DNA/protein interaction and cytotoxicity of palladium(II) complexes of thiocarboxamide ligands

Article abstract of DOI:10.1016/j.ica.2014.03.002

Four planar palladium(II) complexes with general formula [Pd(Cl)(L)(PPh₃)] (HL = N-substituted pyridine-2-thiocarboxamide) have been synthesized and characterized by analytical, spectral (IR, UV-Vis and ¹H, ¹³C and ³¹

Full text of DOI:10.1016/j.ica.2014.03.002

Inorganica Chimica Acta 416 (2014) 1–12
Contents lists available at ScienceDirect
Inorganica Chimica Acta
journal homepage: www.elsevier.com/locate/ica
DNA/protein interaction and cytotoxicity of palladium(II) complexes
of thiocarboxamide ligands
b
c
Elangovan Sindhuja a, Rengan Ramesh a, , Nallasamy Dharmaraj , Yu Liu
⇑
a School of Chemistry, Bharathidasan University, Tiruchirappalli 620024, India
b Department of Chemistry, Bharathiar University, Coimbatore 641046, India
c Institute of High Energy Physics, Chinese Academy of Sciences, Beijing 100049, China
a r t i c l e i n f o
a b s t r a c t
Article history:
Four planar palladium(II) complexes with general formula [Pd(Cl)(L)(PPh3)] (HL = N-substituted pyridine-
2-thiocarboxamide) have been synthesized and characterized by analytical, spectral (IR, UV–Vis and 1H,
13C and 31P NMR) and single crystal X-ray methods. Crystal structure of all the complexes indicated a
mono negative bidentate coordination of thiocarboxamide ligands to the palladium center via pyridine
nitrogen and thiol sulfur and reveals a square planar geometry. The interaction of palladium(II) thiocarb-
oxamide complexes with calf thymus DNA (CT-DNA) studied by absorption and emission spectroscopic
methods revealed that complexes 5–8 could interact with CT-DNA through intercalation. Further, the
interaction of the palladium complexes with bovine serum albumin (BSA) was investigated using UV–Vis,
fluorescence and synchronous fluorescence methods. The tryptophan and tyrosine residues in BSA as
model protein was quenched by the complexes in a static quenching process. The radical scavenging
ability of all the complexes was accessed by in vitro antioxidant assays involving DPPH radical, hydroxyl
radical, nitric oxide radical and ABTS radicals and was found to be excellent. Further, the anti-cancer
activity of complexes 5–8 against HeLa, MCF-7 and NIH-3T3 cell line has been studied. Though the
complexes 5, 6 and 8 showed more potent anticancer activity than cisplatin, complex 5 was found to
be superior.
Received 16 November 2013
Received in revised form 13 February 2014
Accepted 10 March 2014
Available online 22 March 2014
Keywords:
Palladium(II) complex
DNA/protein interaction
Antioxidant
Anticancer activity
Ó 2014 Elsevier B.V. All rights reserved.
1. Introduction
agents, sequence specific binding, structural probes, and therapeu-
tic agents [5]. The study on the interaction of transition metal
Over the decades, there has been an active research in pharma-
ceutical chemistry to design effective anticancer drugs, potentially
useful in the treatment of various cancers [1–4]. Cisplatin is one of
the leading chemotherapeutic drug among actinomycin, anthracy-
cline antibiotics and widely used metal-based anticancer drugs for
cancer therapy, but it possesses inherent limitations as serious side
effects such as neurotoxicity, tissue toxicity, nausea, nephrotoxi-
city, emetogenesis, gastrointestinal, bone marrow toxicity and
acquired drug resistance. Therefore, considerable attempts are
being made to replace this drug with suitable alternatives and
diverse transition metal complexes have been synthesized and
tested for their anticancer activities.
complexes with DNA is a vibrant area of research [6]. Bovine serum
albumin (BSA) is soluble protein that has the ability to transport a
multitude of endogenous and exogenous ligands such as fatty
acids, amino acids, steroids, metal ions and drugs in blood stream.
It is used to study the interaction of protein–drug complex in the
circulatory system because it has structural resemblance with
human serum albumin (HSA) [7].
Free radicals including reactive oxygen species (ROS) which is
generated from metabolism and or by the environmental factors
interact directly to the biological systems and the unbelievable
generation of free radicals have been implicated with variety of
chronic diseases including cancer, diabetes, atherosclerosis,
neurodegenerative disorders and arthritis [8,9]. Antioxidants are
compounds those can slow down or totally inhibit the oxidation
of lipid or other molecules by inhibiting the propagation of oxidiz-
ing chain reactions interms of prevention, interception and damage
repair [10,11].
Transition metal complexes play an important role in nucleic
acids chemistry are well-known to produce pronounced changes
in the various physicochemical properties of both DNA and inter-
acting agent for their diverse applications such as foot printing
Thiocarboxamide derivatives have raised considerable interest
due to their well known chelating ability and structural flexibility
⇑
Corresponding author. Tel.: +91 431 2407053; fax: +91 431 2407045/2407020.
E-mail address: ramesh_bdu@yahoo.com (R. Ramesh).
http://dx.doi.org/10.1016/j.ica.2014.03.002
0020-1693/Ó 2014 Elsevier B.V. All rights reserved.

