Aminoacid-derivatized Cu (II) complexes: Synthesis, DNA interactions and in vitro cytotoxicity

Article abstract of DOI:10.1016/j.jorganchem.2017.04.017

Two different series of copper complexes, [Cu(MFL)(FcAA)H₂O] (C1-C4) and [Cu(MFL) (BzAA)H₂O] (C5-C8), where FcAA = ferrocenyl amino acid mannich base conjugates and BzAA = benzaldehyde amino acid mannich base conjugates have been syn

Full text of DOI:10.1016/j.jorganchem.2017.04.017

Accepted Manuscript
Aminoacid-derivatized Cu (II) complexes: Synthesis, DNA interactions and in vitro
cytotoxicity
Rinky Singh, P. Rama Devi, Sharmita S. Jana, Ranjitsinh V. Devkar, Debjani
Chakraborty
PII:
S0022-328X(17)30245-0
10.1016/j.jorganchem.2017.04.017
JOM 19904
DOI:
Reference:
To appear in: Journal of Organometallic Chemistry
Received Date: 23 January 2017
Revised Date: 10 April 2017
Accepted Date: 15 April 2017
Please cite this article as: R. Singh, P.R. Devi, S.S. Jana, R.V. Devkar, D. Chakraborty, Aminoacid-
derivatized Cu (II) complexes: Synthesis, DNA interactions and in vitro cytotoxicity, Journal of
Organometallic Chemistry (2017), doi: 10.1016/j.jorganchem.2017.04.017.
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ACCEPTED MANUSCRIPT
Aminoacid-derivatized Cu (II) complexes: Synthesis, DNA interactions and
in vitro Cytotoxicity.
Rinky Singh a , Sharmita S Janab , P. Rama Devi a, Ranjitsinh V Devkarb, Debjani
Chakrabortya*
aDepartment of Chemistry, Faculty of Science, The M .S .University of Baroda, Vadodara-390002, Gujarat,
India.
bDivision of Phytotherapeutics and Metabolic Endocrinology, Faculty of Science, The M .S .University of
Baroda, Vadodara-390002, Gujarat, India.
Email id: debchak23@gmail.com .Tel: +91-9537521703

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Aminoacid-derivatized Cu (II) complexes: Synthesis, DNA interactions and in
vitro Cytotoxicity.
Rinky Singh a , P. Rama Devi a, Sharmita S Janab, Ranjitsinh V Devkar b, Debjani Chakrabortya*
aDepartment of Chemistry, Faculty of Science, The M .S .University of Baroda, Vadodara-390002, Gujarat, India.
bDivision of Phytotherapeutics and Metabolic Endocrinology, Department of Zoology, Faculty of Science,
The M .S .University of Baroda, Vadodara-390002, Gujarat, India.
Email id: debchak23@gmail.com .Tel: +91-9537521703
Abstract
Two different series of copper complexes, [Cu(MFL)(FcAA)H2O] (C1-C4) and [Cu(MFL)
(BzAA)H2O] (C5-C8), where FcAA= ferrocenyl amino acid mannich base conjugates and
BzAA = benzaldehyde amino acid mannich base conjugates have been synthesized and
characterized by spectroscopic methods. The complexes have been investigated for their
interactions with DNA by employing fluorescence quenching measurements, UV–Vis
spectroscopy and DNA viscosity measurements. High binding constants obtained from the
DNA binding studies (Kb= 106 M-1) prompted the in-vitro cytotoxicity assay of complexes
on A549 human lung carcinoma cells (employing MTT assay). The IC50 values obtained
herein were found to be lower than those of the ligands for A549 cell line. Antiproliferative
effects on A549 tumour cells exerted by the complexes were consistent with their intracellular
uptake properties. The cellular uptake studies indicated that complexes C1-C8 enter the
cytoplasm and accumulate in the nuclei. Rapid change in the nuclear morphology was observed
with DAPI staining. Acridine orange/ethidium bromide dual staining revealed that most of the
A549 cells enter early apoptosis within 12 h of treatment. Further all the complexes showed
effective cell growth inhibition by triggering G0/G1 phase arrest and inducing apoptosis.
FACScan results revealed remarkably high percentage of cell death induced by the complexes in
the A549 cells, as compared to control. Annexin-V/PI staining of cells also indicated that the
complexes induce cell death through the apoptotic pathway. Our data here suggests that the
complexes could be good antitumor agents.
Key words: Copper complexes, ferrocenyl mannich bases, DNA Binding, DNA Cleavage,
A549 cells, Apoptosis.

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1. Introduction
Medicinal inorganic chemistry offers additional opportunities for the design of therapeutic agents
not accessible to organic compounds [1−3].The wide range of coordination numbers and
geometries, available redox states, thermodynamic and kinetic characteristic, and intrinsic
properties of the cationic metal ion and ligand itself offer the medicinal chemist a large variety of
reactivity to be exploited. The widespread success of cisplatin in the clinical treatment of various
types of neoplasia has placed coordination chemistry of metal-based drugs in frontline in the
fight against cancer [1, 4]. Although highly effective in treating a variety of cancers, the cure
with cisplatin is still limited by dose-limiting side effects [5] and inherited or acquired resistance
phenomena, only partially amended by employment of new platinum drugs [6−9]. These
problems have stimulated an extensive search and prompted chemists to develop alternative
strategies, based on different metals, with improved pharmacological properties and aimed at
different targets [10].
The ferrocene derivatives among the metallocenes are of importance for their stability in a
biological medium and for their lipophilic, nontoxic, and reversible redox properties [11]. The
cyclopentadienyl rings of ferrocene could be suitably functionalized, and ferrocenyl derivatives
are known to exhibit antitumor, antimalarial, and antifungal properties [11]. Seio et al. [12]
have reported new DNA binding molecules utilizing structural and conformational
properties of ferrocene. Their design concept is based on the fact that the distance
between the two cyclopentadienyl rings of ferrocene, ca. 3.3 Å is close to the distance
between two aromatic rings stacked with each other. In addition, it is well-known that
the minor groove of DNA can accommodate stacked aromatic rings, as established by the
structural studies of natural or synthetic molecules that recognize the minor groove.
Studies on the binding of ferrocenyl ligands with DNA has revealed binding constants in the
order of 103-105 M-1 [13] thus agreeing with the above mentioned fact. The stability of ferrocene
in aqueous and aerobic media has made ferrocenyl compounds very popular molecules for such
biological applications. Furthermore such favourable characteristics of ferrocene led to the
design of ferrocenyl derivatives that function as highly sensitive detectors of proteins or as
reporters of protein activity. Studies on ferrocenyl conjugates with amino acids and end-labelled
ferrocenyl di- and tripeptides have demonstrated distinctive electrical, structural, and medicinal
properties [14]. Many researchers have shown interest in design of unnatural ferrocenyl amino
acids and peptides which further have been studied for their biomedical applications.

