine ligand coordi-
nates in a dibasic tridentate manner to the ruthenium(II)
ion forming a stable six−/five‐membered chelate ring. The
ruthenium(II) ion is therefore sitting in a core RuCONSP2
FIGURE 1 Perspective view (30% probability ellipsoids) of H2L1 with
atom numbering scheme. Compound H2L1 showing two independent
molecules

VIJAYAN ET AL.
11
interaction, in this complex the presence of CO, a stronger
π‐acidic ligand might have forced the bulky PPh3 ligands
to take trans position for steric reasons. The trans angles
deviate from linearity and N(1), S(1) (six‐membered ring)
leads to small N(1)–Ru(1)–S(1) bite angle of 94.50(19)°.
The ruthenium–ligand distances, namely C(55)─Ru(1)
1.863(10) Å, O(2)─Ru(1) 2.161(5) Å, N(1)─Ru(1) 2.166
(7) Å, S(1)─Ru(1) 2.345(2) Å, P(2)─Ru(1) 2.402(19) Å, P
(4)─Ru(1) 2.403(2) Å, found in 1 agree well with those
reported for other similar ruthenium(II) complexes
containing triphenylphosphine in trans position and the car-
bonyl group (CO) is trans to the nitrogen atom. The same
phenomenon occurred with rhenium complexes in different
oxidation states containing the same type of
benzamidines.[33]
In complex 2, the facile cleavage of the C═N bond during
reaction of HL2 with ruthenium(II) precursor is unexpected
and seems to be metal‐induced hydrolytic cleavage as shown
in Scheme 2. The coordination geometry around ruthenium
(II) ion exhibits hexa‐coordination with an octahedral geom-
etry, where the basal plane is constructed with only thiolate
sulfur atom of the unexpected cleaved HL2 in a neutral
monodentate fashion. In complex 2, the remaining apical
coordination sites are filled by a carbonyl group, chlorine
atom and two triphenylphosphines trans to each other. The
ruthenium atom is in a distorted octahedral environment with
trans angles of [P(2)–Ru(1)–P(1)] 178.18(6)° and [Cl(1)–Ru
(1)–S(2)] 167.54(6)°. The carbonyl group is trans to the coor-
dinated Cl(2) atom [Cl(2)–Ru(1)–C(37)] with an angle of
173.9(2)°. The other two axial sites are occupied by a
carbonyl group and one chlorine atom with Ru(1)─(37) and
Ru(1)─Cl(2) distances of 1.827(7) and 2.4748(16) Å. The
CO group occupies the site trans to the Cl (C(37)–Ru(1)–Cl
(2), 173.8(2)°). This may be a consequence of strong Ru
(II) → CO back donation as indicated by the short Ru(1)─C
(37) (1.827(7) Å) bond and low CO stretching frequency
(1952 cm−1), which prefers σ or π weak donor groups occu-
pying the site opposite to CO to favour the d–π back dona-
tion. The ruthenium–ligand distances, namely Ru(1)─S(2)
2.4096(16) Å, Ru(1)─C(37) 1.826(7) Å, Ru(1)─P(1)
2.4025(16) Å, Ru(1)─Cl(1) 2.4432(16) Å, Ru(1)─Cl(2)
2.4750(16) Å, Ru(1)─P(2) 2.3964(17) Å, found in complex
2 agree well with those reported for other similar ruthenium
(II) complexes containing triphenylphosphine in trans posi-
tion and the carbonyl group (CO) is trans to the chlorine
atom.[34] In addition, complex 2 contains one intramolecular
hydrogen bond which creates a pseudo‐chloro bridged
binuclear structure (Figure S12). Therefore, based on the
crystal structures and spectroscopic properties, we propose
a conceivable mechanism for the formation of complex 2 as
shown in Scheme 3.
