Morphological and: In vitro evaluation of programmed cell death in MCF-7 cells by new organoruthenium(II) complexes

Article abstract of DOI:10.1039/c7nj01707c

Cyclopentadienyl ruthenium(ii) thiosemicarbazone complexes with the general formula [Ru(η⁵-C₅H₅)(Ac-tsc)PPh₃]·Cl (1), [Ru(η⁵-C₅H₅)(Ac-mtsc)PPh₃]·Cl (2), [Ru(η⁵-C₅H₅)(Ac-etsc)PPh₃]·Cl (3) and [Ru(η⁵-C₅H₅)(Ac-ptsc)PPh₃] (4) were synthesized and characterized by various spectroscopic techniques (¹H NMR, ¹³C NMR, IR and UV-vis). The molecular structures of the representative complexes 2 and 4 were studied by single-crystal X-ray diffraction. The interactions of all the ligands and complexes with calf thymus DNA (CT-DNA) and bovine serum albumin (BSA) were studied using UV-vis and fluorescence emission spectroscopy. The results of binding studies revealed that the effective binding potentials of the complexes were higher than those of their parent ligands. All the new complexes 1-4 were investigated for their in vitro cytotoxic activity against MCF-7 human breast cancer cell line. All the complexes significantly inhibited cell proliferation in MCF-7 cells in a dose-dependent manner. Cytological observations via an inverted phase contrast microscope and a Hoechst 33342/PI dual-staining assay showed typical apoptotic morphology of cancer cells upon treatment with complexes 2 and 3. It can thus be suggested that the complexes 2 and 3 are modulated by apoptosis. The findings of the present study indicated that complexes 2 and 3 may become potent drugs for the treatment of cancer-related diseases only after further investigation.

Full text of DOI:10.1039/c7nj01707c

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Reyhaneh, F. Dallemer, R. Jayakumar, K. Palaniappan and R. Prabhakaran, New J. Chem., 2017, DOI:
1
0.1039/C7NJ01707C.
Volume 40 Number
1
January 2016 Pages 1–846
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PAPER
Jason B. Benedict et al.
The role of atropisomers on the photo-reactivity and fatigue of
diarylethene-based metal–organic frameworks
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Morphological and in vitro evaluation of programmed cell death in MCF-
cells by new organoruthenium(II) complexes
7
a
b
c
b
d
G. Devagi , F. Reyhaneh , F. Dallemer , R. Jayakumar , P. Kalaivani
,
a*
R. Prabhakaran
a
Department of Chemistry, Bharathiar University, Coimbatore 641 046, India
Department of Molecular Medicine,University of Malaya, Kuala Lumpur 50603, Malaysia
b
c
Lab MADIREL CNRS UMR 7246, Aix Marseille University, SaintꢀJerome Campus,
MADIREL Bldg., 13397 Marseille Cedex 20, France
d
Department of Chemistry, Nirmala College for Women, Bharathiar University, Coimbatore
641018
Abstract
Cyclopentadienylruthenium(II) thiosemicarbazone complexes, with the general
5
5
5
formula [Ru(η ꢀC
5
H
3
5
)(Acꢀtsc)PPh
3
].Cl (1), [Ru(η ꢀC
5
H
5
)(Acꢀmtsc)PPh
3
].Cl (2), [Ru(η ꢀ
5
C H )(Acꢀetsc)PPh ].Cl (3) and [Ru(η ꢀC H )(Acꢀptsc)PPh ] (4) were synthesized and
5
5
5
5
3
1
13
characterized by various spectroscopic techniques ( HꢀNMR, CꢀNMR, IR and UVꢀVis).
The molecular structure of representative complexes 2 and 4 was studied by single crystal Xꢀ
Ray diffraction. Interaction of all the ligands and complexes with Calf ThymusꢀDNA (CTꢀ
DNA) and Bovine Serum Albumin (BSA) was studied using UVꢀVis and Fluorescence
emission spectroscopy, the results of the binding studies cleared that the effective binding
potential of the complexes are higher than their parent ligands. All the new complexes (1-4)
were evaluated for their in vitro cytotoxic activity against MCFꢀ7 cancer cell line. All the
complexes significantly inhibited cell proliferation in human MCFꢀ7 breast cancer cells in a
doseꢀdependent manner. Cytological observations by an inverted phase contrast microscope
and Hoechst 33342/PI dualꢀstaining assay showed typical apoptotic morphology of cancer
cells upon treatment with complexes 2 and 3. It can thus be suggested that complexes 2 and 3
are modulated by apoptosis. The findings of the present study support that the complexes 2
and 3 can be of potent drugs for the treatment of cancer related diseases only after further
explorations.
Keywords: Organoruthenium(II) complexes, CTꢀDNA, Bovine Serum Albumin, Ethidium
Bromide, Anticancer activity, Apoptosis and Morphology
Corresponding author Tel.: +91ꢀ422ꢀ2428319; Fax: +91ꢀ422ꢀ2422387.
Eꢀmail address: rpnchemist@gmail.com (R. Prabhakaran)

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Introduction
There is considerable interest in metal based chemotherapeutic studies and number of
evidence demonstrating that metalꢀbased compounds are promising candidates for cancer
treatments. For instance, complexes possessing platinum, such as cisplatin, carboplatin and
oxaliplatin, have been used to treat several cancer types such ovary, stomach, and
1,2
colon. The mechanism of cancer cell growth inhibition by platinum complexes achieved
bycausing inter and intraꢀstrand crossꢀlinking with DNA, thus preventing DNA repair or
3
replication. However, former studies confirmed that platinum complexes have severe side
effects and generate resistant to cancer cells, limiting the efficacy of these complexes in
4
clinical trials. However, metal based complexes are the promising antiꢀcancer drugs due to
their ease of chemical modification and wideꢀspectrum of effectiveness against various
origins of cancer. ruthenium complexes are potent growth inhibitors for many cancer cells
5
ꢀ10
such as ovarian, melanoma, and breast. They have been used as an alternative to platinum
complexes for development of novel antiꢀcancer drugs with less or no side effects. Certainly,
11ꢀ13
several ruthenium complexes are under phase I or II clinical trials.
Based on the structureꢀ
activity relationship studies (SARS), ruthenium complexes may function to inhibit tumor
1
4
cells through mechanisms similar to that of cisplatin. Some ruthenium(III) complexes
1
5
(
NAMIA, KP1019 and KP1330) are in Phase II clinical trials. In addition, other
organometallic ruthenium (II) arene complexes, RM175 and RAPTA complexes, have also
16,17
shown promising activity.
The nature of the ligands bound to the metal ion is important
for the activity of the drug. Thus we chose thiosemicarbazide ligands, further they are a class
of Schiff base compounds showed different molecular geometry with metal complexes and
are familiar for their biological activity as antiparasitics, antibacterial, antifungal, antiviral,
18
and antitumoral agents. Cellular homeostasis is conserved via a tightly controlled apoptotic
1
9, 20
(programmed cell death) mechanism.
This is attained via two major pathways—the
extrinsic pathway that occurs through death receptors present in the outer membrane of the
2
1
cell, and the intrinsic pathway, which is a mitochondriaꢀdependent pathway. Critical defect
in the apoptotic signalling pathways may leads to uncontrolled proliferation and cell growth
which may ultimately lead to cancer, and the use of chemotherapeutic agents to induce
apoptosis in cancer is one of the effective ways to overcome this deadly disease. The
connection between apoptosis and cancer has been highlighted, with increasing evidence
suggesting that the linked processes of neoplastic transformation, progression and metastasis
2
2
involve the change of normal apoptotic pathways. Apoptosis afforded a number of hints

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with respect to effective anticancer therapy, and numerous chemotherapeutic agents
23
reportedly exert their antitumor effects by inducing apoptosis in cancer cells. Apoptosis not
only plays an important role in the development and maintenance of tissue homeostasis, but it
23,24
also represents an active mechanism by which harmful cells can be removed.
Typical
morphological types of apoptotic cells can be witnessed through microscopic studies using
the inverted phase contrast and fluorescence microscope. Other features such as chromatin
condensation and nuclear fragmentation can be better observed through the double staining
with Hoechst 33342 and propidium iodide using fluorescence microscopic analysis. This is a
convenient and rapid assay, widely used to identify live and dead cells. Hoechst 33342 is a
blue fluorescing dye that stains chromatin DNA. The red fluorescing dye propidium iodide is
only permeable to dead cells and cannot enter the intact plasma membrane of living cells.
Thus, the staining pattern resulted from the simultaneous use of these dyes make it possible to
2
5ꢀ28
distinguish normal, apoptotic, and dead cells population by fluorescence microscopy.
Also, it is highly desirable to have compounds that can cause cancer cell death via
apoptosis.Herein, the present study focused on synthesis and characterisation of new
cyclopentadienylruthenium(II) complexes and their cytotoxicity on human cancer cell line,
namely, the hormoneꢀdependent breast cancer cell line (MCFꢀ7). The presence of apoptosis
induced by complexes was then investigated through morphological observation of the cancer
cells using inverted phase contrast and fluorescence microscope (Hoechst 33342/PI). Further,
the apoptosis was confirmed with flow cytometry (Annexin VꢀFITC).
Measurements
Melting points were measured in a Lab India apparatus. Infrared spectra were
ꢀ1
measured as KBr pellets on a Jasco FTꢀIR 400ꢀ 4100 cm range. Elemental analyses of
carbon, hydrogen, nitrogen and sulfur were determined by using Vario EL III CHNS at the
Department of Chemistry, Bharathiar University, Coimbatore, India. (TMS).Electronic
absorption spectra of the ligands and complexes were recorded using JASCO 600
spectrophotometer and emission measurements were carried out by using a JASCO FPꢀ6600
1
13
3
spectrofluorometer. H and C NMR spectra were recorded in CDCl at room temperature
with a Bruker 400 MHz instrument, chemical shift relative to tetramethylsilane. The chemical
shifts are expressed in parts per million (ppm). CTꢀDNA, pBR322 plasmid DNA, BSA and
ethidium bromide (EB) were obtained from Himedia. Single crystal data collections and
corrections for the new Ru(II) complexes 2 and 4 were done at 293 K with CCD Kappa
2
9
Diffractometer using graphite mono chromated Mo Ka (k = 0.71073 A) radiation. The