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Modification of proteins by incorporating such unnatural ferrocenyl amino acids helps the study
of protein structure, activity and interaction with other biomolecules [15].
The ferrocenyl compounds are known to hold good potential as anticancer agents. This can
be exemplified by the fact that ferrocifen, the ferrocene-appended anticancer drug tamoxifen,
makes the drug effective against both hormone-independent and hormone-dependent breast
cancer cells [16]. Similarly, ferroquine, the ferrocene-attached antimalarial drug chloroquine, has
been found to be more effective than chloroquine [17]. The ability of ferrocenyl compounds to
cleave DNA may be responsible for these diverse properties [18–20]. Recent reports from our
laboratory have shown that Ru (II) complexes of amino acid conjugated ferrocene mannich bases
are efficient DNA and serum protein binders. Further, in-vitro investigations on A549 and MCF7
cell lines revealed their cytotoxicity and preference for apoptosis [21, 22].
Copper-based complexes have been investigated on the assumption that endogenous metals may
be less toxic for normal cells with respect to cancer cells. The altered metabolism of cancer cells
and differential response between normal and tumour cells to copper are the basis for
development of copper complexes endowed with antineoplastic characteristics. Copper is an
essential element for most aerobic organisms, employed as a structural and catalytic cofactor,
and consequently it is involved in many biological pathways [23-25]. Taking this into account,
much attention has been given to research on the mechanisms of absorption [26], distribution,
[27] metabolism, and excretion of copper [28, 29], as well as on its role in development of
cancer and other diseases [30, 31]. The fundamental aspects of the chemistry and biochemistry
of copper [32], the role of this metal in medicine and the pathology and treatment of Menkes and
Wilson diseases [33], and the chelation therapy approach in neurodegenerative disorders
characterized by accumulation of abnormal protein components mediated by copper [34] have
extensively been surveyed in recent review articles to which readers are addressed for a
comprehensive knowledge of the multifaceted functions played by this metal ion in physiology.
Comparing the extensive literature regarding the use of existing clinical drugs and
development of novel chemical entities based on platinum, the field of anticancer copper
complexes seems only at an infancy stage, even though it shows much promise for future
development. Actually, copper complexes offer the potential over platinum (II) complexes of
reduced toxicity, a novel mechanism of action, a different spectrum of activity, and the prospect
of non-cross-resistance. A recent review by Carlo Santini throws light on the various advances in
copper complexes as anticancer agents [35].

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Moxifloxacin (MFL) is a 4thgeneration fluroquinolone drug with a broad spectrum of
antibacterial properties and thus is an important option in the treatment of bacterial infections
[36]. During the past years, the antitumoral activity of MFL has raised much attention. In vitro
and in vivo studies have confirmed its anticancer effects to be associated with the inhibition of
mammalian DNA topoisomerase I, topoisomerase II, and DNA polymerase. Copper complex of
MFL has been found to bind CT-DNA via the intercalative mode and induce apoptosis in A-549
lung cancer cells by DNA damage [37, 38].
The foregoing facts stimulated our interest on the syntheses, structure, DNA binding,
cleavage and anticancer properties of mixed ligand Cu(II) complexes of amino acid conjugated
ferrocenyl / benzaldehyde mannich bases and the fluoroquinolone antibiotic moxifloxacin. These
ferrocenyl amino acids have been targeted as it has been shown that tethering biologically active
groups to ferrocenyl unit increases their potency possibly due to combined action of the organic
molecule with fenton chemistry of the Fe-centre [39]. The structures of the new complexes were
established on the basis of spectroscopic and electrochemical data. In addition, DNA cleavage
abilities and anticancer activities of the complexes have been rationalized in terms of their DNA
binding affinity. All the complexes show strong binding affinity to DNA, induce apoptosis and
act as a good antitumor agent in a dose & time dependent manner.
2. Experimental
2.1. Reagents and Materials
All the chemicals and solvents used for synthesis and characterization of ligands and
complexes are of analytical grade and were used as purchased. Ferrocene carboxaldehyde
and DAPI stain were purchased from Sigma–Aldrich. Amino acids, CT-DNA, Tri–sodium
citrate and EB (ethidium bromide) were purchased from SRL (Sisco Research Laboratory,
Mumbai, India). Cu (acetate) 2. 2H2O and BSA (bovine serum albumin) were purchased from
Hi media. All the solvents used in the present studies were purchased from Merck and are
of analytical grade. Moxifloxacin (MFL) was kindly donated by Alembic Research Centre
(Gujarat, India).
2.2. Methods and instrumentation
Infrared (IR) spectra (400–4000 cm-1) were recorded on Perkin Elmer RX-1 FTIR with samples
prepared as KBr disks. ESI Mass spectra of the ligands were recorded on Thermoscientific DSQ
– II Mass spectrometer and those of the complexes were recorded on Applied Biosystem API

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2000 Mass spectrometer. C, H and N elemental analysis were performed on a Perkin-Elmer
240B elemental analyser. UV–visible spectra were recorded in solution at concentrations in the
range 10-5–10-3 M on Perkin Elmer Lambda-35 dual beam UV-Vis spectrophotometer.
Fluorescence spectra were recorded in solution on JASCO FP-6300 fluorescence
spectrophotometer. EPR spectra at liquid nitrogen temperature were recorded on X-band EPR
spectrometer (IIT Mumbai). Fluorescence spectra were recorded in solution on JASCO FP-6300
fluorescence spectrophotometer. The % metal content in each of the complexes was determined
by complexometric/gravimetric titration method after decomposition of the complexes.
Thermogravimetric analyses were carried out with a model EXSTAR 6000. Cyclic voltammetric
studies were performed on CHI-600C electrochemical analyser in a three-electrode system
comprising of a glassy carbon working, platinum wire auxiliary, and saturated calomel reference
(SCE) electrode. Tetrabutylammonium perchlorate (TBAP) (0.1 M) was used as a supporting
Electrolyte in DMSO. Electrochemical data were uncorrected for junction potentials.
2.3. Synthesis of ligands
2.3.1. Synthesis of L1-L4
The ferrocenyl amino acids L1-L4 were synthesised according to the reported procedure [21].
The amino acids (5.0 mmol) and NaOH (5.0 mmol) in dry methanol (10 mL) were stirred for 30
min to get a homogeneous solution. A methanolic solution (10 mL) of ferrocene carboxaldehyde
(5.0 mmol) was added dropwise to the above solution, which was refluxed for 90 min, cooled,
and treated with sodium borohydride (10.0 mmol) with constant stirring. The solvent was
evaporated, the resulting mass was dissolved in water and acidified with dilute HCl, and the
solution pH was maintained at 5.6. The ligands precipitated as yellow solids were filtered,
thoroughly washed with water and cold methanol, and finally dried in vacuum over P4O10.