FIGURE 3 Perspective view (30% probability ellipsoids) of complex 2 with
atom numbering scheme
FIGURE 4 Perspective view (30% probability ellipsoids) of complex 3 with
atom numbering scheme
coordination environment, which is distorted octahedral in
nature as reflected in all the bond parameters around ruthe-
nium. In 1, the trans angles are [P(2)–Ru(1)–P(4)] 175.92
(8)° and [O(2)–Ru(1)–S(1)] 171.93(16)°. The carbonyl
group is trans to the coordinated N(1) atom [N(1––Ru(1)–
C(55)] with an angle of 171.7(3)°. Though the PPh3 ligands
usually prefer to occupy mutually cis position for better π‐
In complex 3, the coordination geometry around copper
(I) ion exhibits four‐coordination with tetrahedral geometry,
where the basal plane is constructed with only thiolate sulfur
atom of the unexpected cleaved H2L1 and HL2. In complex

12
VIJAYAN ET AL.
3, the remaining apical coordination sites are filled by chlo-
rine atom and two triphenylphosphine cis to each other. The
copper atom is in a tetrahedral environment with cis angles
of P(1)–Cu(1)–P(2) 121.64(2)°, P(2)–Cu(1)–S(1) 110.80(2)°
and P(2)–Cu(1)–Cl(1) 104.72(2)°. The copper–ligand dis-
tances, namely Cu(1)─Cl(1) 2.3911(6) Å, Cu(1)─S(1)
2.3684(6) Å, Cu(1)─P(1) 2.2986(5) Å, Cu(1)─P(2) 2.3094
(6) Å, found in complex 3 agree well with those reported
for other similar copper(I) complexes containing
triphenylphosphine in cis position.[30,35] In addition, complex
3 contains one intramolecular hydrogen bond which creates a
pseudo‐chloro bridged binuclear structure (Figure S12). The
unit cell packing diagram of the compounds is shown in
Figure S13.
the intensity of electronic transitions is much greater than in
the case of octahedral ones. Similarly, the high value of the
absorption coefficient of the lowest energy absorption band
in the case of 1 also confirms the ligand‐to‐ligand charge
transfer character of this absorption band. The low intensity
of the absorption band with maximum at 420 nm in the case
of 2 results from the considerable participation of ruthenium
d orbitals in HOMO and LUMO levels. So, the lowest
energy absorption band has MLCT character with admixture
of d → d (LF) transitions.
3.5 | DNA interaction sudies
3.5.1
| Stability of compounds
To carry out a biological evaluation of complexes, checking
their stability in solution is an important requirement.[36–38]
To check the stability of complexes 1–3, electronic spectra
have been recorded in DMSO at four different time points
(0–12 h) through the UV–visible absorption method. The
UV–visible spectral results (Figure S4) do not reveal any
appreciable changes in either the intensity or the position
of the absorption bands in DMSO solution. This demon-
strates that the complexes retain a stable structure under
physiological‐type conditions.
3.4 | Computational details for theoretical
calculations
To obtain further insight into the electronic structures of the
compounds, optimizations of H2L1 and 1–3 have been car-
ried out using the DFT/B3LYP methodology. The molecular
geometries of H2L1 and 1–3 in the gas phase singlet states
have been optimized and the selected bond parameters are
given in Table 2. Vibrational frequency calculations were
performed for each of these compounds, to ensure that the
optimized geometries represented the local minima of the
potential energy surface and that only positive eigenvalues
were found. In general, the optimized geometry for the
compounds reproduces the overall details of the X‐ray
structure with a reasonable agreement. Contour plots of
the HOMO and LUMO of the compounds and the energy
gap are shown in Figure 5. The lowering of the HOMO–
LUMO band gap is essentially a consequence of the large
stabilization of the LUMO due to the strong electron‐
accepting ability of the electron‐acceptor group. As one
can see from Figures 5 and S14, and the data collected in
Table S17, the electron density of HOMO in 1 is mainly
localized on the H2L1 ligand (67% H2L1 + 11% dRu).