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3
0
structural solutions were done by using SHELXSꢀ97 and refined by full matrix least square
3
1
on F2 using SHELXLꢀ2014.
EXPERIMENTAL SECTION
All the reagents used were analar grade, were purified and dried according to the
3
2
1–4
5
3 2
standard procedure. The ligands (HL ) and the ruthenium complex [RuCl(PPh ) (η ꢀ
33ꢀ35
5 5
C H )] were synthesized according to the standard literature procedures.
General Procedure for the Synthesis of ligands.
Preparation of acetonethiosemicarbazone.[H-Ac-tsc](HL ).
1
Thiosemicarabazide (1 mmol) and acetone (1 mmol) were combined in methanol (5
mL) with acetic acid (1−2 drops). The mixture was refluxed for 3 h, during which a white
precipitate appeared. After 3 h, the mixture was allowed to cool to room temperature, and
dried in vaccum. Ligands were further purified by recrystalliztion from methanol. Yield: 89
%
. Mp 162 °C.Anal.calcd for C H N S: C, 36.62; H, 6.91; N, 32.03; S, 24.44. Found: C,
4
9
3
ꢀ1
1
3
6.59; H, 6.88; N, 32.01; S, 24.40 %. FTꢀIR (cm ) in KBr: 1596 (ʋC=N), 863 (ʋC=S); H NMR
(CDCl
3
, ppm): δ 1.92 (s, 3H, ꢀC(CH
3
)
2
), δ 2.01 (s, 3H, ꢀC(CH
3
)
2
),δ 8.57 (s, 1H, ꢀNHꢀC=S),δ
6
.44 (s, 2H, terminal –NH ).
2
A similar method was followed to prepare other ligands.
2
Preparation of acetone-4(N)-methylthiosemicarbazone. [H-Ac-mtsc](HL ).
2
1
Ligand HL was prepared by the procedure as described for (HL ) with 4(N)ꢀ
methylthiosemicarbazide. Yield: 91 %. Mp 95 °C.Anal.calcd for C
5 11 3
H N S: C, 41.35; H,
ꢀ1
7
.63; N, 28.93; S, 22.08. Found: C, 41.30; H, 7.65; N, 28.89; S, 22.11 %. FTꢀIR (cm ) in
1
KBr: 1556 (ʋC=N), 850 (ʋC=S); H NMR (CDCl
3
, ppm): δ 1.88 (s, 3H, ꢀC(CH
),δ 8.38 (s, 1H, ꢀNHꢀC=S), δ 7.50ꢀ7.52 (d, 1H, J= 8 Hz, terminal –NH),δ 3.20ꢀ
.21 (d, 3H, J= 4 Hz, terminal –CH ).
3 2
) ), δ 1.99 (s,
3 2
3H, ꢀC(CH )
3
3
3
Preparation of acetone-4(N)-ethylthiosemicarbazone.[H-Ac-etsc](HL ).
3
1
Ligand HL was prepared by the procedure as described for (HL ) with 4(N)ꢀ
6 13 3
ethylthiosemicarbazide. Yield: 85 %. Mp 63 °C.Anal.calcd for C H N S: C, 45.25; H, 8.23;
ꢀ1
N, 26.39; S, 20.13. Found: C, 45.19; H, 8.20; N, 26.31; S, 20.09 %. FTꢀIR (cm ) in KBr:
1
1
637 (ʋC=N), 829 (ʋC=S); H NMR (CDCl
3
, ppm): δ 1.89 (s, 3H, ꢀC(CH
3
2
) ), δ 2.00 (s, 3H, ꢀ
C(CH
.21 (d, 3H, J= 4 Hz, terminal methylene protons), δ 3.16ꢀ3.17 (d, 3H, J= 4 Hz, terminal
methyl protons).
3 2
) ), δ 8.39 (s, 1H, ꢀNHꢀC=S), δ 7.47ꢀ7.53 (d, 1H, J= 24 Hz, terminal –NH), δ 3.20ꢀ
3

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4
Preparation of acetone-4(N)-phenylthiosemicarbazone.[H-Ac-ptsc](HL ).
4
1
Ligand HL was prepared by the procedure as described for (HL ) with 4(N)ꢀ
ethylthiosemicarbazide. Yield: 92 %. Mp 114 °C.Anal.calcd for C10
13 3
H N S: C, 57.94; H,
ꢀ1
6
.32; N, 20.27; S, 15.47. Found: C, 57.91; H, 6.28; N, 20.22; S, 15.45 %. FTꢀIR (cm ) in
1
KBr: 1592 (ʋC=N), 826 (ʋC=S); H NMR (CDCl
3
, ppm): δ 1.94 (s, 3H, ꢀC(CH
3 2
) ), δ 2.05 (s,
3
8
7
H, ꢀC(CH ) ), δ 9.28 (s, 1H, ꢀNHꢀC=S),δ 8.58 (s, 1H, terminal –NH), δ 7.18ꢀ7.20 (t, 1H, J=
3 2
Hz, terminal phenyl proton), δ 7.35ꢀ7.38 (t, 2H, J= 7.2 Hz, terminal phenyl protons), δ
.660ꢀ7.663 (d, 2H, J= 1.2 Hz, terminal phenyl protons).
Synthesis of new Ruthenium(II) complexes
Synthesis of [RuCp(Ac-tsc)(PPh
3
)].Cl (1).
5
3
To a solution of [RuCl(PPh
3
)
2
(η ꢀC
5
H
5
)] (0.20 g, 0.2752 mmol) in methanol (10 cm ),
3
acetonethiosemicarbazone [HꢀAcꢀtsc] (0.036 g, 0.2752 mmol) in methanol (10 cm ) was
added. The reaction mixture was then refluxed for 5 h. The reddish orange suspension
gradually turned to red color. The solvent was removed under reduced pressure. The orange
solidformed was washed with petroleum ether (60ꢀ80°C) 2ꢀ3 times. The complex was
crystallized from CHCl
3
/ nꢀheptane. Yield: 69 %. Mp 112 °C.Anal.calcd for
C H ClN PRuS: C, 54.49; H, 4.91; N, 7.06; S, 5.39. Found: C, 54.52; H, 4.89; N, 7.04; S,
27
29
3
ꢀ1
5
2 2
.37 %. FTꢀIR (cm ) in KBr: 1599 (ʋC=N), 839 (ʋC=S); UVꢀVis (CH Cl ), λmax: 242 (140,352)
3
−1
−1
3
−1
−1
nm (dm mol cm ) (intraꢀligand transition) and 321 (45,752) nm (dm mol cm ) (LMCT);
1
H NMR (CDCl , ppm): δ 2.10 (s, 3H, ꢀC(CH ) ), δ 2.29 (s, 3H, ꢀC(CH ) ), δ 4.37 (s, 5H, –
3
3 2
3 2
Cp protons), δ 12.51 (s, 1H, ꢀNHꢀC=S), δ 9.35 (s, 2H, terminal –NH
2
), δ 7.15ꢀ7.69 (m, 15H,
13
triphenylphosphine(aromatic region)). C NMR (CDCl
3
, ppm): δ 180.83 (C=S), δ 166.96
(
C=N), δ 78.12 (Cp), δ 22.36 (C(CH ) ), δ 128.30ꢀ135.49 (aromatic PPh ).
3
2
3
A similar method was followed to synthesize other complexes.
Synthesis of [RuCp(Ac-mtsc)(PPh )].Cl (2).
3
Complex 2 was prepared by the procedure as described for (1) with acetoneꢀ4(N)ꢀ
methylthiosemicarbazone [HꢀAcꢀmtsc] (0.039 g, 0.2752mmol). Orange solid formed was
washed with petroleum ether (60–80°C). The complex was crystallized from CHCl
heptane. Yield: 71 %. Mp 101 ⁰C.Anal.calcd for C28 PRuS: C, 55.21; H, 5.13; N,
.90; S, 5.26. Found: C, 55.19; H, 5.09; N, 6.87; S, 5.29 %. FTꢀIR (cm ) in KBr: 1580
3
/ nꢀ
H31ClN
3
ꢀ1
6
3
−1
−1
(ʋ
C=N), 837 (ʋC=S); UVꢀVis (CH
2 2
Cl ), λmax: 255 (216,963) nm (dm mol cm ) (intraꢀligand
3
−1
−1
1
transition) and 317 (118,229) nm (dm mol cm ) (LMCT); H NMR (CDCl
3
, ppm): δ 2.09