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SCHEME 1
2.3.2 Synthesis of L5-L8
The amino acids (2.0 mmol) were initially dissolved in dry methanol (20 mL) with addition of
NaOH (2.0 mmol) and continuous stirring. Benzaldehyde (2.0 mmol) was subsequently added to
the above solution. The mixture was refluxed for 1 h, cooled, and then treated with an excess of
solid NaBH4 with constant stirring. After stirring for∼15 min, the solvent was removed on rotary
evaporator and the resulting mass was dissolved in water followed by treatment with dilute HCl
to maintain a pH of∼5.5. White solids thus precipitated were filtered off, thoroughly washed
with water and cold methanol, and finally dried in vacuum over P4O10.
SCHEME 2

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2.3.3 Synthesis of complexes C1-C8
Complexes C1-C8 were prepared by a general synthetic procedure in which 1.0 mmol of
copper(II) acetate in 15 mL of methanol was reacted with moxifloxacin (MFL, 1.0 mmol) in
methanol while stirring at room temperature for 0.5 h followed by addition of solid fFc-AA/Bz-
AA (1.0 mmol) ligands in small portions with continuous stirring. The reaction mixture was
further stirred for 2 h, and the product was isolated as a green solid in∼75% yield. The solid was
washed with water and cold methanol, and finally dried in vacuum over P4O10 (scheme 1&2).
SCHEME 3

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SCHEME 4
Physicochemical data of the synthesized ligands and complexes have been provided
supplementary material.
2.4 DNA binding experiments
2.4.1. UV absorption studies
The interaction of compounds with CT DNA has been studied with UV spectroscopy in
order to investigate the possible binding modes to CT DNA and to calculate the binding
constants (Kb). Absorption studies were performed with fixed compound concentrations
while varying the CT-DNA concentration within. While measuring the absorption, equal
increments of CT-DNA were added at different ratios to both the compound solution and
the reference solution to eliminate the absorbance of CT-DNA itself.
2.4.2. Competitive binding studies with EB using fluorescence spectroscopy
The competitive binding study of each compound with EB has been investigated with
fluorescence spectroscopy in order to examine whether the compound can displace EB
from DNA–EB complex. The DNA–EB complex was prepared by adding 2.6 µM EB and 2.0

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µM DNA in tris buffer. The binding mode of complexes C1–C8 with CT-DNA was studied
by adding a certain amount of a solution of each complex step by step into the solution
of the DNA–EB complex. The influence of the addition of each complex has been
obtained by recording the variation in the fluorescence emission spectra of the DNA–EB
complex. The fluorescence intensities of EB bound to CT-DNA were measured at 609 nm
(524 nm excitation) after addition of different concentrations of the complexes at
different ratios.
2.4.3. Competitive binding studies with DAPI using fluorescence spectroscopy
The competitive binding study of C1–C8 with known groove binder DAPI has been
investigated with fluorescence spectroscopy. The DNA–DAPI complex was prepared by
adding DAPI (5 µM) and DNA (20 µM) in tris buffer. The binding mode of complexes
C1–C8 with CT-DNA was studied by adding a certain amount of a solution of each
complex step by step into the solution of the DNA–DAPI complex. The influence of the
addition of each complex has been obtained by recording the variation in the fluorescence
emission spectra of the DNA–DAPI complex. The fluorescence intensities of DAPI bound
to CT-DNA were measured at 451 nm (340 nm excitation) after addition of different
concentrations of the complexes at different mixing (r) ratios.
2.4.4. Viscosity measurements
Cannon–Ubbelohde viscometer maintained at a constant temperature of 32.0 ± 0.1 °C in a
thermostat was used to measure the relative viscosity of DNA (200 µM) solutions in the
presence of complexes C1–C8 (with [complex]/[DNA] ratio of 0, 0.04, 0.08, 0.12, 0.16,
0.20 and 0.24) in Tris–HCl buffer (pH 7.2). Digital stopwatch with least count of 0.01 s.
was used for flow time measurement with accuracy of ±0.1 s. The flow time of each
sample was measured three times and an average flow time was calculated. Data are
1/3
presented as (η/η0) versus [complex]/[DNA], where h is the viscosity of DNA in the
presence of complex and h0 is the viscosity of DNA alone. Viscosity values were
calculated from the observed flow time of DNA-containing solutions (t) corrected for that
of the buffer alone (t0), η = (t - t0)/t0.
2.5. Nuclease activity
The solution containing metal complex was taken in a clean Eppendorf tube and 1µg of plasmid
pBR322 DNA was added. The contents were incubated for 30 minutes at 37 °C and loaded on

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0.8% agarose gel after mixing 3µl of loading buffer and 0.25% bromophenol blue + 0.25%
Xylene cynaol +30% glycerol. Electrophoresis was performed at constant voltage till the
bromophenol blue reached to the 3/4 of the gel. The gel was stained for 10 min by immersing it
in ethidium bromide solution (5 µg/ml) and then de-stained for 10 min by keeping it in sterile
distilled water. The plasmid bands were visualized by photographing the gel under a UV Trans
illuminator. The efficiency of DNA cleavage was measured by determining the ability of the
complex to form open circular (OC) or nicked circular (NC) DNA from its super coiled.
2.6. Cytotoxicity
A549 cells (5.0X103 cells well-1) were placed in 96-well culture plates (Tarson India Pvt.,
Ltd.) and grown overnight at 37°C in a 5% CO2 incubator. Compounds to be tested were
then added to the wells to achieve final concentrations ranging from 10 to 500 mg/ml.
Control wells were prepared by addition of culture medium without the compounds. The plates
were incubated at 37°C at 5% CO2 incubator for 24 h. Upon completion of the incubation,
MTT dye solution (prepared using serum free culture medium) was added to each well to
a final concentration of 0.5 mg/ml. After 4 h of incubation with MTT, the culture media
was discarded and the wells were washed with Phosphate Buffer Saline (Hi-Media, India
Pvt., Ltd.), followed by addition of DMSO to dissolve the formazan crystals so formed
and subsequent incubation for 30 min. The optical density of each well was measured
spectrophotometrically at 563 nm using Biotek-ELX800MS universal ELISA reader (Bio-
Tek instruments, Inc., Winooski, VT). The IC50 values were determined by plotting the
percentage viability versus concentration on a logarithmic graph and reading off the
concentration at which 50% of cells remained viable relative to the control. Each
experiment was repeated at least three times to obtain mean values.
2.6.1. Cellular uptake study
A549 cells (4.0X104 cells well-1) plated on cover slips, were incubated with C1 and alone for
different time intervals from 1 to 6 h, fixed with 4% paraformaldehyde for 10 min at room
temperature (RT) and washed with PBS. This was followed by treatment with Propidium iodide
for 10 min at RT. The cells were washed, mounted in 90% glycerol solution containing Mowiol,
an anti-fade reagent, and sealed. Images were acquired using confocal fluorescence microscope
(Carl Zeiss, Germany)

