Cytotoxicity activity, in silico molecular docking, protein- and DNA-binding study of a new Ni(II) Schiff base complex

Article abstract of DOI:10.1080/00958972.2018.1492118

One new nickel(II) complex, [Ni(L)] (1), was synthesized from the Schiff base ligand derived from pyrrole-2-carboxaldehyde and 1,3-diaminopropane. Complex 1 was characterized by elemental analysis, IR, UV-Vis and ESI mass spectroscopy, cyclic voltammetry, and single-crystal X-ray structure analysis. Crystallographic results show that two Ni(II) monomeric moieties are present with similar structural features but with slightly different bond lengths and bond angles. The geometry around the Ni(II) center is distorted square planar. DNA-binding properties of complex 1 were well explored by employing UV-Vis and fluorescence spectral methods, cyclic voltammetry, and by viscosity measurements. Similarly the protein-binding study was studied by multispectroscopic techniques using both BSA and HSA. The cytotoxicity study of the compound has also been evaluated. Notably, the in vitro cytotoxicity of complex 1 on two cancer cell lines (AGS and A549) demonstrates that complex 1 has very good anticancer activity. MTT assay, cell-cycle analysis, and annexin-V assay have been performed to know the extent of effect of complex 1 as anticancer agent. Further, in silico molecular docking study revealed that the nickel(II) complex fits into the minor groove of duplex DNA by hydrophobic interaction with functional groups of B-DNA.

Full text of DOI:10.1080/00958972.2018.1492118

Journal of Coordination Chemistry
ISSN: 0095-8972 (Print) 1029-0389 (Online) Journal homepage: http://www.tandfonline.com/loi/gcoo20
Cytotoxicity activity, in silico molecular docking,
protein- and DNA-binding study of a new Ni(II)
Schiff base complex
Niladri Biswas, Sumit Khanra, Arnab Sarkar, Shamee Bhattacharjee, Deba
Prasad Mandal, Ankur Chaudhuri, Sibani Chakraborty & Chirantan Roy
Choudhury
To cite this article: Niladri Biswas, Sumit Khanra, Arnab Sarkar, Shamee Bhattacharjee, Deba
Prasad Mandal, Ankur Chaudhuri, Sibani Chakraborty & Chirantan Roy Choudhury (2018):
Cytotoxicity activity, inꢀsilico molecular docking, protein- and DNA-binding study of a new Ni(II)
Schiff base complex, Journal of Coordination Chemistry
To link to this article: https://doi.org/10.1080/00958972.2018.1492118
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Cytotoxicity activity, in silico molecular docking, protein- and DNA-binding
study of a new Ni(II) Schiff base complex
NILADRI BISWASa, SUMIT KHANRAb, ARNAB SARKARc, SHAMEE BHATTACHARJEEc,
DEBA PRASAD MANDALc, ANKUR CHAUDHURId, SIBANI CHAKRABORTYd and
CHIRANTAN ROY CHOUDHURY*a
aDepartment of Chemistry, West Bengal State University, Barasat, Kolkata-700126, India
bDepartment of Chemistry, Indian Institute of Science Education and Research, Kolkata Mohanpur - 741 246,
West Bengal, India
cDepartment of Zoology, West Bengal State University, Barasat, Kolkata-700126, India
dDepartment of Microbiology, West Bengal State University, Barasat, Kolkata-700126, India
One new nickel(II) complex, [Ni(L)] (1), was synthesized from the Schiff base ligand derived
from pyrrolle-2-carboxaldehyde and 1,3-diamino-propane. Complex 1 was characterized by
elemental analysis, IR, UV-vis and ESI mass spectroscopy, cyclic voltammetry and single-
crystal X-ray structure analysis. Crystallographic results show that two Ni(II) monomeric
moieties are present with similar structural features but with slightly different bond lengths and
bond angles. The geometry around the Ni(II) center is distorted square planar. DNA-binding
properties of complex 1 were well explored by employing UV-vis and fluorescence spectral
methods, cyclic voltammetry and by viscosity measurements. Similarly the protein-binding study
was studied by multispectroscopic techniques using both BSA and HSA. The cytotoxicity study
of the compound has also been evaluated. Notably, the in vitro cytotoxicity of complex 1 on two
cancer cell lines (AGS and A549) demonstrates that complex 1 has very good anticancer activity.
MTT assay, cell-cycle analysis and annexin-V assay have been performed to know the extent of
effect of complex 1 as anticancer agent. Further, in silico molecular docking study revealed that
the nickel(II) complex fits into the minor groove of duplex DNA by hydrophobic interaction with
functional groups of B-DNA.
Keywords: Nickel(II) complex; X-ray structure; DNA Binding; Protein binding; Cytotoxicity;
In silico molecular docking
*Corresponding author. Email: crchoudhury2000@yahoo.com