While in the case of 2 and 3, the share of 1,1‐DT ligand
decreases with the increased involvement of the metal ion
orbitals with the significant participation of chloride ligand
in the case of 3. Similar situation occurs in the case of
the LUMO which in 1 is localized mainly on the H2L1
ligand whereas in 2 in phosphine (52%) and dRu (35%)
orbitals and in the case of 3 the LUMO comprises the
PPh3 ligands. Thus, taking into account the composition
of the HOMO and LUMO, it can be assumed that the low-
est energy electronic absorption bands, with maxima above
400 nm, have MLCT character. However, in the case of 3
the participation of the inter‐ligand charge transfer transi-
tions (πCl → π*PPh3) is significant, which confirms the high
value of the absorption coefficient determined for the band
with maximum at 474 nm (ε = 10400). On the other hand,
for tetrahedral complex the share of ligand p orbitals in
the HOMO is justified because of the symmetry, and hence
3.5.2
| Fluorescence emission titration
Fluorescence emission titration offers an effective method
for following binding of metal complexes with CT‐DNA.[39]
Intercalation of metal complexes with base pairs of DNA is
usually shadowed by a hypochromic shift with a small red/
blue shift. On the other hand, non‐intercalative, electrostatic
interactions or damage of the DNA double helix result in a
hyperchromic shift. From the fluorescence emission titration
curves, the binding mode and propensity of 1–3 were
studied in the absence and presence of CT‐DNA at different
increasing concentrations.
From the emissive titration spectra (Figure 6), it is
apparent that upon the addition of increasing concentrations
of CT‐DNA (0–50 μM) to a solution of 1–3 there is
hypochromism of about 13.24% (358 nm), 35.12%
(355 nm) and 25.04% (288 nm), respectively, along with a
small red shift (ca 2–3 nm). The observed hypochromism
is due to an intercalative mode of binding involving a strong
stacking interaction between extended aromaticity of the
ligands and the base pairs of DNA. Based on the emission
enhancement, the intrinsic binding constant was obtained
according to the Scatchard equation:
CtðF−F0Þ
Cb ¼
Fmax−F0
where Ct is the total compound concentration, F the observed
fluorescence emission intensity at given DNA concentration,
F0 the intensity in the absence of DNA and Fmax the fluores-
cence of the totally bound compound. The fluorescence

VIJAYAN ET AL.
13
FIGURE 5 Frontier molecular orbitals of the compounds and their HOMO–LUMO energy gap and ground state optimized geometry (top). For the HOMOs
and LUMOs hydrogen atoms are omitted for clarity. Positive and negative regions are shown in grey and green, respectively
spectra of the tested compounds with increasing concentration
of CT‐DNA are depicted in Figure 6. Intrinsic binding con-
stant (Kb) was cast in the form of a Scatchard plot of r/Cf ver-
sus r, where r is the binding ratio of Cb/[DNA] and Cf is the
free ligand concentration. Kb and binding site size (s) values
for 1–3 with CT‐DNA are listed in Table 3. From available
data it is concluded that the complexes bind to DNA via
intercalative mode similar to earlier reports,[40] where the
order of DNA binding affinity is 3 > 1 > 2. This may be
ascribed to the presence of different functional groups in the
complexes and the lower value of s indicates intercalative
binding mode. Nevertheless, the binding mode needs to be
confirmed through further experiments.
3.5.2.1
|
EB displacement study
The fluorescence spectral technique is an effective method
for studying the interaction of metal complexes with DNA.

14
VIJAYAN ET AL.
FIGURE 6 Fluorescence titrations of complexes 1–3 (25 μM) with CT‐DNA (0–50 μM) (left). Scatchard plots of r/CF versus r for complexes 1–3
(right)
Hence fluorescence quenching experiments with EB‐bound
DNA were undertaken to understand the mode of DNA
interaction with complexes 1–3. EB, a cationic conjugated
planar molecule, is a well‐known DNA intercalator. It is
known that EB emits intense fluorescence in the presence
of DNA owing to its strong intercalation between adjacent
base pairs. The extent of fluorescence quenching of EB by
competitive displacement from DNA is a measure of the
strength of the interaction between the second molecule
and DNA.[41] The emission spectra of EB bound to CT‐
DNA in the absence and presence of complexes 1–3 are
shown in Figure 7.
TABLE 3 Fluorescence spectral parameters for complexes 1–3 bound to
CT‐DNA
Complex
Kb
s
Ksv
Kapp
1
2
3
5.26 × 104
3.98 × 104
7.26 × 104
0.02
0.10
0.08
2.36 × 104
1.19 × 104
1.03 × 105
3.17 × 105
6.30 × 105
7.25 × 106
As the concentration of complexes increases, the emission
band at 606 nm (Figure 7) for EB exhibits hypochromism of

VIJAYAN ET AL.