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3 2 3 2
(s, 3H, ꢀC(CH ) ), δ 2.25 (s, 3H, ꢀC(CH ) ), δ 4.35 (s, 5H, –Cp protons), δ 12.08 (s, 1H, ꢀNHꢀ
C=S), δ 9.72 (s, 1H, , terminal –NH), δ 3.007ꢀ3.019 (d, 3H, J= 4.8 Hz, terminal –CH ), δ
3
1
3
7
.15ꢀ7.41 (m, 15H, triphenylphosphine(aromatic region)). C NMR (CDCl
C=S), δ 173.10 (C=N), δ 83.66 (Cp), δ 26.40 (C(CH ), δ 31.30 (NHꢀCH ), δ 128.03ꢀ133.37
aromatic PPh ).
3
, ppm): δ 182.12
(
(
3
)
2
3
3
3
Synthesis of [RuCp(Ac-etsc)(PPh )].Cl (3).
Complex 3 was prepared by the procedure as described for (1) with acetoneꢀ4(N)ꢀ
ethylthiosemicarbazone [HꢀAcꢀetsc] (0.040 g, 0.2752 mmol). Reddish orange solid formed
was washed with petroleum ether (60–80°C). The complex was crystallized from CHCl
heptane. Yield: 73 %. Mp 92 ⁰C.Anal.calcd for C29 PRuS: C, 55.89; H, 5.34; N, 6.74;
S, 5.15. Found: C, 55.92; H, 5.30; N, 6.73; S, 5.09 %. FTꢀIR (cm ) in KBr: 1571 (ʋC=N), 841
3
/ nꢀ
H33ClN
3
ꢀ1
3
−1
−1
(
ʋC=S); UVꢀVis (CH Cl ), λmax: 254 (214,398) nm (dm mol cm ) (intraꢀligand transition)
2 2
3
−1
−1
1
and 309 (106,424) nm (dm mol cm ) (LMCT); H NMR (CDCl
C(CH ), δ 2.25 (s, 3H, ꢀC(CH ), δ 4.36 (s, 5H, –Cp protons), δ 12.03 (s, 1H, ꢀNHꢀC=S), δ
.78 (s, 1H, , terminal –NH), δ 3.42ꢀ3.50 (m, 2H, J= 4 Hz, terminal methylene protons), δ
.206ꢀ1.242 (t, 3H, J= 7.2 Hz, terminal –CH ), δ 7.17ꢀ7.37 (m, 15H,
, ppm): δ 179.66 (C=S), δ 164.60
C=N), δ 82.16 (Cp), δ 22.27 (C(CH ) ), δ 39.99 (NHꢀCH ), δ 32.45 (NHCH ꢀCH ), δ
3
, ppm): δ 2.10 (s, 3H, ꢀ
3
)
2
3 2
)
9
1
3
1
3
triphenylphosphine(aromatic region)). C NMR (CDCl
3
(
3
2
2
2
3
3
128.23ꢀ135.62 (aromatic PPh ).
Synthesis of [RuCp(Ac-ptsc)(PPh )] (4).
3
Complex 4 was prepared by the procedure as described for (1) with acetoneꢀ4(N)ꢀ
phenylthiosemicarbazone [HꢀAcꢀptsc] (0.056 g, 0.2752mmol). Orange solid formed was
washed with petroleum ether (60–80°C). The complex was crystallized from chloroform and
nꢀheptane. Yield: 69 %. Mp 107 ⁰C.Anal.calcd for C33
32 3
H N PRuS: C, 62.44; H, 5.08; N,
ꢀ1
6
.62; S, 5.05. Found: C, 62.39; H, 5.05; N, 6.58; S, 4.99 %. FTꢀIR (cm ) in KBr: 1539
3
−1
−1
(ʋC=N), 740 (ʋCꢀS); UVꢀVis (CH
2
Cl
2
), λmax: 264 (234,923) nm (dm mol cm ), 295
3
−1
−1
3
(
240,650) nm (dm mol cm ) (intraꢀligand transition) and 349 (108,653) nm (dm
−1
−1
1
mol cm ) (LMCT); H NMR (CDCl , ppm): δ 2.00 (s, 3H, ꢀC(CH ) ), δ 2.30 (s, 3H, ꢀ
3
3 2
C(CH
3 2
) ), δ 4.31 (s, 5H, –Cp protons), δ 5.29 (s, 1H, terminal –NH), δ 6.72ꢀ7.417 (m, 20H,
1
3
phenyl protons and triphenylphosphine(aromatic region)). C NMR (CDCl
3
, ppm): δ 176.31
(
C=S), δ 170.21 (C=N), δ 81.12 (Cp), δ 22.72 (C(CH ) ), δ 122.57ꢀ133.14 (aromatic and
3
2
3
PPh ).

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Binding studies
Various concentration (5–50 ꢁM) of Calf thymus DNA (CT DNA) were prepared by
dissolving in TrisHCl buffer with pH equal to 7.2. The Absorption spectra was recorded in a
JASCO spectrophotometer with 1 cm path length of the cuvettes after equilibrium attained at
20 °C for 10 min. The binding titration experiment has been carried out by fixing the
complex concentration as constant and by varying the DNA concentration. The absorbance
intensity (A) for each complex was recorded after the addition of DNA to the complexes.
Ethidium Bromide Displacement Assay
Emission intensity measurements were carried out by using a 5% DMF/ 5 mMTrisꢀ
HCl/ 50 mMNaCl buffer solution as a blank to make preliminary adjustments. The excitation
wavelength was fixed and the emission range was adjusted before measurements. DNA was
preꢀtreated with EB in the ratio [DNA/EB] = 10 for 30 min at 27 °C. The metal complexes
were then added to this mixture, and their effect on the emission intensity was measured.
DNA Cleavage study
The cleavage of DNA was checked using agarose gel electrophoresis. Supercoiled
pBR322 DNA (100 ng) in 5% DMSO and 95% Tris buffer (5 mM, pH 7.2) with 50 mMNaCl
was incubated at 37 °C in the absence and presence of compounds (50 ꢁM). The DNA,
compound, and sufficient buffer were premixed in a vial, and the reaction was allowed to
proceed for 2 h at 37 °C before the addition of ethylene glycol and loading onto an agarose
gel. Agarose gel electrophoresis of plasmid DNA was performed at 50 V in 1% slab gels
−
1
containing 0.5 ꢁg mL ethidium bromide in Tris buffer. DNA was visualized by
photographing the fluorescence of intercalated ethidium bromide under a UV illuminator.
The cleavage efficiency was measured by determining the ability of the compounds to
convert the supercoiled (SC) DNA to the nicked circular (NC) form and linear circular (LC)
form. After that the gel was documented on a Digital Gel Documentation system unit
(Syngene, USA).
BSA binding experiments
The protein binding study was performed by tryptophan fluorescence quenching
experiments using bovine serum albumin (BSA, 10 ꢁM) as the substrate in phosphate buffer
(pH 6.8). Quenching of the emission intensity of tryptophan residues of BSA at 347ꢀ349 nm
1
4
(
excitation wavelength at 280 nm) was monitored using ligands (HL -HL ) and complexes
–4 as quenchers with increasing complex concentration. The excitation and emission slit
1

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widths were 5 and 10 nm, respectively. Fluorescence measurements were performed using a
cm quartz cell on a JASCO F6500 spectrofluorimeter.
HSA-binding experiments
The protein binding study was performed by tryptophan fluorescence quenching
experiments using human serum albumin (HSA, 10 ꢁM) as the substrate in trisꢀHcl buffer
pH 7.2). A 3.0 mL portion of aqueous solution of HSA (10 ꢁM) was titrated by successive
1
(
1
4
additions of the ligands (HL -HL ) and complexes 1–4. For every addition, the mixture
solution was shaken and allowed to stand for 10 min, and then the fluorescence intensities
were measured with the excitation and emission wavelength set at 280 nm and 300–500 nm,
respectively. In the meantime, the synchronous fluorescence intensity of the mixture solution
was measured at ꢂλ = 60 nm and ꢂλ = 15 nm, respectively. For fluorescence enhancement
experiments, the Ru(II) complex (10 ꢁM) in 0.05 M Tris–HCl/0.15 M NaCl buffer of pH 7.2
was titrated with HSA. The fluorescence emission spectrum was recorded in the wavelength
range of 310–600 nm with an excitation wavelength of 300 nm.
In vitro evaluation of anticancer activity
Cell viability.
The 3ꢀ(4,5ꢀdimethylthiazolꢀ2ꢀyl)ꢀ2,5ꢀdiphenyltetrazolium bromide (MTT) assay is a
quantitative, sensitive and reliable colorimetric assay generally used for assessing viability
and proliferation of the cells. In this assay mitochondrial dehydrogenase enzymes of live cells
convert the yellow water soluble substrate, MTT into a purple formazan crystal product
36
which is insoluble in water or medium.
The cytotoxic potential of the compounds were assessed using a colorimetric MTT
reduction assay using MCFꢀ7 breast cancer cells and HaCaT(human immortalized, but ‘nonꢀ
cancerous’) cells. Briefly, cells were seeded in a 96ꢀwell flat bottom transparent plate at a
3
concentration of 7× 10 and incubated for 24 hours at 37 °C in an incubator supplied with 5%
CO
2
. After 24 hours, the cells were treated with different concentrations of compounds (3.12,
6.25, 12.5, 25, 50, 100 and 200 ꢁg/mL) and 0.1% DMSO was used as a vehicle control and
incubated for 24 h. After the incubation, a 50 ꢁl of MTT solution (2 mg/mL in phosphateꢀ
buffered saline) was added to each well and the plate was incubated for an additional 2h.Then
the supernatant in each well was removed and 100 ꢁL of DMSO was added to dissolve the
produced formazan crystal. Plates were then shaken for 15 min at room temperature for
complete solubilization. The absorbance was measured at 570 nm using a Tecan infinite
M1000Pro microplate reader (Tecan, Männedorf, Switzerland). The assay was performed in