2
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
which leads to high pharmacological properties. In some cases the
highest biological activities is associated with a metal complexing
agent [12]. Though several palladium(II) and platinum(II) thiosem-
icarbazone complexes having potential antitumor activity [13,14]
have been reported, the biological application of palladium com-
plexes with thiocarboxamide is not well explored [15]. Subscribing
to a similar point of view, we decided to inspect the potential of
palladium complexes as anticancer agents. Recently, we have re-
ported the catalytic activity of palladium(II) complexes containing
pyridine-2-thiocarboxamide ligands [16]. Herein, we wish to
investigate the biological properties of a different series of palla-
dium(II) thiocarboxamide complexes.
128.05, 126.17, 126.17, 126.02, 123.23, 121.42, 20.75, 17.80. dP
(160 MHz, ppm) 31.18.
2.2.3. [Pd(Cl)(j
2-S,N-C6H4CS = N-(4-BrPh)(PPh3)] (7)
Yield: 89%. M.p. 204 °C (with decomposition). Anal. Calc. for
C
30H23BrClN2PPdS: C, 51.75; H, 3.33; N, 4.02; S, 4.60. Found: C,
51.70; H, 3.36; N, 4.01; S, 4.63%. IR (KBr, cmꢀ1
)
m
= 1548(m),
1276(s). UV–Vis: kmax/nm (
e
max/dm3 molꢀ1 cmꢀ1): 374 (14465).
NMR (CDCl3): dH (400 MHz, ppm) 9.65 (t, 1H, C-1), 8.19–6.88 (m,
23H, ArH, PPh3). dC (100 MHz, ppm) 160.38, 149.67, 148.51,
138.73, 134.76, 134.65, 132.16, 132.06, 131.97, 131.28, 131.24,
129.22, 128.66, 128.60, 128.48, 128.31, 128.20, 126.34, 123.49,
123.40, 117.24. dP (160 MHz, ppm) 30.91.
2. Experimental
2.2.4. [Pd(Cl)(j
2-S,N-C6H4CS = N-(4-Bz)(PPh3)] (8)
Yield: 92%. M.p. 198 °C (with decomposition). Anal. Calc. for
2.1. Materials and instrumentation
C31H26ClN2PPdS: C, 58.96; H, 4.15; N, 4.44; S, 5.08. Found: C,
59.11; H, 4.17; N, 4.45; S, 5.06%. IR (KBr, cmꢀ1
)
m
= 1587(m),
Chemical reactions were carried out under atmosphere of air.
Solvents were dried and freshly distilled prior to use. Calf-thymus
DNA (CT-DNA), ethidium bromide (EtBr) and PdCl2(PPh3)2 pur-
chased from Sigma-Aldrich Chemie were used as received. Bovine
serum albumin (BSA) was purchased from Himedia Company.
The human breast cancer cell line (MCF-7), human cervical cancer
cell line (HeLa) and non-cancerous NIH-3T3 mouse embryonic
fibroblasts cell line were obtained from the National Centre for Cell
Science (NCCS), Pune, India. All other chemicals and reagents used
for the biological studies were of high quality in biological grade.
Elemental analyses were performed on a Vario EL III CHNS ele-
mental analyser. IR spectra were recorded on a Perkin–Elmer 597
spectrophotometer, using KBr pellets. 1H, 13C and 31P NMR were re-
corded on a high resolution Bruker Avance 400 spectrometer. Melt-
ing points were performed with an electrical instrument and are
uncorrected. UV–Vis spectroscopy was recorded on a Varian Cary
300 Bio UV–Vis spectrophotometer using cuvettes of 1 cm path
length. Emission intensity measurements were carried out using
a Jasco FP-6200 spectrofluorometer.
1267(s). UV–Vis: kmax/nm (
e
max/dm3 molꢀ1 cmꢀ1): 381 (14838).
NMR (CDCl3): dH (400 MHz, ppm) 9.53 (t, 1H, C-1), 7.35–7.14 (m,
23H, ArH, PPh3), 4.59 (s, 2H, CH2). dC (100 MHz, ppm) 160.35,
149.57, 148.34, 138.63, 134.75, 134.64, 131.14, 130.10, 129.40,
129.34, 128.84, 128.13, 126.08, 125.52, 123.98, 123.39, 119.3,
45.81. dP (160 MHz, ppm) 31.00.
2.3. X-Ray structure determinations
Single crystals of all the complexes suitable for X-ray diffraction
were grown by slow evaporation of dichloromethane-ethanol mix-
ture at room temperature. A single crystal of suitable size was cov-
ered with Paratone oil, mounted on the top of a glass fibre, and
transferred to a Bruker SMART APEX II single crystal X-ray diffrac-
tometer using monochromated Mo-K
a radiation (kI = 0.71073 Å).
Data were collected at 293 K. Structures were solved with direct
method using SHELXS-97 or SIR-97 and were refined by full matrix
least-squares method on F2 with SHELXL-97. Non-hydrogen atoms
were refined with anisotropy thermal parameters. All hydrogen
atoms were geometrically fixed and allowed to refine using a rid-
ing model. Frame integration and data reduction were performed
using the Bruker SAINT-Plus (Version 7.06a) software. The multi-
scan absorption corrections were applied to the data using SADABS
software. The crystal data and structure refinement parameters
for the structures 5–8 are given in Table 1.
2.2. Synthesis
The N-substituted pyridine-2-thiocarboxamide ligands (1–4)
and palladium(II) thiocarboxamide complexes (5–8) were
prepared according to literature methods [17,16] respectively.
Further, their identity was confirmed using elemental analysis,
X-ray crystallography and IR, NMR, UV–Vis spectroscopic methods.
2.4. DNA binding studies
2.2.1. [Pd(Cl)(j
2-S,N-C6H4CS = N-(3-MePh)(PPh3)] (5)
Electronic absorption spectral measurements were carried out
in twice distilled buffer containing tris(hydroxymethyl)-amino-
methane (Tris, 5 mM) and sodium chloride (50 mM) and adjusted
to pH 7.2 with hydrochloric acid. Solutions of calf thymus DNA in
TrisꢀHCl buffer gave a ratio of UV absorbance at 260 and 280 nm
is of about 1.89–2.01 which indicates the free of protein in DNA.
Concentrated stock solutions of metal complexes were prepared
by dissolving them in a 5% dimethylsulfoxide (DMSO) and 95%
TrisꢀHCl buffer and diluting suitably with the corresponding buf-
fer to required concentrations for all the experiments. Absorption
spectral titration experiments were performed by maintaining a
Yield: 75%. M.p. 1890 °C (with decomposition). Anal. Calc. for
C
30H23Cl2N2PPdS: C, 58.96; H, 4.15; N, 4.44; S, 5.08. Found: C,
58.99; H, 4.13; N, 4.46; S, 5.07%. IR (KBr, cmꢀ1
)
m
= 1545(m),
1269(s). UV–Vis: kmax/nm (
e
max/dm3 molꢀ1 cmꢀ1): 388 (15501).
NMR (CDCl3): dH (400 MH, ppm) 9.59 (d, 1H, C-1), 7.89–6.83 (m,
23H, ArH, PPh3), 2.21 (s, 3H, CH3). dC (100 MHz, ppm) 159.84,
148.48, 148.18, 137.50, 136.78, 133.75, 133.65, 130.09, 130.07,
128.38, 127.09, 124.07, 122.37, 121.08, 118.02, 20.49. dP
(160 MHz, ppm) 31.01.
2.2.2. [Pd(Cl)(
j
2-S,N-C6H4CS = N-(2,4,6-TriMePh)(PPh3)] (6)
fixed concentration of the complex (25
otide concentration (0ꢀ25 M).
lM) and varying the nucle-
Yield: 69%. M.p. 193 °C (with decomposition). Anal. Calc.for
l
C33H30ClN2PPdS: C, 60.10; H, 4.58; N, 4.25; S, 4.86. Found: C,
59.96; H, 4.55; N, 4.26; S, 4.87%. IR (KBr, cmꢀ1
)
m
= 1561(m),
2.4.1. Competitive binding measurements
1267(s). UV–Vis: kmax/nm (
e
max/dm3 molꢀ1 cmꢀ1): 369 (12953).
DNA was pretreated with ethidium bromide in the ratio [DNA]/
[EtBr] = 10 for 30 min at 27 °C. EtBr was non-emissive in Tris–HCl
buffer solution (pH 7.2) due to fluorescence quenching of the free
EtBr by the solvent molecules. In the presence of DNA, EtBr showed
enhanced emission intensity due to its intercalative binding to
NMR (CDCl3): dH (400 MHz, ppm) 9.58 (t, 1H, C-1), 7.66–6.66 (m,
21H, ArH, PPh3), 2.16, 1.91 (s, 9H, CH3). dC (100 MHz, ppm)
160.78, 149.64, 138.61, 134.74, 134.69, 134.63, 134.58, 132.09,
131.13, 131.01, 130.98, 129.46, 128.91, 128.23, 128.17, 128.11,