15
FIGURE 7 Fluorescence titrations of 1–3 (0–50 μM) with EB‐bound CT‐DNA (7.5 μM) (left). Scatchard plots of I0/I versus [Q] for complexes 1–3 (right)
up to 53.39, 36.87 and 82.34% with a slight red/blue shift from
theinitialfluorescenceintensity.Further,thequenchingparam-
eters for the magnitude of interactionwereascertained from the
classical Stern–Volmer equation:
a plot of I0/I versus [Q] as shown in Figure 7. From these
results it is evident that EB is displaced quite efficiently
from the binding sites of CT‐DNA by the complexes that
also strongly bind with DNA.
Further, apparent DNA binding constants (Kapp) have
been calculated using the following equation:
I0
¼ 1 þ Ksv½Qꢀ
I
KEB½EBꢀ ¼ Kapp½M50%ꢀ
where I0 and I are the emission intensities of EB‐bound
CT‐DNA in the absence and presence of the quencher
(complex) concentration [Q], respectively, which gives the
Stern–Volmer quenching constant (Ksv). The quenching
constant values (Table 3) were obtained from the slope of
where KEB = 1.0 × 10−7 M−1 is the DNA binding con-
stant of EB, [EB] is the concentration of EB (7.5 μM)
and [M50%] is the concentration of the compound used

16
VIJAYAN ET AL.
to obtain a 50% reduction in fluorescence intensity of
DNA pretreated with EB. The Kapp values for 1–3 are
given in Table 3. The DNA binding ability of the com-
pounds follows the order 3 > 1 > 2, which is in accord
with the results obtained from the fluorescence spectral
studies discussed above, and we conclude that all the
complexes may bind with DNA via intercalation.
of the binding of a drug with a protein will facilitate interpre-
tation of the metabolism and transporting processes of the
drug and explain the affiliation between structures and func-
tions of a protein.[43] In this regard, BSA is utilized exten-
sively because of its structural homology with human serum
albumin. The type of interaction between complexes 1–3
and BSA, whether static or dynamic quenching, was deter-
mined from UV–visible absorption studies. The absorption
spectra of BSA solutions (1 μM) in the absence and presence
of 1–3 (25 μM) are shown in Figure 9. The addition of the
complexes causes an increase in the intensity of BSA absorp-
tion at ca 280 nm with a small blue shift of about 2 nm,
suggesting a static interaction between complexes and BSA.
These UV–visible results clearly indicate a static interaction
from the formation of a complex between the quencher
(1–3) and the fluorophore (BSA) in the ground state.
3.5.3
| Viscometric study
These measurements were implemented to authenticate and
further clarify and investigate the conformational changes of
DNA upon interaction with the metal complexes. These mea-
surements can lend strong support in favour of intercalative
binding. In general, this method popularly accepts that a par-
tial and/or non‐classical intercalation of a ligand could bend
(or kink) the DNA helix, reducing its effective length con-
comitantly.[42] The viscosity of CT‐DNA increases when a
compound binds with DNA in intercalating mode but
remains unchanged when a compound binds with DNA in
electrostatic mode. From these considerations, this measure-
ment could lend strong support in favour of intercalative
binding. The changes in the relative specific viscosity of
CT‐DNA (200 μM) in the presence and absence of com-
plexes 1–3 (0–120 μM) were plotted as (η/η0)1/3 versus [com-
pound]/[DNA]. The relative viscosity of DNA solution
increases upon addition of all complexes in the order
3 > 1 > 2 (Figure 8). The viscosity changes might be consid-
ered evidence of all three complexes intercalate between the
base pairs of CT‐DNA. This result is consistent with our fore-
going fluorescence spectral studies.