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triplicate in 3 independent studies and the concentration of complexes with 50% reduction in
cellular viability was expressed as the half maximal inhibitory concentration (IC ) value.
50
Morphological changes of nucleus with Hoechst 33342/PI Staining
Cell death is discriminated into two main forms, apoptosis and necrosis. In contrast to
3
7
necrosis, apoptosis is a programmed and regulated pathway of cell death. Morphological
hallmarks of apoptosis in the nucleus are chromatin condensation and nuclear
3
8
fragmentation. This condensation and fragmentation can be followed by fluorescence
microscopy after staining with DNA binding fluorescence dyes. Hoechst and PI double
staining provide a fast and suitable assay for nucleus morphology and apoptosis. Hoechst
binds in the minor groove of double stranded DNA preferably AT rich region while PI
intercalates between the bases without any sequence preference. PI is normally used to
identify dead cells as it is membrane impermeable and gets excluded from viable cells while
Hoechst dye is permeable to the cell membrane and binds to DNA in live or fixed cells.
4
For this experiment, 8× 10 MCFꢀ7 cells were seeded into a 24ꢀwell plate. After
incubation, cells were treated with or without tested compounds at IC50 concentration and
further incubated for 24h at 37°C in an incubator supplied with 5% CO . After incubation, the
2
media was discarded and the cells were stained with Hoechst 33342 solution (10 ꢀg/mL). The
plate was further incubated for 10 min in the dark at 37°C. Then, the cells were counter
stained with propidium iodide (2.5 ꢀg/mL) for 5 min in the dark at room temperature and
then fluorescence was imaged by fluorescence microscope at 100 × magnifications.
Morphological Changes of MCF7 cells using phase contrast inverted microscope
Observation of morphological changes of apoptotic cells was performed according to
2
5
the method with slight modifications. Briefly, MCFꢀ7 cells were seeded into 24ꢀwell plate
and incubated overnight to attach. Then, the cells were treated with or without test
compounds(control) at different concentrations of 3µg/ml,6µg/ml and 12 µg/ml for complex
2
and 3.5µg/ml,7 µg/ml and 14 µg/ml for complex 3 was incubated 24h at 37°C with 5%
CO . The morphological changes of the cells were observed using an inverted light
2
microscope at 100X magnification.
Annexin V/PI double stainig assay
The procedure was conducted with Annexin VꢀFITC Apoptosis Detection Kit
5
(
eBioscience, USA) according to manufacturer’s protocol. Briefly, 2ꢀ5 × 10 cells were
seeded in 6ꢀwell plate and incubated overnight. The cells were treated with complex 2
6µg/ml and 12µg/ml) and Complex 3 (7µg/ml and 14µg/ml) compounds and incubated for
(

New Journal of Chemistry
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24 h at 37°C with 5% CO2.Then, the treated cells were harvested, washed with PBS and
resuspended in binding buffer. The cells were incubated for 10 minutes in dark with 5µg/ml
Annexin VꢀFITC in room temperature. After incubation, 10µg/ml of PI was added to the cell
suspension and the fluorescence of the cellswere measured by flow cytometry system (FACs
CANTO II).
Results and Discussion
5
The reactions of [RuCl(PPh
3
)
2
(η ꢀC
5
H
5
)] with an equimolar amount of various 4(N)ꢀ
1
4
substituted thiosemicarbazones (HL –HL ) in methanol resulted in the formation of new
complexes (Scheme 1), the analytical data of which confirmed the stoichiometry of the
complexes (1–4). The structure of the complexes 2 and 4 were confirmed by Xꢀray
crystallographic studies and attempts were made to grow single crystals of complex 1 and 3
in various organic solvents were unsuccessful. The complexes are soluble in common organic
solvents such as dichloromethane, chloroform, benzene, acetonitrile, ethanol, methanol,
dimethylformamide and dimethylsulfoxide.
S
S
Ph3P
H C
3
N
C
R
Ph3P
H C
N
C
N
N
3
N
N
Ru
H
H
Ru
S
.Cl
R
H3C
H
H
H3C
PPh3
, Methanol, 5 h
S
Ru
Cl
H C
3
H3C
H3C
N
C
N
,
Methanol, 5 h
C
C H
6 5
N
N
Ph3P
N
N
H
H
H
H3C
R = H, CH , C H
3
2
5
Complex 4
Complex (1-3)
Spectroscopic Studies
1
The IR and UV spectral details are given in the supporting information.The HꢀNMR
1
4
spectra of [HL ꢀHL ] showed a singlet at δ 8.38ꢀ9.28 ppm corresponding to (N(2)HꢀC=S)
39
group, in the complexes 1-3, a singlet occurred at δ 12.03ꢀ12.51 ppm due to (N(2)HꢀC=Sꢀ)
4
0
group indicating that the ligand remains in its thionic form. This shift showed that the
thiosemicarbazone is coordinated to the ruthenium through thione sulphur atom rather than
42,43
thiolate sulphur.
However, in complex 4 there was no resonance attributable to N(2)H,
indicating coordination of the thiolate sulphur ligand in the anionic form after deprotonation
at N(2). In complexes (1-4) a sharp singlet appeared at δ 4.31ꢀ4.37 ppm due to
4
4
1
cyclopentadienyl protons. A singlet observed in the ligand HL at δ 6.44 ppm due to
40
terminal (ꢀNH ) was appeared at 9.35 in the complex1. A doublet observed in the ligands
2
2
3
HL and HL around δ 7.47ꢀ7.53 ppm due to terminal (ꢀNH) was appeared in the complexes

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4
0
4
2
and 3 as a singlet at δ 9.72 ppm and δ 9.78 ppm respectively. Whereas, in HL and
complex 4, terminal (ꢀNH) was observed as a singlet at δ 8.58 ppm and 5.29 ppm
41
2
respectively. A doublet observed in the HL and complex 2 at δ 3.00ꢀ3.21 ppm due to the
3
presence of terminal methyl protons. For HL and complex 3 methylene protons of ethyl
group appeared as a doublet at δ 3.20ꢀ3.50 ppm and methyl protons of ethyl group appeared
4
5
1
4
at δ 1.20ꢀ3.17 ppm. Two singlets for each ligands (HL -HL ) and complexes (1-4) observed
40
13
1
at δ 1.88ꢀ2.05 and δ 2.1ꢀ2.3 ppm was due to methyl protons of acetone. The C { H} NMR
spectra of the complexes (1-4) contain resonances for the cyclopentadienyl ring carbons
44
around δ 78.12ꢀ83.66 ppm. The resonance observed around δ 164.60ꢀ173.10 ppm assigned
4
2
to the (C=N) carbon of the ligands. In complex 4, thioliccarbon signal of the ligand was
observed at δ 176.31 ppm whereas in the complexes (1-3) this signal was observed around
179.66ꢀ182.12 ppm as more deshieldedindicating the thione sulphur coordination to
46,47
ruthenium.
The spectra also showed resonance in the range of δ 122.57ꢀ135.62 ppm
42
corresponding to the aromatic carbons. In all the complexes resonances observed around δ
2.27ꢀ26.40 ppm was due to the methyl carbons of acetone group of ligand. In complex 2 a
signal observed at δ 31.30 ppm has been assigned to terminal HNꢀCH carbons. In complex
2
3
3
, signals observed at δ 39.99 ppm and δ 32.45 ppm were corresponding to the presence of
4
8
methylene and methyl carbons of ethyl group. The molar conductance for complex 1 and 3
ꢀ1
2
ꢀ1
ꢀ1
2
ꢀ1
was found to be 101 Ω cm mol and 109 Ω cm mol is found to be in agreement with 1:1
49
electrolytic behaviour.
X-Ray Crystallographic studies
Complexes 2 and 4 crystallized in monoclinic space group P21/c(Fig. 1 and 2,
2
Table 1). In complex2, the ligand [Hꢀ(Acꢀmtsc)] (HL ) is coordinated to ruthenium ion
through the N(3) nitrogen and thione sulphur atoms, forming a stable five member chelate
ring with a bite angle [N(3)−Ru(1)−S(1)] of 81.04(8)°. The Ru(1)−N(3) and Ru(1)−S(1)
distances found as 2.145(3) Å and 2.3651(10) Å respectively. The other sites are occupied by
triphenylphosphine Ru(1)−P(1) distances of 2.310(10) Å andacyclopentadienylligandwith
Ru(1)−C(6), Ru(1)−C(7), Ru(1)−C(8) Ru(1)−C(9) Ru(1)−C(10) distance of 2.157(4),
2.200(4), 2.208(4), 2.202(4), 2.160(4) Å respectively. The observed bond distances are
comparable with those found in other reported ruthenium complexes containing
55
triphenylphosphine and cyclopentadiene. In addition, complex 2 contains two intermolecular
hydrogen bonding through the hydrogen atom of the terminal nitrogen(N1) with the chlorine
atom(Cl2(A)) employing as charge compensating anion having N(1)−H(1)···Cl2(A) distance