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2.6.2. AO/EB staining technique (induction of apoptosis)
A549 cells were grown in triplicates using a 12 well tissue culture plate and allowed to
acclimatize overnight. Next day, cells were treated with C1-C8 incubated for 12 h. After the
incubation, cells were stained with acridine orange and ethidium bromide dyes for 5 min
in dark and immediately washed three times with PBS. The cells were then suspended in
PBS and photographed on confocal LSM-710 fluorescence microscope.
2.6.3. Nuclear staining
Nuclear staining was performed by reported procedures. Briefly, the cells after exposure with
complexes for 12-16 h were washed with PBS and fixed in 3.7% paraformaldehyde for 10 min.
The fixed cells were then permeabilized with TBST [ 50 mM Tris-HCl (pH 7.4), 150 mM
NaCl, and 0.1% Triton X-100] for 5 min. Cells were washed with PBS and then DAPI
solution (10 µgmL-1 in PBS) was added and kept for 5 min. After several washings with PBS,
the cells were observed under a fluorescence microscope (Leica DM IL microscope with
integrated Leica DFC 320 R2 camera and IL50 image software)
2.6.4. FACScan analysis
A549 cells (5x106 cells/ml) plated overnight were treated with different concentrations of C1-C8
in DMEM. The cells were then cultured overnight, harvested and fixed using chilled 70%
ethanol. The cells were then treated with 50mg/ml RNase A for 6 h at 37 °C, stained with
propidium iodide staining solution (20mg/ml PI and in PBS) for 1 h at 4°C and analysed using
flow cytometry (FACSCanto, Beckton Dickenson) and the percent cell death as that of the
control population was determined.
To determine the pathway of cell death, A549 cells (1x106 cells/ml) were treated with IC50
values of C1-C8 in DMEM, cells were then cultured for 12 h in complete medium, harvested
and washed twice with chilled PBS at 4° C. The cells were re suspended in 100mL annexin-V
binding buffer (100 mM HEPES/ NaOH, pH 7.4 containing 140 mM NaCl and 2.5 mM CaCl2)
and stained with annexin-V FITC and PI. The cells were gently vortexed and incubated for 15
min at RT in the dark. After incubation, 400mL of binding buffer was added to the cells and
analysed immediately using flow cytometry.

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3.0 Results and discussions
3.1. Characterization
3.1.1. Mass spectrometry
The ESI-MS spectra of the complexes of the ligands L1-L8 and complexes C1–C8 showed
molecular ion peaks at m/z values equivalent to their molecular weights. The m/z values
of all the complexes are in well agreement with the proposed composition (Fig S3).
3.1.2. FTIR Spectroscopy
The FTIR spectra of the complexes C1–C8 displayed characteristic strong stretching
bands at 1518–1582 cm-1 and weaker bands at 1490–1514 cm-1 due to asymmetric and
symmetric carboxylate (COO-) stretches respectively, which were found as strong bands in
the fingerprint region at 1580–1612 cm-1 in the spectra of free mannich bases (L1-L8). The
distinct broad bands at ~3450 cm-1 owing to the O-H stretching of free carboxylic acid groups
found in the ligands are completely lost in the IR spectra of the complexes indicating
complexation of the ligands with Cu(II) via the carboxylate oxygen. Furthermore the medium
secondary amine N-H stretching bands in the region of 2921–2963 cm-1 observed in the
spectra of the free ligands, shifted to 2920–2930 cm-1 in the complexes
indicating
complexation via the nitrogen of secondary amine. Similarly the shifts in the pyridone
carbonyl ν(CO)MFL and the carboxylate ν(COO)MFL stretching frequencies of moxifloxacin in
complexes indicate the binding of these groups with the metal ion. All the important stretching
values have been tabulated in Table S1.
3.1.3. Electronic Spectra
The electronic absorption spectra of C1-C8 in DMSO solution were recorded in the region 200-
900nm. The electronic spectra of ferrocenyl ligands displayed intense absorption bands in
the UV region ascribed to π-π* intra ligand transition of the cyclopentadienyl rings of ferrocene
[40], which were observed to be blue shifted in the spectra of the complexes C1-C4 (Table
1).
Additional intra ligand transition bands corresponding to the MFL ligands were observed
at 290nm and between 335-340 nm for C1-C8 slightly shifted to longer wave length due to
coordination with Cu (II). All the complexes showed peaks in the region 400-420nm
corresponding to MLCT transitions and broad absorptions in the range 620-640 nm (Fig.S4)
attributed to d–d transitions for Cu (II) in square pyramidal geometry [41].

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The ESR spectra of C2 (Fig.1) and C7 were recorded in DMSO at 10 K using 90 KHz field
modulation and the ꢀ factors were quoted relative to the standard marker TCNE (ꢀ = 2.00277).
The EPR spectra of the complexes exhibit a good hyperfine splitting (Fig.1) and the
corresponding g||, g┴ and A|| values are tabulated in Table 2. The values (gII > g﬩> 2.0023) for the
complexes indicate that the unpaired electron of Cu (II) most likely resides in dx2-y2 orbital
2
having B1g as ground state and the complexes have distorted square pyramidal geometry [42].
The axial symmetry parameter (G>4) indicates that there are no spin exchange interactions
between the copper centres.
Table 1: Electronic spectral data of complexes C1-C8.
Compound
Intraligand charge transfer
transitions
CT
Nm
d-d Transition
nm
π-π*
205, 289, 339
406
417
402
409
402
402
402
402
628
631
619
627
629
630
637
632
C1
C2
C3
C4
C5
C6
C7
C8
206, 290, 337
205, 290, 338
204, 290, 336
216, 290, 338
205, 289, 337
206, 289, 339
204, 290, 338
Fig 1: ESR Spectra of C2 in DMSO.
EPR conditions: Temperature, 10K; microwave
power, 5.0 mW; Modulation amplitude, 1G;
microwave frequency, 9.1GHz.
Fig 2: Cyclic voltammogram of C2 in DMSO.
0.1 M TEAP at a scan speed of 50 mV s-1.
Inset: Cyclic voltammogram of C5 in DMSO
-0.1 M TEAP at a scan speed of 50 mV s-1.