3.6.2
| Fluorescence spectral study
In the fluorescence study, a solution of BSA (1 μM) was
titrated with increasing concentrations of 1–3 (0–50 μM)
in the range 290–440 nm (λex = 280 nm) with negligible
redshift. The observed red shift is mainly due to the active
site in protein hydrophobic environment. Figure 10 demon-
strates the effect of increasing concentration of complexes
on the fluorescence emission of BSA. The fluorescence
quenching constant (Kq) is defined by the following Stern–
Volmer equation:
I0
I
¼ 1 þ Ksv½Qꢀ ¼ 1 þ Kqτ0½Qꢀ
where I0 and I are the fluorescence intensity in the absence
and presence of the quencher, respectively. Kq is the
quenching rate constant, τ0 is the fluorescence lifetime of
biopolymer BSA (τ0 = 10−8 s)[43] and Ksv and [Q] are the
Stern–Volmer quenching constant and the concentration of
quencher, respectively. Kq values were calculated from plots
of I0/I versus [Q] (Figure 10). It can be seen that the Kq value
of BSA is higher than 1.0 × 1012, which is much higher than
3.6 | BSA interaction studies
3.6.1
| UV–visible spectral study
Serum albumins are the most extensively studied and applied
proteins due to their ability to transport amino acids and drug
molecules. Interaction of metallodrugs with proteins is signif-
icant for obtaining information about their properties such as
uptake, bio‐distribution, overall toxicity, mechanism of action
and shuttling of complexes to cancer cells. Therefore, studies
FIGURE 8 Relative viscosity (η/η0)1/3 of CT‐DNA in Tris–HCl buffer solu-
tion in the presence of complexes 1–3 in increasing amounts (r = 0–1)
FIGURE 9 Absorption spectra of complexes 1–3 with BSA

VIJAYAN ET AL.
17
FIGURE 10 Fluorescence titrations of complexes 1–3 (0–50 μM) with BSA (1 μM) (left). Stern–Volmer plots of I0/I versus [Q] and Scatchard plots of log[(I0 –
I)/I] versus log[Q] for complexes 1–3 (right)
TABLE 4 Quenching parameters of BSA for complexes 1–3
Complex
Ksv
Kq
Kbin
n
1
2
3
1.002 × 105
1.187 × 105
1.335 × 105
1.002 × 1013
1.187 × 1013
1.335 × 1013
7.50 × 106
1.03 × 1010
2.77 × 1010
1.45
2.01
2.10
SCHEME 5 Catalytic oxidation of 3,5‐DTBC to 3,5‐DTBQ in DMF in oxy-
gen atmosphere

18
VIJAYAN ET AL.
FIGURE 11 Oxidation of 3,5‐DTBC by complexes 1–3 monitored by fluorescence spectroscopy (left). Lineweaver–Burk plots for tested compounds (right)
the molecular fluorescence diffusing constant as evident from
Table 4. From the values of Kq, it is concluded that complex 3
interacts with BSA more strongly than the other complexes.
where Kbin is the binding constant of the compound with
BSA and n is the number of binding sites. Kbin and n have
been found from plots of log[(I0 − I)/I] versus log[Q]
(Figure 10). The calculated Kbin and n are gathered in
Table 4. The calculated value of n is around 2 indicating
the existence of two binding sites available in BSA for all
the complexes. The higher values of Kq and Kbin indicate
strong interaction between the BSA protein and the com-
plexes. It is noticed that the values of binding constant of
the complexes follows the order 3 > 2 > 1, suggesting that
the complexes effectively bind with BSA and hence can have
potential anticancer properties.
3.6.3
| Binding constant and binding site number.
The binding constant Kbin was calculated using the Scatchard
equation:
ꢀ
ꢁ
I0−I
I
log
¼ logKbin þ n log½Qꢀ

VIJAYAN ET AL.
19
3.7 | Catechol oxidase study
In order to check the ability of complexes 1–3 to show
catacholase mimic properties, the catalytic oxidation of 3,5‐
DTBC, which is taken as an exemplary substrate, showing a
low quinone–catechol reduction potential, was carried out.
In general, it is easy to oxidize and the bulky tert‐butyl
groups prevent further over‐oxidation such as ring opening
(Scheme 5). So far, only a limited number of ruthenium(II)/
copper(I) complexes have been reported to mediate catalytic
oxidation of 3,5‐DTBC.[44]
The reactions were carried out at 30 °C in presence of air
and monitoring was done by fluorescence spectroscopy.
Complexes 1–3 (1 × 10−4 M) were added to 100 equivalents
of a DMF solution of 3,5‐DTBC (1 × 10−2 M) under aerobic
conditions and the progress of the reaction was studied every
15 min up to 2 h. It is seen that on addition of 3,5‐DTBC, a
new band (Figure 11) appears and its intensity gradually
increasing at 434 nm may be attributed to LMCT band due
to the formation of the oxidized product 3,5‐di‐tert‐
butylbenzoquinone (3,5‐DTBQ). Thus, the experiment
clearly proves that the oxidation of 3,5‐DTBC to 3,5‐DTBQ
is catalysed by the new complexes, as it is well established
that 3,5‐DTBQ shows a maximum at λemis = 434 nm in pure
DMF.