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of 3.180 Å and a hydrogen atom of the(N2) imine nitrogen with the chlorine atom(Cl2(A))
having N(2)−H(2)···Cl2(A) distance of 3.141 Å (Fig. S14). In complex 4, the ligand [Hꢀ(Acꢀ
4
ptsc)] (HL ) coordinated to ruthenium in NS fashion by utilizing its, N1 nitrogen, and thiolate
sulphur atoms with the formation of stable five member ring with a bite angle
N(1)−Ru(1)−S(1) of 80.36(6)°. The Ru(1)−N(1) and Ru(1)−S(1) distances found as 2.137(2)
Å and 2.3522(7) Å respectively. The other site is occupied by triphenylphosphine with
Ru(1)−P(1) distance of 2.2847(2) Å and a cyclopentadienyl ligand with Ru(1)−C(35),
Ru(1)−C(36), Ru(1)−C(37), Ru(1)–C(38), Ru(1)−C(39) 2.157(3), 2.166(3), 2.204(3),
2.194(3), 2.197(3) Å respectively. The observed bond distances are comparable with those
found in other reported ruthenium complexes containing triphenylphosphine and
50
cyclopentadiene. The selected bond length and bond angles are given in Table. 2. Further,
there were no greater variations observed in the bond lengths and bond angles of complex 2
and complex 4.
3
Fig. 1.ORTEP for [RuCp(Acꢀmtsc)(PPh )].Cl (2)

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3
Fig. 2.ORTEP for[RuCp(Acꢀptsc)(PPh )] (4)

New Journal of Chemistry
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Table 1. Crystallographic data of complexes[RuCp(Acꢀmtsc)(PPh )].Cl (2) and [RuCp(Acꢀ
3
3
ptsc)(PPh )] (4)
[
RuCp(Ac-mtsc)(PPh )].Cl (2)
[RuCp(Ac-ptsc)(PPh )] (4)
3
3
CCDC No.
1525515
1525516
Empirical formula
C H Cl N PRuS
33 32 3
C H N PRuS
3
0
32
7
3
8
46.83
634.71
293 K
Formula weight
Temperature
293 K
Wavelength
0.71073 Å
Monoclinic
0.71073 Å
Monoclinic
Crystal system
Space group
P 2 /c
P 2 /c
1
1
1
1
1
9
9
9
2
4
1
0
1
0
3.5872(3) Å
1.6120(2) Å
8.5182(5) Å
0 °
a
15.5078(6)Å
8.1447(3) Å
29.4236(10) Å
90.0°
b
c
α
β
γ
9.189(2) °
0 °
93.075(3) °
90.0°
3
3
884.20(12) Å
Volume
3711.1(2) Å
Z
4
3
3
.462 Mg/m
Calculated density
Absorption coefficient
F(000)
1.516 Mg/m
ꢀ1
ꢀ1
.699 mm
304.0
.18 × 0.1 × 0.07 mm
1.051 mm
1708.0
Crystal size
Theta range for data
collection
0.18 x 0.18 x 0.08 mm
6.934 to 57.346
6.938 to 57.284
ꢀ
20 ≤ h ≤ 20, ꢀ10 ≤ k ≤ 10, ꢀ37 ≤ l ≤ ꢀ17 ≤ h ≤ 17, ꢀ14 ≤ k ≤ 15, ꢀ24 ≤
Limiting indices
3
8
l ≤ 23
Reflections collected /
unique
80452/ 8840 [R(int) = 0.0769]
64094 /6709 [R(int) = 0.0484]
Completeness to theta
Absorption correction
Max. and min. transmission
Refinement method
24.66° 99.59%
24.66° 99.63 %
Semiꢀempirical from equivalents
1.00000 and 0.72409
Semiꢀempirical from equivalents
1.00000 and 0.81326
2
2
Fullꢀmatrix leastꢀsquares on F
Fullꢀmatrix leastꢀsquares on F
6
709/0/352
.059
Data / restraints / parameters 8840/24 / 445
2
1
Goodnessꢀofꢀfit on F
Final R indices
1.070
R1 = 0.0598, wR2 = 0.1104
R = 0.0376, wR = 0.0845
1
2
[
I>2sigma(I)]
R = 0.0518, wR = 0.0911
R indices (all data)
R1 = 0.0899, wR2 = 0.1200
1
2
ꢀ3
ꢀ3
Largest diff. peak and hole
0.83 and ꢀ0.79 e.A
0.51 and ꢀ0.39 e.A

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Table 2.Selected bond lengths (Å) and angels (°) for Complex 2 and Complex 4
Bond lengths
Complex 2
2.3651(10)
2.3105(10)
2.145(3)
2.157(4)
2.200(4)
2.208(4)
2.202(4)
2.160(4)
Complex 2
92.92(3)
81.04(8)
95.06(8)
Bond lengths
Complex 4
2.3522(7)
2.2847(7)
2.137(2)
Ru(1)—S(1)
Ru(1) — S(1)
Ru(1) —P(1)
Ru(1) — P(1)
Ru(1) —N(3)
Ru(1) — N(1)
Ru(1) —C(6)
Ru(1) — C(35)
2.157(3)
Ru(1) —C(7)
Ru(1) — C(36)
2.166(3)
Ru(1) —C(8)
Ru(1) — C(37)
2.204(3)
Ru(1) —C(9)
Ru(1) — C(38)
2.194(3)
Ru(1) —C(10)
Bond Angles
Ru(1) — C(39)
2.197(3)
Bond Angles
Complex 4
91.26(2)
P(1)—Ru(1) — S(1)
N(3) —Ru(1) —S(1)
N(3) —Ru(1) —P(1)
P(1) —Ru(1) —S(1)
N(1) —Ru(1) —S(1)
N(1) —Ru(1) —P(1)
N(1) —Ru(1) —C(35)
N(1) —Ru(1) —C(36)
N(1) —Ru(1) —C(37)
N(1) —Ru(1) —C(38)
N(1) —Ru(1) —C(39)
C(35) —Ru(1) —S(1)
C(35) —Ru(1) —P(1)
C(35) —Ru(1) —C(36)
C(35) —Ru(1) —C(37)
C(35) —Ru(10)—C(38)
C(35) —Ru(1) —C(39)
C(36) —Ru(1) —S(1)
C(36) —Ru(1) —P(1)
C(36) —Ru(1) —C(37)
C(36) —Ru(1) —C(38)
C(36) —Ru(1) —C(39)
C(37) —Ru(1) —S(1)
C(37) —Ru(1) —P(1)
C(38) —Ru(1) —S(1)
C(38) —Ru(1) —P(1)
C(38) —Ru(1) —C(37)
C(38) —Ru(1) —C(39)
C(39—Ru(1) —S(1)
C(39) —Ru(1) —P(1)
C(39) —Ru(1) —C(37)
80.36(6)
95.04(6)
N(3) —Ru(1) —C(6) 155.01(16)
N(3) —Ru(1) —C(7) 117.48(15)
N(3) —Ru(1) —C(8) 95.45(15)
N(3) —Ru(1) —C(9) 107.06(17)
N(3) —Ru(1) —C(10) 143.02(18)
155.76(12)
144.52(13)
108.27(13)
96.90(12)
118.58(12)
122.06(11)
93.61(10)
36.79(14)
62.18(14)
61.65(14)
37.32(14)
93.97(10)
120.20(11)
37.70(15)
61.84(15)
62.04(14)
99.35(12)
155.64(10)
133.21(14)
135.27(14)
36.86(16)
36.83(15)
156.01(10)
100.85(11)
62.09(15)
C(6) —Ru(1) —S(1)
C(6) —Ru(1) —P(1)
121.60(15)
94.18(15)
C(6) —Ru(1) —C(7) 37.69(18)
C(6) —Ru(1) —C(8) 62.27(18)
C(6) —Ru(1) —C(9) 62.8(2)
C(6) —Ru(1) —C(10) 37.85(19)
C(7) —Ru(1) —S(1)
C(7) —Ru(1) —P(1)
154.85(13)
101.59(14)
C(7) —Ru(1) —C(8) 36.82(18)
C(7) —Ru(1) —C(9) 62.30(19)
C(8) —Ru(1) —S(1)
C(8) —Ru(1) —P(1)
C(9) —Ru(1) —S(1)
C(9) —Ru(1) —P(1)
131.21(14)
135.71(14)
97.18(14)
156.83(15)
C(9) —Ru(1) —C(8) 37.17(18)
C(10) —Ru(1) —S(1) 92.10(15)
C(10) —Ru(1) —P(1) 121.67(17)
C(10) —Ru(1) —C(7) 62.81(19)
C(10) —Ru(1) —C(8) 62.26(19)
C(10) —Ru(1) —C(9) 37.44(19)