E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
3
Table 1
Crystallographic data for complexes 5–8.
Complex
5
6
7
8
Empirical formula
Formula weight
Wavelength (Å)
Crystal system
Space group
a (Å)
C
31H26ClN2PPdS
C33H30ClN2PPdS
659.47
0.71073
orthorhombic
Pbca
C30H23BrClN2PPdS
696.29
0.71073
monoclinic
C2/c
25.0113(4)
10.6446(2)
22.9703(5)
90
110.792(2)
90
5717.23(2)
8
1.618
2.292
2768
C31H26ClN2PPdS
631.42
0.71073
631.42
0.71073
monoclinic
P2(1)/n
11.5287(1)
11.2841(1)
21.8774(2)
90
95.636(1)
90
2832.29(4)
4
1.481
0.903
1280
0.06 ꢁ 0.05 ꢁ 0.05
1.87–25.00
44913
triclinic
ꢀ
P1
9.1309(2)
14.858(3)
42.253(9)
90
90
90
5732(2)
8
1.528
0.896
2688
9.2061(3)
12.2173(5)
13.0731(5)
92.483(2)
99.141(2)
102.537(2)
1412.47(9)
2
1.485
0.905
640
0.07 ꢁ 0.06 ꢁ 0.05
1.58–26.00
25565
b (Å)
c (Å)
a
(°)
b (°)
c
(°)
V (Å3)
Z
Dcalc (Mg/m3)
Absorption Coefficient (mmꢀ1
F(000)
)
Crystal size (mm3)
h Range for data collect (°)
Reflections collected
Independent reflection
0.07 ꢁ 0.06 ꢁ 0.05
1.93–28.32
35258
0.06 ꢁ 0.06 ꢁ 0.05
1.74–24.92
46688
4999
7024
4974
5558
Maximum and minimum transmission
Refinement method
0.9562 and 0.9469
full-matrix least squares on F2
4999/0/335
1.033
0.9559 and 0.9392
0.8940 and 0.8747
0.9558 and 0.9387
Data/restraints/parameters
7024/0/335
1.047
4974/0/334
1.039
5558/0/334
1.038
Goodness-of-fit (GOF) on F2
Final R indices [I > 2
R indices (all data) R1, wR2
Rint
r
(I)] R1, wR2
0.0213, 0.0516
0.0251, 0.0543
0.0281
0.0613, 0.1434
0.1157, 0.1839
0.1201
0.0403, 0.1051
0.0603, 0.1170
0.0443
0.0249, 0.0600
0.0308, 0.0638
0.0267
DNA. The excitation wavelength 545 nm was fixed for EtBr bound
to DNA were recorded with an increasing amount of the palla-
dium(II) complex concentration and the emission range was ad-
justed before measurements. The metal complexes (0–50 lM)
were then added to this mixture and their effect on the emission
solution, and 1.0 mL of DMSO. The reaction was started by adding
0.5 mL of ascorbic acid and incubated at 80–90 °C for 15 min in a
water bath. The reaction was then terminated by the addition of
1.0 mL of ice-cold trichloroacetic acid. Three milliliters of Nash
reagent was added and left at room temperature for 15 min. The
intensity of the color formed was measured at 412 nm against re-
agent blank.
intensity was measured.
Nitric oxide (NO) radical scavenging activity was determined
based on the procedure by Griess Illosvoy reaction [20]. The reac-
tion mixture (3 mL) containing sodium nitroprusside (10 mM,
2 mL), phosphate buffer saline (0.5 mL) at different concentration
2.5. Protein binding studies
The excitation wavelength of BSA at 280 nm and the emission at
335 nm were monitored for the protein binding studies. The exci-
tation and emission slit widths and scan rates were maintained
constant for all of the experiments. Concentrated stock solutions
of complexes were prepared by dissolving the complex in DMSO:
phosphate buffer (1:100) and diluted suitably with phosphate buf-
fer to required concentrations for all the experiments. A 2.5 mL
and standards (50–250 l
g mLꢀ1) were incubated at 25 °C for
150 min. After incubation, 0.5 mL of the incubated solution
containing nitrite was pipetted and mixed with 1 mL of sulfanilic
acid reagent and allowed to stand for 5 min for completing diazo-
tization. Then, 1 mL of N-1-naphthyl ethylene diamine dihydro-
chloride was added, mixed and allowed to stand for 30 min at
25 °C. The absorbance of pink colored chromophore formed during
diazotization was measured at 540 nm.
solution containing appropriate concentration of BSA (1
titrated by successive additions of a 10 mL stock solution of
complexes (0–50 M). For synchronous fluorescence spectra the
similar concentrations of BSA and the complexes were used. The
lM) was
l
The ABTS (2,20-Azino-3-ethylbenzthiazoline-6-sulfonic acid
diammonium salt) cationic radical was studied according to the re-
ported procedure [21]. The ABTS was dissolved in water to a 5 mM
concentration and its radical was produced by reacting with potas-
sium persulfate (2.4 mM) for 12 h at room temperature in the dark.
The solution was then diluted in 60 mL methanol to obtain an
absorbance of 0.706 0.001 at 734 nm. BHT of various concentra-
spectra were measured at two different
and 60 nm.
Dk values, such as 15
2.6. Antioxidant activity determination
The DPPH (2-20-diphenyl-1-picrylhydrazyl) radical scavenging
activity were performed based on the procedure described by Blios
tions (100–500 l
g mLꢀ1) were allowed to react with 1.0 mL of
the ABTS cationic radical solution and the absorbance was taken
at 30 °C exactly after 30 min, the initial mixing and the reaction
mixture without test sample was used as control.
[18]. Various concentrations (10–30 lg/mL) of sample were mixed
with 1 mL of 0.1 mM methanol solution of DPPH and final volume
was made upto 4 mL with double distilled water. The tubes were
allowed to stand at room temperature for 20 min. The control
was prepared as above without any extract, and methanol was
used for the baseline correction. Changes in the absorbance of
the samples were measured at 517 nm.
For the above four assays, all the tests were run in triplicate, and
various concentrations of the complexes were used to fix a concen-
tration at which the complexes showed around 50% of activity. In
addition, the percentage of activity was calculated using the
formula, % of suppression ratio = [(A0 ꢀ AC)/A0] ꢁ 100. A0 is the
absorbance of the control and AC is the absorbance of the complex.
The 50% activity (IC50) can be calculated using the percentage of
activity.
The hydroxyl (OH) radical scavenging activity of the BHT (Butyl-
ated hydroxy toluene) was determined according to the Nash
method [19]. Different concentrations (50–250
lg/mL) of extract
were added with 1.0 mL of iron-EDTA solution, 0.5 mL of EDTA

4
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
2.7. In vitro anticancer activity. maintenance of cell lines
formation of complexes (Scheme 1). The analytical and spectral data
(IR, UV–Vis and NMR) confirmed the stoichiometry of the complexes.
The IR spectra of the free ligands show a medium to strong band
in the region 3310–3214 cmꢀ1 which is characteristic of the N–H
functional group. The free ligands also display mC=S absorptions in
the region 824–776 cmꢀ1. The bands due to mN–H and mC=S stretch-
ing vibrations are not observed in the complexes indicating that
the ligands undergo tautomerization and subsequent coordination
of the thiolate form during complexation. The IR spectra of the
complexes did not display mS–H at 2589–2572 cmꢀ1 suggesting
the deprotonation of the thiol proton prior to coordination.
Moreover, a new band is appeared around 1278–1269 and 1587–
1545 cmꢀ1 which corresponds to mC-S and mC=N for the thiocarbox-
amide complexes. This bidentate coordination is further confirmed
by the presence of band at 495–450 cmꢀ1, which is assigned to mPd-
The assays were carried out in monolayer cells were detached
with trypsin-ethylenediaminetetraacetic acid (EDTA) to make sin-
gle cell suspensions and viable cells were counted using a hemocy-
tometer and diluted with medium containing 5% FBS to give final
density of 1 ꢁ 105 cells/mL. One hundred microlitres per well of
cell suspension were seeded into 96-well plates at plating density
of 10000 cells/well and incubated to allow for cell attachment at
37 °C, 5% CO2, 95% air and 100% relative humidity. After 24 h the
cells were treated with serial concentrations of the test samples.
They were initially dissolved in neat DMSO to prepare the stock
(200 mM) and stored frozen prior to use. At the time of drug addi-
tion, the frozen concentrate was thawed and an aliquot was diluted
to twice the desired final maximum test concentration with serum
free medium. Additional three, 10 fold serial dilutions were made
Coordination of the thiolate sulfur is indicated by a decrease in
S.
to provide a total of four drug concentrations. Aliquots of 100
of these different drug dilutions were added to the appropriate
wells already containing 100 L of medium, resulted the required
final drug concentrations. Following drug addition the plates were
incubated for an additional 48 h. The medium containing no sam-
ples served as a control and triplicate was maintained for all
concentrations.
l
L
energy of thioamide band as well as by the presence of a band at
392–370 cmꢀ1 that is assignable to
mPd–N. In addition, other
l
characteristic bands due to triphenylphosphine are also present
in the region 1467–1424 cmꢀ1 in the spectra of all the complexes
[16]. A week intensity band is observed at 1098–1087 cmꢀ1 char-
acteristic of the coordinated pyridine.
The 1H, 13C and 31P NMR spectra of complexes 5-8 were
recorded in CDCl3 and are shown in Figs. S1-12 (Supporting infor-
mation). The 1H NMR spectrum of the free ligands show the signal
at d 2.37 ppm for ligand 1 and d 2.35 and 2.24 ppm for ligand 2 are
due to the methyl protons, and the signal at d 4.67 ppm is corre-
sponding to methylene protons of ligand 4. The 1H NMR spectra
of the complexes causes a significant downfield shift for the proton
adjacent to the nitrogen (H-1) and appeared in the range d 9.65–
2.7.1. Evaluation by MTT assays
MTT is a yellow water soluble tetrazolium salt. A mitochondrial
enzyme in living cells, succinate-dehydrogenase, cleaves the tetra-
zolium ring, converting the MTT to an insoluble purple formazan.
Therefore, the amount of formazan produced is directly propor-
tional to the number of viable cells. After 48 h of incubation,
9.59 ppm, this arise from a
r-effect based on electron donation
15 lL of MTT (5 mg/mL) in phosphate buffered saline was added
to Pd(II) via the nitrogen lone pairs. The thiocarboxamide proton
is lost upon coordination and is not observed in the spectra of
the palladium(II) complexes, suggesting coordination of this ligand
to Pd(II) through pyridine nitrogen and thiol sulfur. All the
complexes show a multiplet in the range d 8.34–6.66 ppm for the
aromatic protons including triphenylphosphine. A sharp singlet
that appeared for the –NH protons of the free ligand in the region
d 11.92–11.34 ppm are absent in all the complexes, further
supporting thione-thiol tautomerization and coordination of the
thiolate sulfur to the palladium(II) ion. In the case of methyl
protons in the phenyl ring, a singlet is appeared at d 2.21 ppm
for complex 5 and two singlets at d 2.16 and 1.91 ppm for complex
6 whereas the methylene signal of the benzyl unit is observed as a
singlet at d 4.59 ppm in the complex 8. The I3C NMR spectra
showed one signal at d 20.49 for complex 5 and two signals at d
20.75 and 17.80 ppm for complex 6, which are assigned to methyl
carbons. Similarly, the resonance at d 45.81 ppm attributable to the
to each well and incubated at 37 °C for 4 h. The medium with
MTT was then flicked off and the formed formazan crystals were
solubilized in 100 lL of DMSO and then the absorbance at
570 nm was measured using micro plate reader. Nonlinear regres-
sion graph was plotted with the percentage of cell inhibition versus
log10 concentration. From this, the IC50 value was calculated by
using the following formula.
%cell inhibition ¼ 1 ꢀ AbsðsampleÞ=AbsðcontrolÞ ꢁ 100:
3. Results and discussions
3.1. Synthesis and spectroscopic study
The reaction between PdCl2(PPh3)2 and pyridine-2-thiocarboxa-
mide ligands at reflux for 3 h in ethanol medium resulted in the
Na2S.9H2O
H
N
S8
R-NH2
N
CH3
N
R
S
Ethanol, 80oC
[PdCl2(PPh3)2]
1
R = 4-BrPh
R = 3-MePh
3
R = 2,4,6-TriMePh
2
4
R = Bz
CH3
N
N
N
N
CH3
N
N
N
N
Cl Pd
S
Cl Pd
S
Cl Pd
PPh3
S
Cl Pd
S
CH3
Br
H3C
PPh3
PPh3
PPh3
5
6
7
8
Scheme 1.