1. Introduction
Nature has already used different transition elements for various biological processes. Based on
these ideas, researchers aim at the development of functional models [1-7]. Both the metal ions
and ligand parts largely influence the stereochemistry as well as properties of complexes [8].
Nowadays, systematic investigation of anticancer property of metal complexes is one of the main
areas of research [8, 9]. But cis-platin, which has application in cytotoxic study, is found to have
numerous side-effects which ultimately creates the need to develop more effective as well as less
toxic anticancer drugs [10, 11]. Recent study focused on the interaction of metal complexes with
DNA and understanding the mechanism of action of the probe [12]. Different drugs mostly target
DNA and can cause damage of DNA in the target cell [10, 13-15]. In recent time, much research
is focused on the development of efficient, target specific, non-toxic drugs [16, 17]. Till now,
DNA-targeted nickel complexes are very rare in literature compared to other transition metal
complexes like copper, platinum, etc. [12, 18-21]. Very few mono- and di-nuclear Ni(II)
complexes are available in literature which show very good biological activity [22, 23]. From
literature, it is observed that suitable anticancer activity is shown by compounds when aromatic
ligands contain N4 or N2O2 coordination [24-30]. Due to the similarity with HSA, low-cost BSA
is mostly used as target protein for interaction study [31-34]. Increased interaction of metal
complexes with protein molecules will directly dictate the effective concentration of drugs
in vivo [10]. So model study of binding of metal complexes in vitro can be considered very
effective in determining the binding nature of proteins [10, 35].
This fact promoted us to work with Ni(II) metal for the synthesis of metal complexes.
The present work reports the synthesis of a Schiff base ligand derived from pyrrole-2-
carboxaldehyde and 1,3-diaminopropane (scheme 1) and its nickel(II) complex [Ni(L)] (1). The
entitled nickel(II)-Schiff base complex is well characterized by IR and UV-vis spectroscopy,
cyclic voltammetry and single-crystal X-ray structure analysis. Interaction of complex 1 with
CT-DNA was examined by using UV-vis and fluorescence spectral methods, viscosity
measurement and cyclic voltammetric study. The protein (BSA and HSA) binding studies were
determined with the help of UV-vis as well as fluorescence spectral methods. Theoretical
molecular docking studies were also carried out to ascertain the interaction pattern of nickel(II)
complex with B-DNA. The cytotoxicity profiles of the titled nickel(II) complex along with cell
line study have also been investigated. The significant groove binding of complex 1 with DNA

was explained on the basis of its low intrinsic binding constant value and this groove binding
pattern was further explained with the help of viscosity measurements and theoretical molecular
docking studies.
Scheme 1. Formation of ligand H2L.
2. Experimental
2.1. Materials
All the required chemicals were of reagent grade and used without purification. Nickel(II) nitrate
was purchased from E-Merck and used as received. Pyrrole-2-carboxaldehyde, HSA, BSA and
TBAP were purchased from Sigma-Aldrich. Tris-HCl-buffer (pH 7.2) and CT-DNA were
purchased from SRL. 1,3-Diaminopropane and ethidium bromide (EB) were purchased from
Spectrochem.
2.2. Physical measurements
FT-IR spectra were recorded from 400-4000 cm-1 for complex 1 in solid KBr pellets by using a
Perkin-Elmer SPECTRUM - 2 FT-IR spectrophotometer. The UV-vis spectra of the complex