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DNA binding studies
1
4
Addition of CTꢀDNA to ligands (HL -HL ) and the complexes (1-4), a increase in the
molar absorption was observed (Fig. 3 and Fig.S17). The absorption spectra of the complexes
at constant concentration (10 ꢁM) in the presence of different concentrations of CTꢀDNA (5–
1
4
5
0 ꢁM). The absorption spectra of the ligand (HL -HL ) mainly consist of only one resolved
1
4
band. The absorption spectra of ligands (HL -HL ) showed one resolved band in their
absorption spectra at 243 nm, 245 nm, 260 nm and 249 nm (intraligand transition)
respectively. While increasing the concentration of DNA all the ligand exhibited
hyperchromic red shift of 10 nm (A = 0.9008–1.8150), 10 nm (A = 0.5256–1.3154), 1nm (A
=
0.2367–1.5429) and 8 nm (A = 0.2783–1.3145).Whereas, the absorption spectra of
complex 1 consist of one resolved band centred at 259 nm (intraligand transition). As the
DNA concentration is increased, hyperchromism (A = 0.2599–2.0788) with a blueshift of 1
nm (up to 258 nm) was observed. The spectra of complex 2 consist of one resolved band
centred at 251 nm (intraligand transition). As the DNA concentration is increased,
hyperchromism (A = 0.1042–1.7081) with a redshift of 4 nm was observed in the intraligand
band. The spectra of complex 3 consist of one resolved band centred at 260 nm (intraligand
transition). As the DNA concentration is increased, hyperchromism (A = 0.1290–1.4745)
with a blueshift of 2 nm was observed in the intraligand band.Complex 4 exhibited
hyperchromism in the intraligand band at 263 nm (A = 0.2057–1.0546) with 4 nm blue shift.
The observed hyperchromic effect with a blue and red shift has suggested which has been
5
1,52
suggested to reside primarily at the outer electrophilic coat of CT DNA
predominantly via
electrostatic interaction with the negatively charged deoxyriboseꢀ phosphate backbone.
To identify the quantitative comparison of the DNA binding affinities, we obtained
the intrinsic binding constants,
CT DNA) by using the Stern Volmerequation :
DNA]/(ε ꢀε ) = [DNA]/(ε
where [DNA] is the concentration of DNA in base pairs,
coefficient obtained by calculating obs/[complex], ε corresponds to the extinction
coefficient of the complex in its free form, and ε
complex in the bound form. Each set of data, when fit to the above equation, gives a straight
line with a slope of 1/(ε ꢀ ε ) and a intercept of 1/ (ε ꢀ ε ), and was determined from
the ratio of the slope to intercept (Fig.3 and Table. 3). The intrinsic binding constants,
obtained for the complexes follow the order 2>3>4>1, suggesting that the complexes are
b
K , of the ligands and complexes for binding with calf thymus
(
[
a
f
b
f
ꢀε ) + 1/
K
b
(ε ꢀε )
b f
ε
a
is the apparent extinction
A
f
b
refers to the extinction coefficient of the
b
f
y
K
b
b
f
K
b
b
K ,

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involved in DNA binding. All the complexes showed better binding ability compared to that
if their corresponding ligands.
0
0
0
0
0
0
0
0
.014
.012
.010
.008
.006
.004
.002
.000
(
HL1)
(HL2)
(
(
(
(
(
(
HL3)
HL4)
1)
2)
3)
4)
0
10
20
30
40
50
[
DNA]
1
4
a f
Fig.3.Plot of [DNA]/{[DNA]/(ε ꢀε )} for ligand (HL -HL ) and complexes (1- 4).
b
Table 3.The binding constant K , for the interactions of ligands and complexes with CTꢀ
DNA
-
1
Complex Kb/M
1
5
HL
HL
HL
HL
1
0.83×10
2
3
4
5
5
5
5
6
5
5
1.63 ×10
2.10 ×10
2.35 ×10
3.71 ×10
1.96 ×10
9.52 ×10
5.36 ×10
2
3
4
Ethidium Bromide Displacement Assay
Further, the binding ability of the ligands and complexes with the CTꢀDNA has been
confirmed by Ethidium Bromide (EB) displacement studies (Fig. 5 and Fig.S18). Ethidium
Bromide competitive binding studies using ligands and new ruthenium(II) complexes (1-4) as

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quenchers may give further information about the binding of them to DNA. The quenching
extent of fluorescence of EB bound to DNA is used to determine the extent of binding of
metal complexes to DNA. Upon the addition of ligands and complexes (1-4) (0ꢀ50 ꢁM) to CT
DNA preꢀtreated with EB in a 5% DMF/5 mMTrisꢀHCl/50 mMNaCl buffer at pH 7.2, the
emission intensity at 607ꢀ 609 nm of DNAꢀbound EB decreases. Based on the classical Sternꢀ
Volmer equation the quenching constant has been analysed (Fig.5).
I
0
/I = Ksv[Q] + 1
From a plot of the observed intensities against the complex concentration, the values of the
53
apparent DNA binding constant (Kapp) were calculated using the equation
EB [EB] = Kapp[complex]
Where [complex] is the value at 50% reduction in the fluorescence intensity of EB, KEB
K
7
ꢀ1
(
1.0×10 M ) is the DNA binding constant of EB, [EB] is the concentration of Ethidium
Bromide (EB) (10 ꢁM). Kapp values are found in Table 4. From these experimental data, it is
inferred that the complex 2 replaces the EB more effectively than other complexes. From this
observation, it may conclude that all the complexes can bind to DNA through the
intercalation mode.
140
120
100
140
120
100
(1)
(2)
80
60
40
20
0
8
0
60
40
20
0
550
600
650
700
750
550
600
650
700
750
Wavelength (nm)
Wavelength (nm)
1
1
1
40
1
40
20
00
20
00
(
3)
(4)
1
1
80
60
40
20
0
8
6
4
2
0
0
0
0
0
550
600
650
700
750
550
600
650
700
750
Wavelength (nm)
Wavelength (nm)

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Fig.4.The emission spectra of the DNA–EB system (λexc = 515 nm, λem = 550–750 nm), in
the presence of complexes(1-4). [DNA] = 10 ꢁM, [Complex] = 0ꢀ50 ꢁM, [EB] = 10ꢁM. The
arrow shows the emission intensity changes upon increasing complex concentration.
1.22
1.20
1.18
1.16
1.14
1.12
1.10
1.08
1.06
1.04
1.02
1.00
(
(
(
HL1)
HL2)
HL3)
(HL4)
1)
2)
3)
(4)
(
(
(
0
20
40
60
80
100
[Q]
Fig.5.Stern–Volmer plots of the EB–DNA fluorescence titration for ligands and complexes
Table 4.Ksv, Kappvalues for the interactions of ligands and complexes with CTꢀDNA+EB
-
1
-1
Complex
K
sv/M
Kapp/M
1
3
3
3
3
3
3
3
3
6
6
6
HL
1.47 ×10
1.49 ×10
1.44 ×10
1.45 ×10
1.85 ×10
2.50 ×10
2.16 ×10
1.75 ×10
1.56 ×10
1.62 ×10
1.58 ×10
2
HL
3
HL
4
6
HL
1.50×10
1.53×10
1.51×10
1.57×10
1.50×10
6
1
2
3
4
6
6
6
DNA cleavage study
To assess the DNA cleavage ability of the free ligands and ruthenium(II) complexes,
supercoiled (SC) pBR322 DNA was incubated with complexes and ligands in 5
mMTrisHCl/50mM NaCl buffer at pH 7.2for2 h without addition of a reductant. Upon gel

New Journal of Chemistry
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DOI: 10.1039/C7NJ01707C
electrophoresis of the reaction mixture, a DNA cleavage was observed (Fig. 6). The relatively
fast migration is the intact supercoil form (Form I) and the slower migration is the open
circular form (Form II), which was generated from supercoiled when scission occurred on its
54,55
one strand.
The intensity of supercoiled SC (Form I) diminished and partly converted to
1
3
nicked form NC (Form II) by ligands (HL -HL ) and by complexes (1- 4). Whereas in ligand
4
(
HL ) the intensity of supercoiled SC (Form I) increased and partly converted to nicked form
NC (Form II). The intensity of the NC (Form II) band increases, whereas the production of
linear form LC (Form III) of DNA diminished. It is obvious that the ligands and
ruthenium(II) complexes has the ability to cleave the supercoiled plasmid DNA and this
cleavage system does not require the addition of any external agents.
Fig. 6.Gel electrophoresis diagram showing the cleavage of supercoiledpBR322 DNA
(
100ng) by ligands and complexes in 5% DMSO and 95% 5 mMTrisꢀHCl/50 mMNaCl buffer
1
4
at pH 7.2 and 37 °C with an incubation time of 2 h. Lane 1ꢀ4: Ligands (HL -HL ), Lane 5ꢀ8:
Complexes (1-4), Lane 9:buffer+plasmid, Lanes 10: plasmid alone, and Lane 11: DNA
ladder.
Protein binding studies
Tryptophan quenching measurements
The majority of intrinsic fluorescence of BSA when excited at 295 nm is provided by
tryptophan residues alone and other amino acid residues like tyrosine and phenylalanine
5
6
contribute to fluorescence only weakly. In order to determine the protein binding affinity of
the complexes, tryptophan quenching experiments were carried out by adding the complexes
in increasing concentration (5–50 ꢁM) to BSA (10 ꢁM) at room temperature and following
the decrease in fluorescence intensities. Generally, fluorescence quenching can be illustrated
57
by the wellꢀknown Stern–Volmer equation:
/Icorr = 1+ Ksv [Q]
I
0