E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
5
methylene carbon for complex 8. In 13C NMR spectra the signal as-
signed to the thioketone carbon, which moves upfield from d
197.32–184.06 ppm in the ligand to d 160.38–149.64 ppm in the
palladium(II) complexes, resulting from the reduced C–S bond
order on coordination. The 31P NMR spectra for all the complexes
5–8, show singlet in the range d 31.18–30.91 ppm indicates the
presence of one PPh3 group in the complexes.
The formation of the reaction product can be monitored by con-
ventional spectrophotometer in the near-UV–Vis region have been
recorded in dichloromethane solution. The d–d spin allowed sin-
glet–singlet and three spin forbidden singlet–triplet transitions
are predicted for square-planar complexes of palladium(II) but
strong charge-transfer transitions may interfere and prevent the
observation of some of the expected bands, which is actually
observed in the electronic absorption spectra of the complexes.
The absorption spectra of all the palladium(II) complexes were
recorded in chloroform in the range 800–200 nm at room temper-
ature and all the complexes showed three bands with absorption
maxima in the region 401–241 nm. The high intensity bands in
the 277–241 nm region were assigned to ligand-centered (LC)
Fig. 2. The molecular structure of [Pd(Cl)(PPh3)(L2)], 6. (hydrogen atoms are
omitted for clarity; displacement parameters are drawn at 50% probability level).
transitions and have been designated as intraligand
p–
p⁄ and
n–p⁄ transitions involving energy levels higher in energy than
the ligand LUMO levels. In the spectra of all the complexes, the
lowest energy bands observed in the region 401–369 nm were
attributed to the charge transfer due to metal and ligand transi-
tions. These prominent strong bands of both intraligand and charge
transfer absorptions support the idea of a square planar environ-
ment for the Pd metal ions.
3.2. Structure of the complexes
The molecular structures of the complexes 5–8 has been deter-
mined by single crystal X-ray diffraction to find out the coordina-
tion mode of the thiocarboxamide in the complexes and are shown
in Figs. 1–4. The summary of the data collection and refinement
parameters are shown in Table 1 and selected bond angles and
bond distances are gathered in Table 2. The thiocarboxamide
ligand coordinates in a bidentate fashion to palladium ion via pyr-
idine nitrogen and thiolate sulfur in addition one PPh3 and one
chlorine groups. The complexes 5 and 7 crystallized in the mono-
clinic space group ‘P21/n’ and ‘C2/c’ where the complex 6 crystal-
lized in the orthorhombic space group ‘Pbca’ whereas the
ꢀ
complex 8 crystallized in the triclinic space group ‘P1’. Palladium
Fig. 3. The molecular structure of [Pd(Cl)(PPh3)(L3)], 7. (hydrogen atoms are
is therefore sitting in a NSPCl coordination environment, which is
square planar in nature as reflected in all the bond parameters
around palladium. In the complex 5, the thiocarboxamide ligand
omitted for clarity; displacement parameters are drawn at 50% probability level).
Fig. 1. The molecular structure of [Pd(Cl)(PPh3)(L1)], 5. (hydrogen atoms are
Fig. 4. The molecular structure of [Pd(Cl)(PPh3)(L4)], 8. (hydrogen atoms are
omitted for clarity; displacement parameters are drawn at 50% probability level).
omitted for clarity; displacement parameters are drawn at 50% probability level).