SCHEME 6 Conceivable mechanism for the catalytic cycle of the oxidation
of 3,5‐DTBC
interesting to note that no blue coloration was observed in
the absence of 3,5‐DTBC.
According to the catecholase reaction mechanism
(Scheme 6), an electron transfer occurs in the rate‐deter-
mining step resulting in a substrate–catalyst 1:1 intermedi-
ate indicating that in the catalytic cycle there is only one
ruthenium/copper ion accepting one electron from the
substrate.[45] The Michaelis–Menten constant (Km) and
maximum initial rate (Vmax) were determined by lineariza-
tion using Lineweaver–Burk plots (Figure 11). The rate
constants for dissociation of substrates S (i.e. turnover
number, kcat) were calculated from graphs of 1/V versus
1/[S] (Figure 11), known as Lineweaver–Burk plots, using
the above equation and all these parameters are listed in
Table 5. Upon comparison of the data in Table 5, it
might be stated that complex 2 is a highly efficient cata-
lyst and has an appreciable turnover rate, and the order of
their activity is 2 > 1 > 3.
3.8 | Kinetic study of catecholase activity
The rate constant of the catalyst was determined by the initial
rate method. Furthermore, the catalytic behaviour showed
saturation kinetics, and a treatment based on the Michaelis–
Menten model seemed to be appropriate under excess sub-
strate conditions. Plots of ki versus [3,5‐DTBC] give
nonlinear curves of decreasing slope (Figure S15) which are
best designated by the following equations:
k2
k1
C þ S ⇌ CS ⇌ C þ P
k−1
k−2
Vmax½Sꢀ
V ¼
TABLE 5 Kinetic parameters for the catecholase activity of complexes 1–3
KM þ ½Sꢀ
Catalyst
Km (M)
Vmax (M m−1
)
Kcat (h−1
)
1
2
3
3.08 × 10−2
9.88 × 10−1
9.31 × 10−3
5.05 × 10−3
2.85 × 10−2
6.02 × 10−4
308
988
93
1
V
KM
1
1
¼
þ
Vmax ½Sꢀ Vmax
A feasible mechanism of 3,5‐DTBC oxidation promoted
by catalysts 1–3 is shown in Scheme 6 based on the report
of Chyn and Urbach.[45] Unfortunately we were unable to
characterize the intermediates. However, when a mixture of
starch–potassium iodide solution was added to a mixture of
complexes and 3,5‐DTBC, blue coloration formed, indicating
that H2O2 was produced during the course of reaction. It is
SCHEME
7
Hydrolysis of 4‐NPP to 4‐NP ion in DMF in oxygen
atmosphere

20
VIJAYAN ET AL.
3.9 | Phosphate hydrolysis activity
evolution of p‐nitrophenolate (4‐NP) ion in DMF at
λemis = 486 nm (λex = 435 nm) through a wavelength scan
from 400 to 800 nm over 15 min intervals up to 2 h with
40 equivalents of substrate being used relative to the catalyst
(Scheme 7).
Phosphates are widely found in biological organisms. Phos-
phate diesters such as DNA and RNA have metabolic inter-
mediates that exist as phosphomonoesters.[46] Their kinetic
stability makes monoesters particularly suitable for their bio-
logical roles. 4‐NPP ester is negatively charged and under
neutral conditions is very resistant towards hydrolysis. There-
fore the development of artificial catalysts capable of cleav-
ing phosphate ester bonds is of fundamental importance as
they may find applications in gene technology or as therapeu-
tic antisense oligonucleotides. Its hydrolytic tendency was
investigated using emissive intensities by monitoring the time
3.10 | Kinetic study of phosphatase activity
In order to obtain the rate constant of the catalyst the ini-
tial rate method was used. Moreover, the catalytic behav-
iour showed saturation kinetics, and a treatment based on
the Michaelis–Menten model seemed to be appropriate
FIGURE 12 Hydrolysis of 4‐NPP by complexes 1–3 monitored by fluorescence spectroscopy (left). Lineweaver–Burk plots for complexes 1–3 (right)

VIJAYAN ET AL.