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0
where I and I are the fluorescence intensities of BSA in the absence and presence of
the complexes respectively, Ksv is the Stern–Volmer quenching constant, [Q] is the
concentration of the quenching complex. The value of Ksv obtained as a slope of the linear
plot of I
0
/I vs. [Q] (Fig. 8). On increasing the concentration of the ligands and complexes
(
(
Fig. 7 and Fig. S19 ), uniform decrease in the fluorescence intensity by blue shift in ligands
1
4
HL ꢀ HL ) with 2 nm, complexes (1and3) with 3 nm and 1 nm respectively, also red shift in
complex 2 with 1 nm and the complex 4 with no shift was detected, which is due to the
increase in hydrophobicity of the microenvironment around tryptophan site in the
58
protein. Also, the linear fit obtained for Stern–Volmer quenching (Table. 5) analysis
represents a single quenching mechanism for all the complexes. However binding affinity of
complexes with BSA proportionately followed with binding ability observed for DNA
binding with complexes, 2>3>4>1.In order to investigate further whether the quenching
mechanism is static or dynamic, the quenching of tryptophan emission of BSA was studied at
room temperature. The absorption intensity of BSA was increased with the addition of the
complexes (1-4) (Fig. S20). The increasing intensity in the absorbance spectra for BSA +
59
complexes indicate the interaction of the complexes with BSA. It is well known that
dynamic quenching only affects the excited state of fluorophore and does not change the
absorption spectrum. However, the formation of nonꢀfluorescence groundꢀstate complex
induced the change in the absorption spectrum of fluorophore. Thus, possible quenching
mechanism of fluorophoreꢀquencher (BSA) by the complexes was found as static
60
quenching.
2
2
1
1
50
00
50
00
250
200
150
100
50
(1)
(2)
50
0
0
-
50
-50
-100
-150
-
100
150
-
300
350
400
Wavelength (nm)
450
500
300
350
400
Wavelength (nm)
450
500

New Journal of Chemistry
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DOI: 10.1039/C7NJ01707C
3
50
250
200
150
100
300
250
200
(4)
(3)
5
0
1
1
50
00
0
-50
50
-
-
100
150
0
300
350
400
Wavelength (nm)
450
500
300
350
400
450
500
Wavelength (nm)
Fig.7.The emission spectrum of BSA (10 ꢁM; λexc = 278 nm; λemi = 347 nm) in the
presence of increasing concentration of complexes (0 – 50 ꢁM). The arrow shows the
emission intensity changes upon increasing complexes concentration.
0
.4
2.5
2.4
2.3
2.2
2.1
2.0
1.9
1.8
1.7
1.6
1.5
1.4
1.3
1.2
1.1
1.0
0.9
(
(
HL1)
HL2)
0.2
0.0
(
(
HL1)
HL2)
(HL3)
(
(
HL4)
1)
(HL3)
(HL4)
-0.2
-0.4
-0.6
(2)
(3)
(
(
1)
2)
(4)
(3)
(4)
-
-
0.8
1.0
-1.2
1.4
-1.6
1.8
-
-
0
20
40
60
80
100
-
5.0
-4.8
-4.6
-4.4
-4.2
-4.0
[
Q]
Log[Q]
Fig. 8.Stern–Volmer plot
Fig.9. Scatchard plot
Determination of binding parameters and binding force
When static quenching interaction occurs, if it is assumed that the complex binds
independently to a set of equivalent binding sites in BSA, the binding parameters can be
61
determined according to the Scatchard equation:
log [(F −F)/F]= log
is the binding constant for the binding of the complex with BSA and n is the
number of binding sites per BSA molecule. By fitting the linear plot of log(F − F)/F) vs.
0
b
K + n log [Q]
where K
b
0
1
4
b
log[Q] (Fig. 9, Table 5) at room temperature, the K values for the ligands (HL -HL ) and
complexes (1–4) were calculated, which confirm that Ru(II) complexes having a large
hydrophobic area can interact efficiently than ligands with BSA through a static pathway.
Also, the value of n (1.1–1.6), which is approximately equal to 1, indicates that the binding
62
site in BSA is unique and accessible by the complexes. Synchronous fluorescence spectral

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DOI: 10.1039/C7NJ01707C
study was used to obtain information about the molecular environment in the vicinity of the
63
fluorophore moieties of BSA. Synchronous fluorescence spectra showed tyrosine residues of
BSA only at the wavelength interval ꢂλf 15 nm whereas tryptophan residues of BSA at ꢂλof
60 nm (Fig. 10 and Fig. S21). While increase the concentration of complexes (0–50 ꢁM)
added to BSA (10 ꢁM), a decrease in the fluorescence intensity in the tryptophan emission
maximum is observed for all the complexes(Fig. 10). In contrast, the emission intensity of
tyrosine residue increases without any change in the wavelength of emission. These
observations indicate that the test compounds did not affect the microenvironment of tyrosine
residues during the binding process significantly.
300
250
200
150
100
350
3
2
2
1
1
00
50
00
50
00
1
2
50
50
0
0
300
320
340
360
380
400
300
320
340
360
380
400
Wavelength (nm)
Wavelength (nm)
4
00
3
3
2
2
1
1
50
3
350
00
50
00
50
00
4
3
2
2
1
1
00
50
00
50
00
50
0
5
0
0
300
320
340
360
380
400
300
320
340
360
380
400
Wavelength (nm)
Wavelength (nm)
Fig. 10.Synchronous spectra of BSA (10 ꢁM) in the presence of increasing concentration
ofcomplexes1-4 for a wavelength difference of ꢂλ= 60 nm. The arrow shows the
emissionintensity changes upon increasing concentration of complex.

New Journal of Chemistry
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Table 5. Quenching constant (Ksv), binding constant (Kbin) and number of binding sites (n)
for the interactions of complexes (1–4) with BSA
-
1
-1
Complex
HL
K
b
/M
K
sv/M
n
1
2
3
4
4
4
4
4
4
6
6
4
3
3
3
3
3
3
3
3
0.23 ×10
0.63 ×10
0.50 ×10
0.03 ×10
3.44 ×10
3.97 ×10
1.12 ×10
9.32 ×10
5.54×10
1.28×10
8.84×10
6.42×10
9.17×10
1.14×10
1.51×10
8.62×10
0.92
0.93
0.95
0.67
1.16
1.61
1.49
1.25
HL
HL
HL
1
2
3
4
HSA-binding studies
The fluorescence titrations of the Ru(II) complex with HSA were carried out in Tris–
HCl buffer of pH 7.2. Fluorescence spectroscopy is an effective method used to explore the
interaction between small molecules and biomacroꢀmolecules. The fluorescence emission
spectra of HSA recorded in the range of 300–500 nm by exciting the HSA at 280 nm at
various concentration of the Ru(II) complex are shown in Fig.11. The HSA shows a strong
fluorescence emission with a peak at 342ꢀ343 nm, while ligands and the complexes has no
intrinsic fluorescence under the present experiment conditions. The fluorescence intensities
of the HSA decreased continually with increasing concentration of the complexes,
accompanied with a blue shift (from 342 to 340 nm), indicating that the binding of the
complexes to HSA changed the local microenvironment around the Trpꢀ214 residue in HSA,
and concomitantly the tertiary structure of the HSA. Commonly, fluorescence quenching can
64
be described by the following Stern–Volmer equation:
I
0
/I = 1+ Ksv [Q]
where I and I are the fluorescence intensities of HSA in the absence and presence of
0
the complexes respectively, Ksv is the Stern–Volmer quenching constant, [Q] is the
concentration of the quenching complexes. The value of Ksv obtained as a slope of the linear