6
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
Table 2
Selected bond lengths (Å) and angles (°) for the complexes 5–8.
Complex
5
6
7
8
Pd(1)–N(1)
Pd(1)–S(1)
Pd(1)–P(1)
Pd(1)–Cl(1)
2.1141(15)
2.2400(6)
2.2624(5)
2.3449(6)
1.754(2)
2.109(4)
2.108(3)
2.1088(17)
2.2567(6)
2.2495(6)
2.3608(6)
1.767(2)
2.2641(14)
2.2418(14)
2.3376(14)
1.758(Å)
2.2587(13)
2.2630(12)
2.3682(11)
1.751(5)
S(1)–C(6)
N(2)–C(6)
1.275(3)
1.289(7)
1.288(6)
1.273(3)
N(1)–Pd(1)–S(1)
N(1)–Pd(1)–P(1)
S(1)–Pd(1)–P(1)
N(1)–Pd(1)–Cl(1)
S(1)–Pd(1)–Cl(1)
P(1)–Pd(1)–Cl(1)
C(5)–N(1)–Pd(1)
C(6)–S(1)–Pd(1)
84.44(5)
176.11(5)
91.93(2)
94.17(5)
177.58(2)
89.51(2)
85.11(13)
177.13(13)
94.74(5)
94.67(13)
177.19(5)
85.62(5)
84.57(11)
176.56(11)
92.83(4)
93.48(11)
175.54(5)
89.27(4)
84.87(5)
175.77(5)
90.91(2)
94.44(5)
172.49(2)
89.75(2)
118.97(13)
101.91(8)
118.8(4)
99.37(19)
118.5(3)
100.65(17)
118.48(14)
100.00(8)
binds the metal center at N and S forming one five membered che-
late ring with bite angles of 84.44(5)° N(1)–Pd(1)–S(1), 91.93(2)°
S(1)–Pd(1)–P(1), 94.17(5)° N(1)–Pd(1)–Cl(1) and 89.51(2)° P(1)–
Pd(1)–Cl(1). The bond lengths of Pd(1)–N(1) and Pd(1)–S(1) are
2.1141(15) and 2.2400(6) Å respectively. This shows high planarity
of the PPh3, Cl and N, S donor atoms of pyridine thiocarboxamide in
which PPh3 are trans to pyridine nitrogen and Cl are trans to thio-
late suphur. These bond distances are very similar to those ob-
served in other palladium(II) complexes. The Pd(1)–P(1) bond
length 2.2624(5) Å is in agreement with other structurally charac-
terized palladium phosphine complexes. Further, it was observed
that the complexes 6-8, adopt similar geometry as in the complex
5, with slight changes in bond angles and bond lengths.
respectively. The binding constants (Kb) is the ratio of the slope
to the intercept which has been estimated to be
6.3123 ꢁ 105 Mꢀ1
,
2.1724 ꢁ 105 Mꢀ1
,
9.8914 ꢁ 104 Mꢀ1 and
1.9189 ꢁ 105 Mꢀ1 for complexes 5-8 respectively and the Kb values
show that the complexes bind to DNA through the intercalation
[25]. As the DNA concentration is increased, the
p –
p⁄ transition
bands of this complex 5 exhibited a hypochromism of about
38.64% together with a hypsochromic shift of 4 nm and showed
only hypochromism (29.85%) without any shift respectively, at
277 and 405 nm. Whereas, the absorption band of complex 6 at
271 and 402 nm exhibited a hypochromism of about 37.29% and
25.11% with a hypsochromic shift of 3 nm and without any shift,
respectively. Similar observation is obtained for complexes 7 and
8 at 267, 358 and 273, 352 nm, respectively.
The observed binding constants revealed that the complex 5 has
higher magnitude of binding than complexes 6,7 and 8 with
CT-DNA and the order of binding affinity is 7 < 8 < 6 < 5. The incor-
poration of three methyl groups on the phenyl ring in the complex
6 hindered the intercalation of the complex and hence resulted in
lower Kb value. The electron withdrawing group in complex 7 has
lower binding value and the complex 8, which contains a benzyl
substituent, has a similar binding constant to complex 6. From this,
it is inferred that the increase in electron donating ability of the
substituent at the terminal nitrogen of the ligands increases the
DNA binding ability of the complexes. This may be due to the
increase in the electron density on the electron deficient metal
centers. These results suggested an intimate association of the
complexes with CT-DNA, and it is also likely that these complexes
bind to the DNA helix via intercalation [26].
3.3. DNA binding properties: absorption spectral studies
The interactions of metal complexes with DNA have been the
subject of interest for the development of effective chemothera-
peutic agents. Electronic absorption spectroscopy is one of the use-
ful techniques in DNA-binding studies. The binding of metal
complex to DNA grooves or intercalative p–p stacking interactions
has been characterized by the significant changes in the absor-
bance intensities (hypochromism) and appreciable shifts in wave-
length (blue or red shift) due to interactions between the planar
aromatic chromophore and the base pairs of DNA [22,23]. Any
interaction between the complex and DNA is expected to perturb
the ligand centered spectral transitions of the complex. Intensity
of the spectral band of the complexes (25
l
M) is found to decrease
M) and Figs. 5
with an increasing concentration of CT-DNA (0ꢀ25
l
and S13 . Metal complexes containing planar aromatic ligands
bind strongly to DNA by intercalation, while complexes with
non-planar ligands may bind to DNA through non-intercalative
modes. The complexes intercalate to the base pairs of DNA, the
3.3.1. Ethidium bromide displacement assay
The competitive DNA binding of complexes has been studied by
monitoring changes in emission intensity of ethidium bromide
(EtBr) bound to CT-DNA by the addition of complexes 5–8 as
quenchers. EtBr is a planar cationic dye can intercalate non-specif-
ically into DNA and it can form soluble complexes with nucleic
acids which causes it to fluoresce strongly. The emission intensity
of EtBr in buffer medium is enhanced due to its stacking interac-
tion of the planar phenanthridinium ring between the adjacent
DNA base pairs, eventhough it is quenched by the solvent
molecules and the fluorescence intensity is highly enhanced upon
addition of CT-DNA. When complex was added to DNA pretreated
with EtBr, the emission intensity was decreased due to competi-
tion by the complex for the EtBr intercalating sites. But at higher
binding ratios the intercalation is disrupted due to either DNA
bound strongly than EtBr or accepting an excited state electron
from EtBr and also the regularity of base stacking. The quenching
of emission intensity of DNA bound EtBr is occurred due to the
p⁄ orbital of the intercalated complexes could couple with
p
orbi-
tals of the base pairs, thus decreasing the
p ?
p⁄ transition ener-
gies, hence resulting in hypochromism [24]. DNA-binding
affinities of these complexes quantitatively, their intrinsic binding
constants with CT-DNA were obtained by monitoring the changes
in absorption at intraligand band with increasing concentrations of
DNA from the following equation:
½DNAꢂ=ðea
ꢀ
ef Þ ¼ ½DNAꢂ=ðeb
ꢀ
ef Þ þ 1=Kbðeb
ꢀ
ef
Þ
Where, [DNA] is the concentration of DNA in base pairs and the
apparent absorption coefficient ( a) is Aobs/[complex], ef is the
e
extinction coefficient of the free palladium complex and eb is the
extinction coefficient of the palladium complex bound to DNA,
respectively. The plot of [DNA]/(ea
and intercept which are equal to 1/(eb
ꢀ
e
f) versus [DNA] gave a slope
ꢀ
e
f) and 1/Kb(eb ef)
ꢀ

E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
7
Fig. 5. Absorbance quenching curves of complex bound to DNA: 5 (A) and 6 (B). [DNA] = 0–25 lM, [Complex] = 25 lM. (Inset: Plot of [DNA/(ea-ef) vs [DNA]).
planar rings in complexes can efficiently compete with EtBr for
intercalative binding sites on DNA by replacing EtBr. The changes
observed in the fluorescence spectra of EtBr on its binding to
CT-DNA are often used for the interaction study between DNA
and metal complexes. Quenching constant has been calculated
from the following SternꢀVolmer equation:
complexes 5–8 respectively. The fluorescence quenching spectra
of DNA-bound EtBr by complexes 5–8 shown (Fig. 6) illustrate that,
as the concentration of the complexes increases, the emission band
at 605 nm (545 nm excitation) exhibited hypochromism up to
35.01, 31.70, 29.14 and 30.31% with hypsochromic shift of 4, 3, 2,
2 nm of the initial fluorescence intensity for 5–8 respectively. The
observed decrease in the fluorescence intensity clearly indicates
that the EtBr molecules are displaced from their DNA binding sites
and are replaced by the complexes under investigation [27]. These
data suggests that the interaction of the palladium(II) complexes
with CT-DNA is stronger, which is consistent with the above absorp-
tion and emission spectral observations. Since these changes indi-
cate only one kind of quenching process, it may be concluded that
all of the complexes bind to CT-DNA via the same mode. Further-
more, such quenching constants and binding constants of the Pd(II)
complexes suggest that the interaction of all of the complexes with
DNA should be intercalation [28–30]. On the basis of all the spectro-
scopic studies, we concluded that the palladium(II) complex can
bind to CT-DNA in an intercalative mode and that the complex 5
binds to CT-DNA more strongly than the other complexes.
I0=I ¼ Kq½Qꢂ þ 1
Where, I0 is the emission intensity in the absence of a quencher,
I is the emission intensity in the presence of a quencher, Kq is the
quenching constant, and [Q] is the quencher concentration. The Kq
value is obtained as a slope from the plot of I0/I versus [Q]. The
quenching plots illustrate that the quenching of EtBr bound to
CT-DNA by the palladium(II) complexes are in good agreement
with the linear SternꢀVolmer equation. In the SternꢀVolmer plots
of I0/I versus [Q], the Kq values for complexes 5–8 were found to be
1.4608 ꢁ 104 Mꢀ1
,
3.8838 ꢁ 103 Mꢀ1
,
3.1326 ꢁ 103 Mꢀ1 and
3.8438 ꢁ 103 Mꢀ1 respectively. Further, the apparent binding
constant (Kapp) were calculated from the following equation:
KEtBr½EtBrꢂ ¼ Kapp½Cplxꢂ
(where the complex concentration has the value at a 50% reduc-
3.4. Protein binding studies
tion of the fluorescence intensity of EtBr, KEtBr = 1.0 ꢁ 107 Mꢀ1 and
[EtBr] = 10
l
M) were found to be 6.3174 ꢁ 105 Mꢀ1
,
It is a well known fact that BSA is the primary pharmacological
target of many antitumor complexes, and hence, the interaction
1.9789 ꢁ 105 Mꢀ1
,
1.5940 ꢁ 104 Mꢀ1 and 1.9784 ꢁ 105 Mꢀ1 for