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Table 2: ESR and cyclic voltammetry data of complexes C1-C8.
Compounds
C1
-
C2
2.23
C3
-
C4
-
C5
-
C6
-
C7
2.32
C8
-
g
||
g┴
-
2.09
-
-
-
-
2.12
-
A
|| (cm-1)
-
140×10-4
48×10-4
4.8
-
-
-
-
146×10-4
50×10-4
4.2
-
A┴ (cm-1)
G
-
-
-
-
-
-
Cu2+/Cu1+(mV)
Cu1+/Cu2+(mV)
∆Ep (mV)
Fc+/Fc (mV)
-450
-290
-160
480
-470
-440
-285
-165
485
-420
-250
-170
480
-380
-250
-130
-
-410
-255
-155
-
-400
-250
-150
-
-385
-245
-140
-
-290
-180
470
Complexes C1-C4 were redox active and displayed a quasi-reversible cyclic voltammetric
response near 480 mV vs SCE in DMSO 0.1 M TEAP assignable to the Fc+-Fc couple, where Fc
is the ferrocenyl moiety (Fig.2). There is a significant positive shift of∼50 mV in the Fe (III)/Fe
(II) potential in these complexes compared to that of only ferrocene (430mV vs SCE). The
complexes also showed a quasi-reversible cyclic voltammetric response in the range -380 to -470
mV assigned to the Cu (II)/Cu (I) reduction and Cu(I)/Cu(II) oxidation in the -250 to -290mV
range (Table 2).
Thermal stability and thermal behaviours of all complexes were studied by
°
thermogravimetric analysis in the temperature range 50–850 C in N2 atmosphere. The
thermal behaviour studies of the complexes were found to be similar. The TGA profiles over
°
the temperature range 30–250 C are usually due to loss of moisture, water of hydration
and coordinated water. The complexes have first decomposition stage in the range 200–250 °C.
This dehydration process probably is due to the loss of coordinated water. Above 250 °C,
complexes decompose in a gradual manner, which may be due to thermal degradation of
°
the organic ligands. The continuous loss of weights is observed up to 700 C, leaving CuO as
residue (Fig. S5).

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3.1 DNA binding studies
3.1.1 UV-Visible absorption titration
The presence of ground state interactions between the biological macromolecule DNA and
complexes under study have been detected using absorption spectroscopy. DNA can
provide three distinctive binding sites (groove binding outside of DNA helix along major
or minor groove, electrostatic binding to phosphate group and intercalation), a behaviour
important for the biological role of antibiotic and anticancer drugs in vivo [43]. The
binding efficiency of the metal complexes to DNA can be effectively investigated
employing electronic spectroscopy since the observed changes in the spectra may give
evidence of the existing interaction mode [44]. Any interaction between C1–C8 and DNA
is expected to perturb the ligand centred transitions of the compounds.
Binding
with
DNA
via
non-
intercalative binding modes, such as
electrostatic forces, van der Waals
interactions, hydrogen bonds and
hydrophobic interactions
generally
results in increase in absorption
intensity (hyperchromism) and blue shift
of the absorption bands upon increasing
the concentration of CT-DNA owing
to the degradation of the DNA
double helix structure. A blue shift may
also be attributed to improper coupling
of π* orbital of the ligand and π orbital
of the base pairs. This unstacking of base
Fig 3: Absorption spectra of C1 showing the increase in
absorption intensity on gradual addition of CT-DNA in 5
mM TrisHCl buffer (pH,7.2) at 25°C. Inset shows the plot of
[DNA]/(εA-εf) vs [DNA].
pairs (distortion in π orbital of base pairs and π* orbital of ligand) causes slight blue shift
(hypsochromic shift). Similarly reduction of face to face base stacking (exposed electrons)
induces enhancement of absorption intensity (hyperchromism). On the other hand intercalation
generally results in hypochromism and a red shift (bathochromism) of the absorption band
due to a strong stacking interaction between an aromatic moiety of the ligand and the
base pairs of the DNA. A red shift can be directly linked with coupling of π* orbital of
intercalated compounds with the π orbital of DNA base pairs, thus decreasing π-π* transition

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energy. On the other hand the coupling π orbital is partially filled by electrons thus decreasing
transition probabilities and concomitantly resulting in hypochromism [45].
Table 3: DNA binding constants (Kb) of
The UV spectra of the complexes C1-C8 (10-6 M),
ligands L1-L8 and complexes C1-C8
have been recorded in the absence and presence of
Compound
%
K
b
varying CT-DNA concentration (1–50 x 10-3 M)
within. The bands centred around 289–290 nm and
337-339 nm in the complexes showed significant
hyperchromism with red shift (Fig.5 representative
data for C1), speculative of primarily groove binding
nature of the complexes. It should be noted that the
exact mode of interaction with DNA cannot be
concluded only by UV spectroscopic studies and more
techniques should be combined in order to come to a
safe conclusion. The results from the UV titration
experiments suggest that all complexes can bind to CT
DNA [46]; in the cases where an hyperchromic effect
exists, a first evidence of binding to CT DNA probably
by groove binding may be suggested, while the
existence of a red-shift is a hint of a stabilization of the
DNA duplex provoked by such interaction [47].
Hypo/Hyper
chromism
18%
L1
L2
L3
L4
L5
L6
L7
L8
C1
C2
C3
C4
C5
C6
C7
C8
6.7×104
1.4×104
1.3×104
5.8×104
2.0×104
1.5×104
4.0×104
3.4×104
7.0×106
2.3×106
4.2×106
6.4×106
6.0×105
2.5×105
3.4×105
5.2×105
12%
19%
20%
15%
20%
17%
19%
36%
25%
30%
34%
39%
34%
30%
36%
The magnitude of binding strength to CT-DNA
may be determined through the calculation of binding constant Kb which is obtained by
monitoring the changes in the absorbance of the compounds with increasing concentrations
of CT-DNA. Kb is given by the ratio of slope to the y intercept in plots [DNA]/(εa− εf)
versus [DNA]. where εa = Aobsd/ [compound], εf is the extinction coefficient for the
unbound ligands and complexes, and ϵb is the extinction coefficient for the compounds in the
fully bound form. The Kb values of metal complexes showed 10–100 folds higher binding
efficacy compared to the ligands. In general, complexes C1–C4 exhibit stronger binding
interactions(106 M-1) than complexes C5–C8 (105 M-1) due to strong groove binding of the
ferrocenyl moiety (Table 3). Complexes C1 and C4 with tryptophan and tyrosine substituted
ferrocenyl moieties bound to the Cu (II) centre respectively show the higher binding
constant values due to additional hydrogen bonding interactions between the NH group of