21
under excess substrate conditions. The dependence of the
initial rate on the concentration of the substrate was mon-
itored at the respective wavelength by fluorescence spec-
troscopy, which corresponds to the increase in 4‐NP
concentration (Figure 12). Plots of ki versus [4‐NPP] give
nonlinear curves of decreasing slope (Figure S16) which
are best designated by first‐order kinetic equations. The
rate versus concentration of substrate data were scrutinized
on the basis of the Michaelis–Menten approach of enzy-
matic kinetics to get Lineweaver–Burk (double reciprocal)
plots as well as the values of the various kinetic parame-
ters Vmax, Km and Kcat. The rate constant for dissociation
of substrates S (i.e. turnover number, kcat) were calculated
from the graphs of 1/V versus 1/[S] (Figure 12) using the
kinetics equation and all these parameters are listed in
Table 6. While comparing all these data it can be con-
cluded that complex 3 is quite an efficient catalyst and
has an appreciable turnover rate and the order of the
activity is 3 > 2 > 1. Therefore, based on the crystal
structures, and physicochemical properties, it is concluded
that the hydrolysis of 4‐NPP follows the mechanism of
catacholase activity.
TABLE 7 In vitro cytotoxicity of compounds against normal and cancer
cell lines
IC50 (μg ml−1 a
)
Compound
Vero
HeLa
HepG2
MCF‐7
1
54.31 1.06
50.60 0.85
52.49 1.65
15.18 1.08
33.56 0.69
28.69 1.18
26.70 0.78
12.29 1.27
34.14 1.10
33.65 1.75
27.03 1.10
11.78 1.52
34.13 1.45
38.54 0.96
32.11 1.29
10.29 1.17
2
3
Cisplatin
a50%percent inhibitory concentration after exposure for 24 h in MTT assay.
behaviour of 3 is very similar to that of the DNA/BSA binding
affinities as discussed previously. Upon ratification of the
excellent cytotoxicity properties of complex 3, further
staining assay was conducted with HeLa and HepG2 cancer
cell lines to understand the mechanism of action.
3.11.2
| Apoptotic event: AO/EB and DAPI staining
Staining property was investigated to elucidate the basis for
the observed induction of cellular death, as it plays a central
role in the execution of the apoptotic process. Apoptosis is
characterized via DNA fragmentation, chromatin condensa-
tion and marginalization, membrane blebbing, cell shrinkage
and chromosomal DNA fragmentation of the cells into mem-
brane‐enclosed vesicles or apoptotic bodies and global
mRNA decay.[49] Activity is significantly enhanced in the
particular cell lines tested, being clear at 24 h for HeLa and
HepG2 cells. To detect cell death at a basic level, microscopic
evidence for apoptosis was obtained using AO, EB and DAPI
staining, the fluorescence patterns for which depend upon
viability and membrane integrity of cells. During morpholog-
ical analysis using AO/EB and DAPI staining method it can
observed that cell death increases due to cell membrane dis-
ruption with an increase in the concentration of each drug,
as nuclei appear light orange with a greenish background.
The morphological changes of images of the control cells
and treated HeLa and HepG2 cancer cells are depicted in
3.11 | Anticancer activity In Vitro
3.11.1
| Cancer cell growth inhibition (MTT assay)
The cytotoxicity of complexes 1–3 was determined using the
MTT assay against normal Vero and cancer cell lines, namely
human cervical cancer (HeLa), human hepatocellular carci-
noma (HepG2) and human breast cancer (MCF‐7) cell lines.