8
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
3.4.1. Fluorescence quenching of BSA by complexes
Fluorescence quenching measurements have been widely used
to study the interaction of metal complexes with proteins such
as (BSA, HSA, lysozyme, etc.,) [31–33]. Fluorescence quenching is
the decrease in intensity of the fluorescence from a fluorophore
that occurs by various types of molecular interactions such as
ground state complex formation, excited reactions, molecular rear-
rangement, energy transfer and collision quenching. Changes in
molecular environment in the vicinity of fluorophore can be ac-
cessed by the change in fluorescence spectra in the absence and
presence of the complexes and hence provide valuable information
regarding the nature of the binding phenomenon. Generally, the
fluorescence of BSA is caused by two intrinsic characteristics of
the protein, namely tryptophan and tyrosine. Therefore, the intrin-
sic fluorescence of BSA can provide considerable information on
their structure and dynamics and is often utilized in the study of
protein folding and association reactions. The interaction of BSA
with all the complexes was studied by fluorescence measurement
at room temperature by Stern–Volmer relation. A solution of BSA
(1
lM) was titrated with various concentrations of the complexes
(0ꢀ50
lM). Fluorescence spectra were recorded in the range of
450–290 nm upon excitation at 280 nm. The effects of the com-
plexes on the fluorescence emission spectrum of BSA are shown
(Fig. 7). The addition of the above complexes to the solution of
BSA resulted in a significant decrease of the fluorescence intensity
of BSA at 335 nm, up to 55.7%, 46.9%, 37.5% and 44.7% from the
initial fluorescence intensity of BSA accompanied by a bathochro-
mic shift of 4, 3, 3 and 3 nm for complexes 5–8 respectively. The
observed red shift is mainly due to the fact that the active site in
protein is buried in a hydrophobic environment. This result sug-
gested a definite interaction of all of the complexes with the BSA
protein [34–36].
When a complex bind to set of equivalent sites on a protein, the
equilibrium binding constant and the number of binding sites can
be analyzed according to the Scatchard equation:
log½ðI0 ꢀ IÞ=Iꢂ ¼ logKbin þ nlog½Qꢂ
where, Kbin is the binding constant of the complex with BSA and n is
the number of binding sites, I0 and I are the fluorescence intensity in
the absence and presence of the complex. The value of Kbin, and n
can be determined from the plot of log(I0ꢀI)/I versus log [Q] as
shown in Fig. 8 and Table 3. The binding constant values indicate
that the complexes bind to BSA in the order of complex 5 > complex
6 > complex 8 > complex 7. The observed red shift can be correlated
with the binding of the complexes with protein that makes it more
hydrophobic environment. Complexes 5 and 6 possess a methyl
group while the other complexes 7 and 8 have bromo and benzyl
substitutions. Complexes 5 or 6 can interact with the active site
by making it more hydrophobic. The larger value of Kbin and Kq indi-
cates a strong interaction between the BSA and the complexes. The
calculated value of n is around 1 for all of the complexes, indicating
the existence of just a single binding site in BSA for all of the com-
plexes. Among the four Pd(II) complexes, the complex 5 has better
interaction with BSA than the other complexes 6, 7 and 8.
3.4.2. UV–Vis spectral studies
UV–Vis absorption measurement is a very simple and effective
method in exploring the structural change and detecting the
complex formation. Quenching mechanisms are usually classified
as either dynamic or static quenching. Dynamic quenching mech-
anism is refers to the excited state fluorescence molecule is influ-
enced by quenchers and the absorption spectra of the fluorescent
substance is not changed while, for static quenching, a formation
of complex between the ground state of the fluorescent substance
and quencher and therefore, the absorption spectra of fluorescence
substance would be considerably influenced. It is revealed that
Fig. 6. Fluoresence quenching curves of EtBr bound to DNA: 5 (A), 6 (B), 7 (C) and 8
(D). [DNA] = 10 lM, [EB] = 5 lM, [Complex] = 0–50 lM. (Inset: Plot of I0/I vs [Q]).
between BSA and metal complexes is of paramount importance in
understanding the mechanism.

E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
9
Fig. 8. SternꢀVolmer plots (A) and Scatchard plots (B) of the fluorescence titration
of complexes 5–8 with BSA.
Table 3
Quenching constant, binding constant and number of binding sites (n).
Complex
Kq (Mꢀ1
)
Kbin (Mꢀ1
)
n
5
6
7
8
7.90( 0.14) ꢁ 105
4.21( 0.25) ꢁ 105
5.65( 0.18) ꢁ 104
3.97( 0.09) ꢁ 105
6.43( 0.21) ꢁ 106
5.87( 0.32) ꢁ 105
6.42( 0.17) ꢁ 104
4.95( 0.04) ꢁ 105
1.05
0.98
0.87
0.92
This indicates that the interaction between BSA and complex 5 is a
static one and it was involved in the formation of ground state
complex of the type BSA-complex 5. But, dynamic quenching
affects only the excited state while it has no function on the
absorption spectrum. Therefore, the mechanism is only static
quenching and the same observed for BSA-complexes 6–8. A
similar type of static interaction between protein with complex
has been previously reported [38].
3.4.3. Characteristics of synchronous fluorescence spectra
According to Llyod, this technique provides information about
the molecular environment near the vicinity of the fluorophore
functional groups and also has advantages like spectral simplifica-
tion, spectral bandwidth reduction and avoid of different perturb-
ing effects. The synchronous fluorescence spectra of BSA are due to
presence of tyrosine and tryptophan residues and give information
on the conformational changes around tryptophan and tyrosine
region upon the addition of our complexes were measured before
and after the addition of complexes. Therefore, synchronous fluo-
rescence spectra are frequently used to characterize the interaction
Fig. 7. The emission spectra of BSA (1
complexes 5 (A), 6 (B), 7 (C) and 8 (D) (0–50
l
M) in the presence of increasing amounts of
lM).
there exists a static interaction between BSA and the added
complexes due to the formation of the ground state complex of
the type of BSA-compound reported earlier [37].
A simple method to explore the type of quenching is UV–Vis
absorption spectroscopy. UV–Vis spectra of BSA in the absence
and presence of the complexes (Fig. 9) show that the absorption
intensity of BSA was enhanced as the complexes 5–8 were added.
between fluorescence probe and proteins [39]. When
Dk of 15 nm
is used, the obtained synchronous fluorescence spectrum indicated