were recorded using a Perkin-Elmer Lambda-35 UV-Vis spectrophotometer in Tris-HCl buffer
medium at 300 K. The fluorescence measurements were carried out with a Perkin-Elmer LS 55
fluorescence spectrophotometer in Tris-HCl-buffer medium. ESI mass spectra were recorded in
an Agilent 6520 Q-Tof Spectrometer at room temperature in the m/z range of 0-1000. Elemental
analyses were measured with a Perkin-Elmer 2400 II elemental analyzer. During measurements
of emission spectra, an excitation and emission slit width of 10 nm was used. All the
electrochemical experiments were carried out with three electrode configuration using a CH
660E cyclic voltammeter in Tris-HCl buffer medium. Saturated calomel electrode (SCE) as
reference, Pt wire-electrode as counter electrode and glassy carbon electrode as a working
electrode were used as three electrode system with tetrabutylammonium perchlorate (TBAP) as a
supporting electrolyte at a scan rate of 50 mV sec-1. All the electrochemical data were recorded
under a dry nitrogen environment. Nitrogen gas was passed into the sample solution at a constant
rate for 1 min. The viscosity measurements were carried out using a semi-micro viscometer at
25 °C.
The doubly-distilled water was used to prepare Tris-HCl-buffer maintaining the pH
around 7.2. The concentration of CT-DNA in base pairs was calculated with the help of molar
extinction coefficient of 6600 M-1cm-1 at 260 nm. The ratio of absorbance at 260 and 280 nm
was greater than 1.8 which is the indication of high purity of CT-DNA [24].
2.3. Synthesis and characterization of ligand and complex
The tetradentate Schiff base ligand (H2L) and complex 1 were prepared according to the
following methods. Chemical structure of 1 is shown in figure 1.
2.4. Synthesis of tetradentate Schiff base ligand (H2L)
Schiff base (H2L) was prepared by mixing equimolar concentration of pyrrole-2-carxaldehyde
(0.4755 g, 5 mmol) and 1,3-diaminopropane (0.418 mL, 5 mmol) in 50 mL methanolic solution.
It was refluxed for 45 min until the color of the solution changed from colorless to straw yellow.
The resultant solution was cooled at room temperature during complex preparation.

2.5. Synthesis of [C13H14N4Ni] (1)
A methanolic solution of Schiff base ligand (1 mmol) was added to a methanolic solution of
Ni(NO3)2∙6H2O (0.2908 g, 1 mmol, 10 mL). After mixing, the solution became light red. Then
the resultant solution was stirred over a magnetic stirrer for 10 min until the color turned into
deep red and it was left for crystallization. After seven days red-colored single-crystals were
appeared which were collected by filtration. Yield: 64% (0.254 g). Anal. Calc. for [C13H14N4Ni]:
C, 57.74; H, 4.91; N, 19.65%. Found: C, 57.71; H, 4.86; N, 19.60%. IR (KBr, cm-1): 3467(b),
1595(s), 1275-1436(w) and 489(m).
2.6. Single-crystal X-ray diffraction study
Good quality single-crystal of 1 was mounted on a Bruker APEX-II CCD diffractometer,
equipped with graphite monochromated Mo Kα radiation (λ = 0.71073 Å) fine-focus sealed
tubes. Intensity data were collected at 292(2) using \ω/2\q scans. Crystal data were collected
using an APEX2 (Bruker, 2008). Data refinement and reduction were performed using SAINT
(Bruker, 2008) software [36]. Multiscan absorption corrections were applied empirically to the
intensity values (Tmin = 0.939 and Tmax = 0.992) using SADABS [36]. The structures were solved
by direct methods using the program SIR97 [37] for 1 and refined with full-matrix least-squares
based on F2 using SHELXL-97 [38]. All non-hydrogen atoms were refined anisotropically.
C-bound hydrogen atoms were placed geometrically and refined using a riding model
approximation. The molecular graphics and crystallographic illustration for 1 was prepared using
DIAMOND [39] and ORTEP [40].
2.7. DNA-binding experiment
2.7.1. Electronic absorption spectrophotometric study. The interaction of complex 1 with
CT-DNA was measured in Tris-HCl-buffer solution having pH 7.2. The bulk solution of
CT-DNA was prepared by adding CT-DNA in Tris-HCl-buffer solution followed by dilution at
4 °C and it was kept for at least four days. The CT-DNA concentration was measured by its
absorption intensity with the help of molar extinction coefficient (6600 M-1 cm-1) at 260 nm. A
ratio of absorption intensity at 260 and 280 nm of about 1.80:1 signifies that the CT-DNA was
sufficiently free from protein molecule and also indicates its purity [41a-b]. The intrinsic binding
constant (Kb) and binding mode of complex 1 with CT-DNA was determined by the