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tryptophan and the –OH group of tyrosine with DNA nucleobases which are accessible both
in major groove and minor groove.
3.1.2 Fluorescence quenching Studies
To further examine the mode of interaction of the compounds with DNA, via
intercalation or groove binding, a competitive binding study with two dyes: EB and
DAPI have been carried out using steady state fluorescence spectroscopy. Ethidium
bromide (=3,8-diamino-5-ethyl-6-phenyl-phenanthridinium bromide) is a phenanthridine
fluorescence dye and is a typical indicator of intercalation, forming soluble complexes
with nucleic acids and emitting intense fluorescence in the presence of CT DNA due to
the intercalation of the planar phenanthridinium ring between adjacent base pairs on the
double helix [48]. Addition of a second molecule, which may replace EB and bind to DNA
via intercalation results in a decrease of the DNA-induced EB emission [49]. The
emission spectra of DNA–EB (λex= 546 nm, λem= 610) in the absence and presence of
increasing amounts of ligands and complexes have been recorded. Addition of the
complexes C1–C8 did not have any kind of effect on the emission intensity or nature of
the emission of DNA–EB complex, indicating that intercalation may not be the binding mode
for these compounds.
3.1.3 Competitive binding studies with DAPI
DAPI (4,6-diamidino-2-phenylindole) is a classical minor groove binder to DNA and binds
specifically to GC regions. The fluorescence of DAPI (5µM) increases approximately 30
times when 20-fold excess (base pairs) of DNA (20µM) is added to the solution of the
dye [50,51]. The fluorescence spectra of a mixture of DNA–DAPI solution with
increasing concentration of C1–C8 (0-100µM) have been recorded . The addition of
aliquots of the complexes cause an initial fluorescence enhancement (Fig.4, red lines) which
on further addition of greater amounts showed subsequent quenching of the DNA–DAPI
fluorescence (Fig.4, blue lines). The initial fluorescence enhancement can be attributed to
partial overlap of the electronic states of the quencher molecules (C1–C8) and DNA–
DAPI complex leading to partial stabilization of the ground state complex and an
increase in the value of the Franck–Condon factor. The fluorescence intensity is

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proportional to the overlap of vibrational wave functions and consequently to the Franck–
Condon factor.
A quencher molecule when approaches a DNA–DAPI complex, forms an intermediate
complex, which is temporarily more stable than the original DNA– DAPI complex. This
shifts the ground electronic state to left and down, and as a result the overlap between
the vibrational functions in the excited and ground electronic states increases due to the
Frank–Condon principle (greater the overlap between the vibrational functions of excited
and ground states of the complex, greater is the Franck–Condon factor), leading to the
increase in fluorescence signal. Further addition of the quencher leads to the complete
displacement of DAPI from the DNA helix and fluorescence depletion [21, 52]. This
phenomenon has been explained schematically in Fig.4. The quenching of the DNA–DAPI
fluorescence is conclusive of the fact that C1–C8 replace DAPI from the minor grooves of
DNA and get bound, indicating their preference for groove binding.
Fig 4: Emission titration of DNA-Dapi complex with Fig 4: Schematic presentation of DAPI displacement
increasing concentration of C1.
from DNA helix by quencher molecule followed by
Red lines = changes in 1st phase (increasing intensity); fluorescence quenching and corresponding energy
blue lines= changes in 2nd phase (decreasing intensity).
diagrams.
3.1.4 Viscosity Measurement
In order to further confirm the mode of binding of complexes C1–C8 to CT-DNA,
viscosity measurements of DNA solutions were carried out in presence and absence of
these complexes. The viscosity of DNA is sensitive to length changes and is regarded as
the least ambiguous and the most critical clues of a DNA binding mode in solution [53].
In general, intercalating agents are expected to elongate the double helix to accommodate
the ligands in between the base pairs, leading to an increase in the viscosity of DNA. In

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contrast, a complex that binds exclusively in the DNA grooves typically causes less
pronounced (positive or negative) or no changes in DNA solution viscosity [54].
The effects of the complexes C1–C8, the classical intercalator EB and the groove binder dapi
on the viscosities of CT-DNA solution are shown in Fig. 5. with increasing [complex] /
[DNA] concentration ratios no significant change in the relative viscosity of CT-DNA
solution was observed which ruled out intercalative binding mode of complexes and is
consistent with the DNA groove binding as indicated by DNA–DAPI competitive binding
studies.
.
Fig 5 : Effect of increasing amounts of the complexes C1 C8, dapi and
ethidium bromide(EB) on the relative viscosity of CT-DNA (200 mM) in
Tris–HCl buffer at 25 oC [Complex]/[DNA] = 0,0.04, 0.08, 0.12,0.16, 0.20
3.1.5 Nuclease activity
In order to explore the DNA cleavage activity of the complexes, pBR322 DNA (30µM) was
incubated with C1-C8 (10µM) in 5mM Tris–HCl / 50mM NaCl buffer solution (pH 7.2).The
DNA cleavage activity was assessed by the conversion of supercoiled form of DNA (Form I,
SC) to linear open circular DNA (Form II, OC) or nicked circular (Form III, NC). It was found
that all the complexes exhibited DNA cleavage by the conversion of SC Form (I) into OC Form
(II) (Fig. 6), which indicates that the complexes are involved in the cleavage through single
strand breaking. The control experiments with the ligands or CuCl2·2H2O or DNA alone do
not reveal any significant cleavage, As the complexes do not require an external agent like

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ascorbic acid, MPA, or H2O2 for cleavage, it is evident that DNA cleavage occurs via an
oxidative
only.
pathway
Fig 6: Photogenic view of interaction of pBR322 DNA (30 µg/mL) with 10 µM
complexes incubated for 2hours at 37 °C: Lane 1, DNA; Lane 2, DNA+C1; Lane 3,
DNA + C2; Lane 4, DNA + C3; Lane 5, DNA + C4; Lane 6, DNA + C5; Lane 7,
DNA +C6; Lane 8, DNA +C8.
4.0 Cytotoxicity studies
4.1 MTT assay
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay was done to test the
ability of compounds to inhibit cell growth and induce cell death in A549 cells (human lung
carcinoma). A549 cells
were treated with the
compounds at varying
concentrations
(15–
250µg/ml) for 12 h which
inhibited the growth of
A549 lung cancer cells
significantly in a dose and
time dependent manner
and
the
complexes
recorded 50–85% higher
cytotoxicity compared to
the ligands. The IC50
Fig.7: MTT data expressed as mean ±S.E.M. for n=3. Where, ***p<0.001
compared to Ligands (L1-L8) and ###p<0.001 compared to MFL. 50%
inhibition concentration (IC50µg/ml) for complexes C1–C8.
values shown in Fig.7 indicate the order of cytotoxicity as C7 < C6 < C8 < C5 < C3 < C2 < C4
<C1.The results reveal enhancement in the antiproliferative activity of the parent
fluoroquinolone ligand (MFL) upon metal ion(Cu2+) conjugation giving credence to the
hypothesis that biological activity of such fluoroquinolones may partly be related to their metal

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chelating ability. Cell viability assay using control compounds showed that the ligands and the
metal salt alone were nontoxic to the cancer cells. Amongst the compounds examined presently,
C1 seems to be the most potent molecule.