10
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
Fig. 9. The absorption spectra of BSA (1 lM) in the presence of complexes 5–8
(5 lM).
the spectral property of tyrosine residues, whereas
indicates that of tryptophan residues [40]. When the scanning
interval between excitation and emission wavelengths ( k) is sta-
Dk of 60 nm
D
bilized at 15 and 60 nm, the spectrum provides the particular
information of tyrosine and tryptophan residues in a protein,
respectively and the difference between the excitation wavelength
and emission wavelength (
D
k = kemi ꢀ kexc) indicates the type of
chromophores.The maximum emission wavelengths of tryptophan
and tyrosine residues in the protein molecule are related to the
polarity of their surroundings, changes of the maximum emission
wavelengths can reflect changes of protein conformation.
The effect of palladium(II) complexes on BSA synchronous fluo-
rescence spectroscopy with
Dk = 60 nm and Dk = 15 nm is shown
in Figs. 10 and 11, respectively. It has been observed that the addi-
tion of the complexes to the solution of BSA resulted in a signifi-
cant decrease of the fluorescence intensity of BSA at 340 nm, up
to 65.9, 58.1, 41.6 and 49.7% of the initial fluorescence intensity
of BSA accompanied with a bathochromic shift of 4, 3, 2 and
3 nm for complexes 5-8 respectively at the investigated concentra-
tion range when
Dk was equal to 60 nm. However, the addition of
the complexes to the solution of BSA resulted in a small decrease of
the fluorescence intensity of BSA at 304 nm, for complexes 5–8, the
decrease in the intensity of the above mentioned band of about
42.8%, 35.8%, 27.7% and 39.6% from the initial fluorescence inten-
sity of BSA was accompanied by a bathochromic shift of 2, 2, 1
and 2 nm, respectively at the investigated concentrations range
when
Dk was equal to 15 nm. The bathochromic shift of the max-
imum emission wavelength indicates that the conformation of BSA
was changed and the polarity around the tryptophan and tyrosine
residues was decreased, whereas the hydrophobicity was in-
creased [41]. The fluorescence intensities of both the tryptophan
and tyrosine were decreased along with red shifted with increasing
concentration of complex 5. It suggests that the interaction of com-
plex 5 with BSA affects the conformation of both tryptophan and
tyrosine micro-region. It reveals that the hydrophobicity around
tryptophan and tyrosine residues are strengthened. The results of
fluorescence and synchronous measurements of BSA confirm that
the binding of complexes with BSA is mainly due to the hydropho-
bic interactions. The order of binding strength of the complexes
with BSA is found to be 5 > 6 > 8 > 7. So, the strong interaction
between the complexes and BSA suggested that these complexes
can easily be stored in protein and can be released in desired target
areas. Thus, all the complexes exhibit a remarkable DNA and pro-
tein binding with high efficiency depending upon mode of binding
suggested that the complexes are potential candidates for further
biological studies such as antioxidant, antibacterial and anticancer
studies.
Fig. 10. Synchronous spectra of BSA (1
of complexes 5 (A), 6 (B), 7 (C) and 8 (D) (0–50
k = 15 nm.
l
M) in the presence of increasing amounts
lM) at a wavelength difference of
D
3.5. Antioxidant activity
Since the palladium complexes exhibit good DNA and protein
binding affinity, it is considered worthwhile to study the
antioxidant activity of these complexes. The radical scavenging

E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
11
indicated that the complexes showed antioxidant activity in the
order of 5 > 6 > 8 > 7 in all of the experiments. The complexes
5–8 displayed almost comparable radical scavenging activity with
respect to standard antioxidant (BHT). Further, the results obtained
against the different radicals confirms that the complexes are more
effective to arrest the formation of the ABTS and OH radicals than
that of DPPH and NO radicals studied. The palladium(II) complexes
exhibited nearly more than 50 times higher activity when
compared to the standards, which clearly indicates that the palla-
dium(II) chelation plays a vital role in determining the antioxida-
tive properties. Among the complexes, complex
5 shows
excellent antioxidant activity and this might be due to the more
electron donating nature and the planarity of the phenyl group.
The lowest activity was observed for complex 6 due to the steric
nature of trimethyl groups. So, the various antioxidant activity as-
say results clearly indicate that all the complexes are responsible
for the scavenging activity.
3.6. In vitro cytotoxic activity
As the palladium complexes bind DNA and protein, and show
antioxidant and antibacterial properties, the in vitro cytotoxicity
of the palladium complexes 5–8 was also evaluated against HeLa,
MCF-7 and NIH-3T3 cell lines using MTT assay after 48 hours of
inhibition. For comparison, the cytotoxicity of the known anti-can-
cer drug cisplatin has also assessed against all the above cell lines.
The results were analyzed by means of cell inhibition expressed as
IC50 values and the values of the four palladium thiocarboxamide
complexes 5–8 are listed in Table 4 and Fig. S14 . All the
complexes have higher cytotoxic activity against HeLa and the
MCF-7 cell lines used. Among the complexes studied here, complex
5 significantly had a stronger inhibitor effect on the proliferation of
HeLa and MCF-7 cells than cisplatin under similar experimental
conditions. This result suggests that complex 5 possess great selec-
tivity towards the both cancer cells and displays potential activity
towards HeLa and MCF-7 cancer cells with IC50 value 1.6 and
2.2
l
M respectively than cisplatin. A slightly lower inhibition cell
M) and MCF-7 (3.8 M) was observed
proliferation of HeLa (2.4
l
l
in the case of complex 6. Interestingly, complex 8 was found to
be more potent cytotoxic agent than the complex 7. The superior
activity of the complex 5 assumes significance in the light of the
fact that presence of electron donating methyl substituent and less
steric hindrance. Further, the activity may be correlated to the rel-
atively more electron rich center in the complex 5. The results of
the in vitro cytotoxic activity have also indicates that the IC50 value
of complex against NIH-3T3 (non-cancerous cells) is found to be
above 232 lM, which confirmed that the complex is very specific
for cancer cells compared to cisplatin. The IC50 values are much
better than those preciously reported for the other Pd complexes
Fig. 11. Synchronous spectra of BSA (1
of complexes 5 (A), 6 (B), 7 (C) and 8 (D) (0–50
k = 60 nm.
l
M) in the presence of increasing amounts
l
M) at a wavelength difference of
D
ability of the palladium complexes 5-8 along with standard BHT in
a cell free system against DDPH, OH, ABTS and NO radicals was
investigated with respect to different concentration of the test
complexes varying from 0 to 50
lM (Fig. 12). The IC50 values
Fig. 12. Trends in the inhibition of radicals by complexes 5–8.