Cells were exposed to a broad range of drug concentrations
(10–100 μg ml−1) for 24 h, and the cell viability was quanti-
fied using the colorimetric method. The amount of cell prolif-
eration significantly decreases in a dose‐dependent manner on
supplementation with complexes 1–3, as observed within 24 h
of incubation with respective cell lines (Figure 13). For com-
parison purposes, the cytotoxicity of cisplatin was evaluated
under the same experimental conditions.[47,48] The results
were analysed by means of cell inhibition expressed as IC50
values and are summarized in Table 7. As evident from
Table 7, the IC50 values show that complex 3 exhibits slightly
higher inhibitory effect than the other complexes towards the
tested cancer cell lines. Moreover, the complexes display
slightly less cytotoxic activity than cisplatin. Importantly,
complexes 1–3 are less toxic towards the normal Vero cell line
as is evident from the higher IC50 values (Table 7). On the
basis of the IC50 values, the cytotoxicity of these complexes
can be arranged in the order 3 > 2 > 1. The cytotoxic
TABLE 6 Kinetic parameters for the phosphatase activity of complexes 1–3
Catalyst
Km (M)
Vmax (M m−1
)
Kcat (h−1
)
1
2
3
1.74 × 10−2
4.26 × 10−3
9.18 × 10−3
1.70 × 10−2
1.01 × 10−3
0.88 × 10−3
34.92
85.30
FIGURE 13 Cytotoxicity of complexes 1–3 after 24 h incubation with nor-
mal Vero and cancer cell lines MCF‐7, HeLa and HepG2
183.70

22
VIJAYAN ET AL.
FIGURE 14 Images of HeLa and HepG2 cancer cells after AO/EB, DAPI and EB staining, incubated for 24 h with treatment by complex 3 with fixed con-
centration. The yellow arrows show early apoptotic cells with blebbing, blue arrows show late apoptosis and red arrows show necrotic cells
Figure 14. In this figure, yellow arrow(s) show early apopto-
tic cells with membrane blebbing which is seen at fixed doses
(10 μg ml−1) of complex 3 and blue arrow(s) indicate late
apoptotic cells with chromatin aggregation that is highly con-
densed chromatin. The red arrow(s) show necrotic cells.
These results clearly suggest a mechanism of action of com-
plex 3 which induces necrosis in HeLa and HepG2 liver can-
cer cells favouring an apoptotic pathway.
undergo intercalative binding with CT‐DNA. Furthermore,
the protein binding results revealed that the complexes
showed high binding affinity with BSA and quenched the
fluorescence of BSA through a static quenching mechanism.
For all the complexes, from a strong piece of experimental
evidence, the difference of the driving force for the reduction
of the metal centres plays an important role in the oxidation
of the model substrate 3,5‐DTBC and the hydrolysis of
4‐NPP in DMF in oxygen atmosphere. Almost all the com-
plexes showed significant cytotoxicity, as evidenced by the
submicromolar IC50 values achieved towards various cancer
cell lines. The morphological changes of the apoptotic event
indicated that complex 3 induces necrosis in HeLa and
HepG2 cancer cells favouring an apoptotic pathway.
4
| CONCLUSIONS
In this work, ligands H2L1 and HL2 having different struc-
tural features as regards ruthenium(II) and copper(I) precur-
sors and new complexes 1–3 have been synthesized and
were characterized using various spectral, analytical and X‐
ray crystallographic techniques. The new ligands exhibited
ONS‐tridentate coordination mode for 1 (in a dianionic form)
and S‐monodentate coordination mode for 2 and 3 (in a
neutral form). The coordination mode is governed by hydro-
lytic cleavage of C═N bond in the benzamidine ligands by
metal precursors. The coordination geometry of complexes
1 and 2 was designated as distorted octahedral around ruthe-
nium ion and tetrahedral geometry was assigned for complex
3. According to the DFT studies, a HOMO–LUMO energy
gap evident for the charge transfer interactions occurred
within the molecules, as well as a lowering of the band gap.
The structural parameters of the compounds that are in good
agreement with results of X‐ray analysis were obtained by
theoretical calculations. The new complexes 1–3 have been
used to study their binding properties with CT‐DNA/BSA.
From the results obtained using fluorescence titrations and
EB competitive assay, it was concluded that all complexes
ACKNOWLEDGMENTS
Authors P.V. and P.V. gratefully acknowledge UGC (F. no.
40‐66/2011 (SR)) for financial support. R.N. is grateful to
DST for financial assistance (project no. SR/FT/CS‐95/
2010). The Gaussian 09 calculations were carried out in the
Wroclaw Centre for Networking and Supercomputing,
WCSS, Wroclaw, Poland (http://www.wcss.wroc.pl). The
authors thank The Director, CAS in Botany, School of Life
Sciences, University of Madras, Chennai for providing labo-
ratory facilities to perform the cell line studies.
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