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4.3 Cellular uptake study
The cellular uptake and localization of C1-C8, was studied by using confocal microscopy [55].
A549 cells were cultured on coverslips (corning22x22 mm), incubated for 24 h until they
reached 70% confluency. These were then serum starved overnight, followed by incubation with
C1-C8 (IC50), for 6h at 37 °C. Confocal microscopy imaging was carried out by staining with PI
(nuclear stain) to identify localization of the complexes and any nuclear disintegration. After
treatment for 6 h, bright blue fluorescence in the cytoplasm of the cells was observed under
confocal microscope (Fig. 8A), indicating the uptake of complexes by cells. Fig.8B shows
uptake of PI by nucleus only and Fig. 8C is merged image of Fig.8A and B, which confirms the
uptake and accumulation of complexes in cytosol of the cells. Cells not treated with complexes
exhibited negligible luminescence. Fluorescence intensity was found to increase with time
indicating increased internalization of the complexes. These results indicated that C1-C8 can
enter and accumulate in the cytosol of cells.
Fig 8: Confocal microscopic images of A549 cells treated with C1 and PI (A) blue emission of C1 (B) nucleus
stained with PI (C) Merged image of A and B (D) Bright field image of A549 cells treated with C1 for 6 hr at
37oC.
4.4 Induction of Apoptosis
Apoptosis induced by compounds is one of the considerations in drug development. The
apoptotic cells usually show characteristic apoptotic features such as nuclear shrinkage and
chromatin condensation. Apoptosis assay was carried out with staining methods using acridine
orange (AO) and ethidium bromide (EB). The AO/EB staining is sensitive to DNA and was used
to assess the changes in nuclear morphology. Apoptotic and necrotic cells can be distinguished
from one another using fluorescence microscopy. In the absence of C1, the living cells were
stained bright green (Fig. 9, control). After treatment of A549 cell line with C1 for a period of 12
h, green apoptotic cells containing apoptotic bodies, stained by acridine orange as well as

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ethidium bromide were found in addition to red necrotic cells stained by ethidium bromide (Fig.
9). Similar results were observed for all the complexes.
Furthermore, to determine the nuclear features and to gain insight into the pathway of cell death,
DAPI staining was carried out in the presence of C1-C8. The control cells exhibit light and
evenly stained contours of the nuclei in contrast to the treated cells that show typical
characteristics of cells undergoing apoptosis. The treated cells are seen to possess fragmented or
highly condensed nuclei while the bright field images provide evidence for cell shrinkage and
membrane blebbing attributed to the typical features of apoptotic cells (Fig. 9). Necrotic nuclei
are not observed with DAPI staining. The DAPI staining indicates apoptotic mode of cell death
in the presence of C1.Similar results were observed with all the other complexes.
Fig 9: Confocal images of A549 cells treated with C1 for 12 hr at 37oC in 5% CO2.
A) Staining with acridine orange/ ethidium bromide. B) Staining with DAPI.
4.5 Cell cycle arrest by flow cytometry
The effect of C1-C8 on the cell cycle of A 549 cells was studied by flow cytometry in propidium
iodide (PI) stained cells after treatment with complexes for 12 h. The representative DNA
distribution histograms of control and A549 cells in the presence of C1are shown in Fig.10.

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Cells go through the cell cycle in several well-controlled phases [56]. The entry into each phase
of the cell cycle is carefully regulated by different checkpoints. One of the major focuses of drug
discovery is to develop agents that target the cell cycle checkpoints that are responsible for the
control of cell cycle phase progression. Cell cycle analysis is used to detect and measure
apoptosis, a form of programmed cell death, by analysing cells with less DNA content ("sub-G1
cells")[56]. Such cells are usually the result of apoptotic DNA fragmentation. Apoptotic cells
often have fractional DNA content due to the fact that the fragmented (low MW) DNA
undergoes extraction during the staining procedure. Some cells also lose DNA (chromatin) by
shedding apoptotic bodies. Thus, only a fraction of the DNA remains within apoptotic cells.
Therefore, nuclei of apoptotic cells contain less DNA than nuclei of healthy G0/G1 cells,
resulting in a sub-G0/G1 peak in the fluorescence histogram that can be used to determine the
relative amount of apoptotic cells in a sample.
Flow cytometry was used to probe the mode of cell death and cell cycle arrest induced by the
complexes, and it was found that after 12 h of incubation the population of G0/G1, S and G2/M
cells decreased without significantly affecting the overall cell count (Fig. 10(a)). Moreover, the
percentage of DNA content (apoptotic cells) is increased in the Sub G0/G1 phase (Fig.10(a) &
(b)) reflecting a significant inhibition of cell cycle progression. Thus, the overall effect of C1
treatment appears to involve the prevention of cell division followed by cell death with a profile
indicative of programmed cell death (apoptosis). Similar results were obtained with complexes
C2-C8 (Fig. 10(b)).
Fig 10 (a): Effects of C1 on A549 tumour cells after 12 h, distribution among cell cycle phases
.

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Fig 10 (b): Cell cycle distribution on exposure of A549cells to complexes C1-C8 (IC50).
An apoptosis study using flow cytometery can be used to distinguish different cell types such as
viable cells, early apoptotic cells, and late apoptotic cells [57]. The measurement of annexin V
binding to the cell surface as an indicator for apoptosis is performed along with a dye exclusion
test to establish the integrity of the cell membrane. Annexin V, a calcium-dependent
phospholipid binding protein with high affinity for phosphatidylserine, binds to the
phosphatidylserine that has migrated outside the cell membrane. Propidium iodide (PI), a
viability stain which binds to the nucleus once the membrane has broken down, is used as an
indicator of membrane structural integrity.
A549 cells were incubated with C1-C8 (IC50) for 12 h, followed by staining with annexin and PI.
Data was collected by a flow cytometer. In the early stages of apoptosis, the cell membrane can
expose phosphatidylserine, which is annexin V-positive. Viable cells did not bind to annexin V
or PI (lower-left quadrant), early apoptotic cells bound to annexin V but excluded PI (lower-right
quadrant), and late apoptotic cells were positive for both annexin V and PI (upper-right
quadrant). The upper-left quadrant contains dead cells. The results shown in Fig.11 (a) & (b)
indicate that the control cells contain 3.59 % late apoptotic cells (upper-right quadrant). The
order of the percentage of late apoptotic A549 cells treated with complexes C1–C8 for 12 h is as
follows: C2 (21.3%) < C7(22.8%) < C5(27.4%) < C8 (27.6%) < C4(29.3%) < C6 (31.1%) <
C3(33.1%) < C (36.6 %). Thus it can be concluded that the complexes induce apoptosis in A-
549 cells.