12
E. Sindhuja et al. / Inorganica Chimica Acta 416 (2014) 1–12
Table 4
Appendix A. Supplementary material
The cytotoxic activity of the complexes.
Complex
aIC50 values (
HeLa
lM)
CCDC 841865, 900505, 830765 and 830764 contains the
supplementary crystallographic data for 5–8. These data can be
obtained free of charge from The Cambridge Crystallographic Data
Centre via www.ccdc.cam.ac.uk/data_request/cif. Supplementary
data associated with this article can be found, in the online version,
at http://dx.doi.org/10.1016/j.ica.2014.03.002.
MCF-7
NIH-3T3
5
6
7
8
1.66 0.8
2.46 2.3
18.82 0.7
11.21 1.6
12.6 3 0.9
2.21 1.9
3.88 0.6
23.99 1.7
8.98 2.5
13.86 0.4
232.88 2.4
235.63 0.6
256.72 1.4
244.64 2.0
243.94 1.9
Cisplatin
a
References
IC50 = concentration of the drug required to inhibit growth of 50% of the cancer
cells ( M).
l
[1] D. Wang, S.J. Lippard, Nat. Rev. Drug Discovery 4 (2005) 307.
[2] P.A. Andrews, J.A. Jones, Cancer Commun. 3 (1991) 93.
[3] M.E. Bianchi, M. Beltrame, G. Paonessa, Science 243 (1989) 1056.
[4] D. Bissett, K. McLaughlin, L.R. Kelland, R. Brown, Br. J. Cancer 67 (1993)
742.
[5] L.S. Lerman, J. Mol. Biol. 3 (1961) 18.
[6] W.O. Foye, Cancer Chemotherapeutic Agents, American Chemical Society,
Washington, DC, 1995.
[7] K.D. Demertzi, M.A. Demertzis, E. Filiou, A.A. Pantazaki, P.N. Yadav, J.R. Miller,
Y. Zheng, D.A. Kyriakidis, Biometals 16 (2003) 411.
[8] J.K. Willcox, S.L. Ash, G.L. Catignani, Crit. Rev. Food Sci. Nutr. 44 (2004) 275.
[9] I. Bjorkhem, A.H. Freyschuss, O. Breuer, U. Diczfalusy, L. Berglund, P.
Henriksson, Arterioscler. Thromb. Vasc. Biol. 11 (1991) 15.
[10] K.J.A. Davies, IUBMB Life 50 (2000) 279.
[11] A.C. Torbergsen, A.R. Collins, Eur. J. Nutr. 39 (2000) 80.
[12] M.N. Xanthopoulou, S.K. Hadjikakou, N. Hadjiliadis, E.R. Milaeva, J.A. Gracheva,
V.Y. Tyurin, N. Kourkoumelis, K.C. Christoforidis, A.K. Metsios, K. Karkabounas,
K. Charalabopoulos, Eur. J. Med. Chem. 43 (2008) 327.
[13] D.K. Demertzia, M.A. Demertzisa, J.R. Millera, C. Papadopouloub, C. Dodorou, G.
Filousis, J. Inorg. Biochem. 86 (2001) 555.
[42]. The result indicates that the palladium complexes 5, 6 and 8
possess great selectivity towards HeLa and MCF-7 cells and display
potential application in cancer chemotherapy and less toxic drug.
The overall cytotoxic behavior of the palladium thiocarboxamide
complexes is very similar to that of the DNA binding activity by
them discussed earlier, indicating better activity for complex
5.Thus, it emerges from our present study that binding of DNA is
one of the essential for a redox active drug to act as an anticancer
agent. Further, investigations are required to fully understand the
detailed molecular mechanism of cytotoxicity and hence verify
over hypothesis.
[14] S. Padhye, Z. Afrasiabi, E. Sinn, J. Fok, K. Mehta, N. Rath, Inorg. Chem. 44 (2005)
1154.
4. Conclusions
[15] M. Akaiwa, T. Kanbara, H. Fukumoto, T. Yamamoto, J. Organomet. Chem. 690
(2005) 4192.
[16] E. Sindhuja, R. Ramesh, Y. Liu, Dalton Trans. 41 (2012) 5351.
[17] M.H. Klingele, S. Brooker, Eur. J. Org. Chem. (2004) 3422.
[18] M.S. Blios, Nature 29 (1958) 1199.
[19] T. Nash, J. Biochem. 55 (1953) 416.
[20] D.C. Garrat, The Quantitative Analysis of Drugs, thire ed., Chapman and Hall
Ltd, Japan, 1964, p. 456.
[21] R. Re, N. Pellegrini, A. Proteggente, A. Pannala, M. Yang, C.R. Evans, Free Radical
Biol. Med. 26 (1999) 1231.
[22] Z.C. Liu, B.D. Wang, Z.Y. Yang, Y. Li, D.D. Qin, T.R. Li, Eur. J. Med. Chem. 44
(2009) 4477.
[23] Z.C. Liu, B.D. Wang, B. Li, Q. Wang, Z.Y. Yang, T.R. Li, Y. Li, Eur. J. Med. Chem. 45
(2010) 5353.
[24] A.M. Pyle, J.P. Rehmann, R. Meshoyrer, C.V. Kumar, N.J. Turro, J.K. Barton, J. Am.
Chem. Soc. 111 (1989) 3051.
[25] M.T. Carter, M. Rodriguez, A.J. Bard, J. Am. Chem. Soc. 111 (1989) 8901.
[26] V.A. Bloomfield, D.M. Crothers, I. Tinoco Jr., Physical Chemistry of Nucleic
Acids, Harper and Row, New York, 1974. p. 432.
[27] H.M. Torshizi, R. Mital, T.S. Srivastava, H. Parekh, M.P. Chitnis, J. Inorg.
Biochem. 44 (1991) 239.
[28] G.Y. Bai, K.Z. Wang, Z.M. Duan, L.H. Gao, J. Inorg. Biochem. 98 (2004) 1017.
[29] S.A. Tysoe, R. Kopelman, D. Schelzig, Inorg. Chem. 38 (1999) 5196.
[30] G.Y. Han, P. Yang, J. Inorg. Biochem. 91 (2002) 230.
[31] L. Messori, P. Orioli1, V. Vullo, L. Alessio, E. Iengo, Eur. J. Biochem. 267 (2000)
1206.
[32] S.J. Lau, B. Sarkar, J. Biol. Chem. 246 (1971) 5938.
[33] Y. Wang, X. Wang, J. Wang, Y. Zhao, W. He, Z. Guo, Inorg. Chem. 50 (2011)
12661.
A simple route to synthesis Pd(II) thiocarboxamide complexes
has been developed. The DNA binding properties of palladium(II)
complexes were examined by the UV–Vis spectra and fluorescence
spectra, which suggest their involvement in intercalative DNA
interaction with different binding affinities. The binding constants
showed that the DNA binding affinity increased in the order
7 < 8 < 6 < 5. The protein binding of the complexes was examined
by fluorescence spectra indicates the binding affinity, increases
based on the substitution in the phenyl ring at the terminal nitro-
gen of the thiocarboxamides. Experimental results reveal these
palladium complexes could bind to the serum albumins by high
affinity and quench the fluorescence of serum albumins through
a static quenching mechanism. All the complexes exhibit excellent
radical scavenging activities and the activity of complex 5 showed
greater IC50 values than the other. The cytotoxic study reveals that
the complexes 5, 6 and 8 show better cytotoxicty against HeLa and
MCF-7 cell lines with a potency significantlty higher than the
cisplatin and complex 5 shows excellent activity among them.
The present study highlights the importance of palladium(II)
complexes, which correlates with its potential to act as a novel
antitumor agent by inducing DNA binding. Further, confirms the
binding of the complex to DNA, which consequently leads to cell
death.
[34] H.Y. Shrivastava, B.U. Nair, J. Inorg. Biochem. 98 (2004) 991.
[35] K.Y. Zhang, S.P.Y. Li, N. Zhu, I.W.S. Or, M.S.H. Cheung, Y.W. Lam, K.K.W. Lo,
Inorg. Chem. 49 (2010) 2530.
[36] A.D. Tinoco, E.V. Eames, C.D. Incarvito, A.M. Valentine, Inorg. Chem. 47 (2008)
8380.
[37] S.S. Bhat, A.A. Kumbhar, H. Heptullah, A.A. Khan, V.V. Gobre, S.P. Gejji, V.G.
Puranik, Inorg. Chem. 50 (2011) 545.
Acknowledgments
[38] Y. Hu, Y. Yang, C. Dai, Y. Liu, X. Xiao, Biomacromolecules 11 (2010) 106.
[39] G.Z. Chen, X.Z. Huang, Z.Z. Zheng, J.G. Xu, Z.B. Wang, Methods of Fluorescence
Analysis, second ed., Science Press, Beijing, 1990.
[40] J.H. Tang, F. Luan, X.G. Chen, Bioorg. Med. Chem. 49 (2006) 3210.
[41] F. Samari, B. Hemmateenejad, M. Shamsipur, M. Rashidi, H. Samouei, Inorg.
Chem. 51 (2012) 3454.
One of the authors (E. S.) thanks University Grants Commission
(UGC), New Delhi, for the award of UGC-RFSMS. We thank
Dr. S. Abraham John, Gandhigram University, Dindigul, for
fluorescence spectral facilities. UGC and Department of Science
and Technology (DST), New Delhi, are greatly acknowledged for
Bruker Avance DPX 400 MHz NMR and Bruker SMART APEX II dif-
fractometer in the School of Chemistry, Bharathidasan University.
[42] S. Ray, R. Mohan, J.K. Singh, M.K. Samantaray, M.M. Shaikh, D. Panda, P. Ghosh,
J. Am. Chem. Soc. 129 (2007) 15042.