Page 25 of 39
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DOI: 10.1039/C7NJ01707C
0
plot of I /I vs. [Q] (Fig.12). On increasing the concentration of the ligands and complexes
(
(
Fig. 11 and Fig.S22), uniform decrease in the fluorescence intensity by blue shift in ligands
1
4
HL ꢀ HL ) and complexes (1-4), which is due to the increase in hydrophobicity of the
microenvironment around tryptophan site in the protein. Also, the linear fit obtained for
Stern–Volmer quenching (Table. 6) analysis represents a single quenching mechanism for all
the complexes.
In order to investigate further whether the quenching mechanism is static or dynamic,
the quenching of tryptophan emission of HSA was studied at room temperature. The
absorption intensity of HSA was increased with the addition of the complexes (1-4) (Fig.
S23). The increasing intensity in the absorbance spectra for HSA + complexes indicate the
interaction of the complexes with HSA. It is well known that dynamic quenching only affects
the excited state of fluorophore and does not change the absorption spectrum. However, the
formation of nonꢀfluorescence groundꢀstate complex induced the change in the absorption
spectrum of fluorophore. Thus, possible quenching mechanism of fluorophoreꢀquencher
65
(HSA) by the complexes was found as static quenching.
6
4
2
0
0
0
0
5
0
0
(1)
(
2)
-
-
-
-
20
40
60
80
-
50
-
100
-
100
300
350
400
450
500
300
350
400
450
500
Wavelength (nm)
Wavelength (nm)
1
00
50
0
50
(
3)
(4)
0
-
50
-
50
-
100
150
-100
-
300
350
400
450
500
300
350
400
450
500
Wavelength (nm)
Wavelength (nm)

New Journal of Chemistry
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Fig. 11.The emission spectrum of HSA (10 ꢁM; λexc = 280 nm; λemi = 340 nm) in the
presence of increasing concentration of complexes (0 – 50 ꢁM). The arrow shows the
emission intensity changes upon increasing complexes concentration.
0
0
.3
.2
3.0
2.8
2.6
2.4
2.2
2.0
1.8
1.6
1.4
1.2
1.0
(
(
(
HL1)
HL2)
HL3)
0.1
.0
0
-0.1
-0.2
-0.3
(HL4)
1)
(2)
3)
(4)
(
(
-
0.4
0.5
0.6
0.7
-0.8
0.9
1.0
1.1
1.2
-1.3
1.4
-
-
(HL1)
(HL2)
(HL3)
-
(
(
(
HL4)
1)
2)
-
-
-
(3)
(4)
-
-
-
5.1
-5.0
-4.9
-4.8
-4.7
-4.6
-4.5
-4.4
-4.3
-4.2
10
20
30
40
50
[Q]
Log[Q]
Fig. 12.Stern–Volmer plot
Fig. 13. Scatchard plot
Determination of binding parameters and binding force
When static quenching interaction occurs, if it is assumed that the complexes binds
independently to a set of equivalent binding sites on a macromolecules, the binding
61
parameters can be determined according to the Scatchard equation:
log [(F −F)/F]= log Kb+ n log [Q]
0
where K
b
is the binding constant for the binding of the complexes with HSA and n is the
− F)/F) vs.
number of binding sites per HSA molecule. By fitting the linear plot of log(F
0
1
4
log[Q] (Fig.13, Table 6) at room temperature, the K values for the ligands (HL -HL ) and
b
complexes (1–4) were calculated, which confirm that Ru(II) complexes having a large
hydrophobic area can interact efficiently than ligands with HSA through a static pathway.
Also, the value of n (0.9–1.5), which is approximately equal to 1, indicates that the binding
site in HSA is unique and accessible by the complexes. Herein, binding affinity of complexes
with BSA proportionately followed with HSA, 2>3>4>1.
Table 6. Quenching constant (Ksv), binding constant (Kbin) and number of binding sites (n)
for the interactions of complexes (1–4) with HSA
-1
-1
Complex
K
b
/M
K
sv/M
n

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DOI: 10.1039/C7NJ01707C
1
2
3
4
4
4
5
4
4
4
4
4
HL
HL
HL
HL
1
2.70 ×10
3.26 ×10
1.28 ×10
3.79 ×10
3.25 ×10
2.82 ×10
1.48 ×10
2.90 ×10
0.98
1.39
1.12
1.05
1.22
1.58
1.36
1.38
5
1.39×10
8.93×10
3.72×10
1.87×10
4.27×10
6.74×10
4.49×10
4
4
5
6
5
5
2
3
4
4
1.00×10
Anticancer evaluation of compounds using MTT assay
To evaluate the cytotoxic effect of the ligands and complexes on the cell viability of
MCFꢀ7 cancer cells andHaCaT (human immortalized, but ‘nonꢀcancerous’) cells, MTT assay
was performed. The results from the triplicate MTT assays showed a significant inhibitory
effect for all the complexes (Fig.14). On the basis of this assay, complex 2 exhibited the
highest cytotoxic activity or lower IC50 value followed by complex 3, whereas complexes 4
and 1 displayed less activity. However ligands displayed no significant cytotoxic activity
against MCFꢀ7 cells.The IC50 values for the complexes 1, 2, 3 and 4 are 27.27 ꢁM (16.23±94
µg/ml), 10.67 ꢁM (6.5±0.19 µg/ml), 11.13 ꢁM (7.7±0.42 µg/ml) and 17.85 ꢁM (11.13±0.24
µg/ml) respectively. From the results obtained, we found that all the complexes (1-4) showed
an enhanced growth inhibitory effect than the recognized drug cisplatin with the IC50 value of
23.7 µM against MCFꢀ7 cell lines. Among the four complexes, 2 and 3 are the most effective
complexesagainst the MCFꢀ7 cell line. Hence, the complexes 2 and 3 have been taken
forapoptotic morphological investigation.While comparing the results of complexes reported
by Ferriet al, our complexes showed good cytotoxicity against the MCFꢀ7 and other cancer
66, 67
cell lines.
Further antiꢀcancer drugs with minimal side effects on normal cells are highly desirable for
68
therapeutic purposes. Hence, the current study also addressed the question of whether
synthesized complexes mediated suppressionof cell viability was selectively to cancer cells.
Nonꢀcancerous normal human keratinocyte line (HaCaT) or Human immortalized but non
cancerous cell line (HaCaT) is a spontaneously transformed aneuploid immortal keratinocyte

New Journal of Chemistry
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DOI: 10.1039/C7NJ01707C
69,70
cell line from adult human skin.
HaCaT cells are utilized for their high capacity
7
1
to differentiate and proliferate in vitro. Their use in research allows for the characterization
of human keratinocyte using a model that is reproducible and addresses issues such as short
culture lifespan and variations between cell lines that would otherwise be encountered. These
cells have allowed the characterization of several processes, such as their utilization as a
7
2
3
model system for vitamin D metabolism in the skin. Cytotoxicity study against human
immortalized, but ‘nonꢀcancerous’ cell line(HaCaT) indicated that the complexes (1-4)
possess less cytotoxic activity against ‘nonꢀcancerous’ cells (Fig. 15).Agents that are capable
of inducing selective apoptosis of cancer cells, without causing much harm to normal cells,
havereceived considerable interest in the development of novel cancer chemotherapeutic
7
3
drugs. We found thatconcentrations of complexes (1-4) were cytotoxic to human breast
cancer MCF7 cells were failed to induce apoptosis in HaCaT cells. These results suggest that
complexes (1-4) possess antiꢀcancer activities in human breast cancer cells withless cytotoxic
effects against human ‘nonꢀcancerous’ cells (HaCaT). Thus, these complexes extracts can be
evaluated further for potential anticancer properties.
1
00
(
(
(
(
(
(
(
3.12)
6.25)
12.5)
25)
50)
100)
200)
8
6
4
2
0
0
0
0
0
HL1
HL2
HL3
HL4
1
2
3
4
cisplatin
Fig.14.The % Cell viabilityfor the ligands, complexes and cisplatin.MTT(3ꢀ(4,5ꢀdimethylthiꢀ
azolꢀ2ꢀyl)ꢀ2,5ꢀdiphenyltetrazoliumbromide) assay.

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1
00
8
6
4
2
0
0
0
0
0
(6.25)
(
(
(
(
(
12.50)
25.00)
50.00)
100)
Control)
Complex 1
Complex 2
Complex 3
Complex 4
Fig. 15.The % Cell viability for the ligands, complexes against HeCaT cells (nonꢀcancerous
cell).
Morphological changes of nucleus with Hoechst 33342/PI Staining
Nuclear fragmentation is one of the prime features of apoptotic cell death. To
investigate the cell death by apoptosis and changes in the nuclear morphology, DNA binding
dyes Hoechst and PI were used to investigate nucleus morphology of the control and treated
cells (Fig.16). Hoechst is permeable to cell membrane and typically used to stain the nucleus
of live/dead cells. Hoechst binds to the minor groove of double strand DNA preferably at AT
rich regions and stains nucleus in blue colour. PI is usually used to identify the dead cells and
its membrane impermeablity and gets excluded from viable cells. PI binds to DNA by
intercalating between the bases without any sequence preference and gives red colour at
respective wavelength.
Morphological analysis of MCFꢀ7 cell nuclei with Hoechst 33342 and PI staining
after treating the cells with complexes 2 and 3 at IC50 concentration at 24 h, displayed
morphological alterations compared to untreated control cells. As shown in Fig.17, the
control cells stained with Hoechst 33342 dye with blue fluorescence represents all cells are
live and healthy, while the treated cells stained with Hoechst 33342 and propidium iodide
resulted in a red fluorescence or pink in the merged photo representing dead cells (late
apoptosis or necrosis), could be clearly visualized due to toxicity of the drugs and cell
membrane disruption.