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Fig 11 (a): Annexin V staining shows induction of apoptosis of A549 cells treated with C1 12 h. The percent of
apoptotic cells were detected by analysing Annexin V and PI binding with the help of flow cytometry. Viable
cells did not bind to Annexin V or PI (lower left quadrant D3), early apoptotic cells bound to Annexin V but
excluded PI (lower right quadrant D4), and late apoptotic cells were both annexin V- and PI-positive (upper right
quadrant D2), the upper left quadrant D1 contains the nuclear debris or dead cell.
Fig 11(b): Graphical representation of flow-cytometric analysis of apoptosis induced in A549 cells on treatment
with C1-C8 for 12 h. Data is represented in the form of percentage of apoptotic population.
5.0 Conclusion
Ternary Cu (II) complexes with ferrocene and benzene conjugated L-Amino acid mannich bases
and the antibiotic drug moxifloxacin were synthesized, characterized and their biological

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potential as cytotoxic agents was studied. Binding of these complexes to CT-DNA has been
investigated in detail by electronic absorption titration, steady state emission, and viscosity
studies. The results suggest that the complexes C1-C8 are efficient DNA groove binders. The
redox active complexes with quasi-reversible Fc+ /Fc (C1-C4) and Cu(II)/Cu(I) couples display
significant chemical nuclease activity (pBR322 DNA) in absence of any external agents. Anti-
proliferative effects on A549 tumour cells exerted by the complexes are consistent with their
intracellular uptake properties. The cellular uptake indicated that complexes C1-C8 enter the
cytoplasm and accumulate in the nuclei. Rapid change in the nuclear morphology with DAPI
staining and acridine orange/ethidium bromide dual staining reveals that most of the A549 cells
enter early apoptosis within 12 h of treatment. Further all the complexes show effective cell
growth inhibition by triggering G0/G1 phase arrest and inducing apoptosis. FACScan results
show remarkably high percentage of cell death induced by the complexes in the A549 cells, as
compared to control. Annexin-V/PI staining of cells also indicated that the complexes induce cell
death through the apoptotic pathway. Our studies suggest that the complexes are potential
anticancer molecules that may interfere with DNA replication. This work makes a significant
contribution to the virtually unknown chemistry of organometallic complexes as synthetic
nucleases.
Acknowledgement(s)
The authors acknowledge University Grants Commission, New Delhi, India for providing
financial support and Department of Science and Technology (DST) for DST INSPIRE
Fellowship. Authors are grateful to the Heads, Department of Chemistry and Department of
Zoology, The Maharaja Sayajirao University of Baroda, for providing with the necessary
laboratory facilities.

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References
[1] T. W. Hambley, Dalton Trans., (2007) 4929.
[2] C. Orvig, M. Abrams, J.Chem. Rev., 99 (1999) 2201.
[3] K. H. Thompson, C. Orvig, Dalton Trans., (2006) 761.
[4] F. Arnesano, G. Natile, Coord. Chem. Rev., 253 (2009) 2070.
[5] Y. W. Jung, S. Lippard, J. Chem. Rev., 107 (2007) 1387.
[6] T. Boulikas, A. Pantos, E. Bellis, P. Christofis, Cancer Ther., 5(2007) 537.
[7] S. van Zutphen, J. Reedijk, Coord. Chem. Rev., 249 (2005) 2845.
[8] R. Gust, W. Beck, G. Jaouen, H. Schoenenberger, Coord. Chem. Rev., 253 (2009) 2760.
[9] R. Gust, W. Beck, G. Jaouen, H. Schoenenberger, Coord. Chem. Rev., 253 (2009) 2742.
[10] P. C. A. Bruijnincx, P. J. Sadler, Curr. Opin. Chem. Biol., 12(2008)197.
[11] (a) F. Tisato, C. Marzano, M. Porchia, M. Pellei, C. Santini, Med. Res. Rev., 30 (2010)
708.
(b) G. Jaouen, A. Vessières and S. Top, Chem. Soc. Rev., 44 (2015) 8802.
[12] (a) K. Seio, M. Mizuta, T. Terada, M. Sekine, J. Org. Chem., 70 (2005) 1031.
(b) S. Tardito, L. Marchio, Curr. Med. Chem., 16 (2009) 1325.
[13] (a) R. Gul, M.K. Rauf, A. Badshah, S.S. Azam, M.N. Tahir, A. Khan, Eur. J. Med. Chem.,
85 (2014) 438-449.
(b) S. Hussain, A. Badshah, B. Lal, R.A. Hussain, S. Ali, M.N. Tahir, A.A. Altaf, J. Coord.
Chem., 67 (2014) 2148-2159.
(c) T.K. Goswami, S. Gadadhar, M. Roy, M. Nethaji, A.A. Karande, A.R. Chakravarty,
Organometallics., 31 (2012) 3010-3021.
[14] A.M. Gellett, P.W. Huber, P.J. Higgins, J. Organomet. Chem., 693 (2008) 2959.
[15] P.J. Higgins, A.M. Gellett, Bioorg. Med. Chem. Lett. 19 (2009) 1614.
[16] A. Nguyen, S. Top, A.Vessieres, P. Pigeon, M. Huche, E. A. Hillard, G. Jaouen, J.
Organomet. Chem., 692 (2007) 1219.
[17] C. Biot, G. Glorian, L. A. Maciejewski, J. S. Brocard, O. Domarle, G. Blampain, P. Millet,
A. J. Georges, H. Abessolo, D. Dive, J.Lebibi, J. Med. Chem., 40 (1997) 3715.
[18] (a) M. F. R. Fouda, M. M. Abd-Elzaher, R.A. Abdelsamaia, A. A. Labib, Appl. Organomet.
Chem., 21 (2007) 613.
(b) C. S. Allardyce, A. Dorcier, C. Scolaro, P. Dyson, J. Appl. Organomet. Chem., 19 (2005)
1.
[19] M. Miwa, H. Tamura, Chem. Lett., (1997) 1177.