spectrophotometric titration of fixed metal complex solution (50 µM) with the continuous
variation of CT-DNA solution (0-90 µM).
2.7.2. Competitive binding fluorescence measurement. The interaction between complex 1
and CT-DNA with EB was further investigated by fluorescence study. The solution of EB was
prepared by dissolving it into Tris-HCl buffer solution (pH 7.2). Generally the emission intensity
of classical intercalator EB increases due to complex formation with CT-DNA [42, 43]. The
solution of complex 1 in Tris-HCl buffer was added into the aliquot of CT-DNA pretreated with
EB. Significant changes of emission intensities at 607 nm (excitation wavelength 508 nm) were
observed. The competitive binding experiment results with the displacement of EB from
prepared DNA-EB mixture which results in significant quenching of fluorescence intensity.
2.7.3. Electrochemical study. To determine the redox behavior of complex 1, cyclic
voltammetric measurements were performed. This experiment was carried out in a single-
compartmental cell at room temperature in Tris-HCl buffer medium using 0.1 M TBAP as
supporting electrolyte. A three electrode system containing calomel, Pt-wire and Pt-working as
reference, counter and working electrode, respectively, were used for this measurement.
Electrochemical experiments in the absence and presence of CT-DNA were carried out to
confirm the binding mode of complex 1 with CT-DNA under dry nitrogen atmosphere.
2.7.4. Viscosity study. Viscosity measurement was carried out using a semi-micro viscometer
suspended vertically in a thermostatic water bath at 25 °C. This experiment was done by keeping
CT-DNA concentration fixed (10 µM) and varying metal complex concentration (10-60 µM).
The average flow times were recorded for three times for each sample with the help of a digital
stop watch. The obtained data was plotted as (η/η0)1/3 versus [complex]/[DN], where η and η0 are
the viscosity of DNA in the presence and absence of complex 1, respectively, in the Tris-HCl
buffer solution [12].
2.8. Protein-binding study
The binding nature of complex 1 with bovine serum albumin (BSA) and human serum albumin
(HSA) were determined by analyzing absorbance as well as fluorescence spectral studies.

Fluorescence spectra were recorded in the range of 280-460 nm (excitation wavelength 279 nm)
for both BSA and HSA. The protein stock solutions were prepared by dissolving BSA and HSA
into 50 mM Tris-HCl buffer and kept under dark conditions at 4 °C. The protein binding studies
were performed by maintaining the fixed concentration of protein solution (40 µM) in Tris-HCl
buffer having pH 7.2 and varying the concentration of the Ni(II) complex from 0-60 µM. UV-vis
spectral titrations were carried out with concentration of proteins (40 µM) and Ni(II) complex
stock solutions (0-70 µM) at room temperature.
2.9. Molecular modeling and docking studies
In silico binding mode analysis between synthesized nickel complex and duplex DNA was
examined. The crystal structure of duplex DNA d(CGCGAATTCGCG)2 dodecamer (PDB ID:
1BNA) was retrieved from Protein Data Bank (PDB). All the water molecules were removed
from the crystal structure. Hydrogen atoms and Momany-Rone partial charges were added to the
duplex DNA. Duplex DNA and complex 1 were typed with CHARMM force-field [44, 45]
before molecular docking study. Docking calculations were performed using CDOCKER
program [46]. Molecular docking poses of complex 1 were ranked according to CDOCKER
interaction energy. Finally energy minimization of the docked receptor-complex 1 was
performed for further analysis. All procedures were carried out using Discovery Studio 3.5
software [47].
2.10. Cytotoxicity study
Annexin V-assay Kit was purchased from eBioscience. Anti-mouse anti-bodies against PCNA
and GAPDH were procured from Santa Cruz (USA), bacitracin, leupeptin, pepstatin A, PMSF,
phosphatase inhibitor cocktails. A-Sepharose beads, RNase, NAC and NBT were purchased from
Sigma (St. Louis, MO). Nitrocellulose membrane and filter papers were obtained from Pall
Corporation, USA.
Human lung (A549) and gastric (AGS) cancer cell lines were obtained from NCCS Pune.
Cells were cultured at 37 °C in humidified atmosphere of 5% CO2 -95% air. Cell culture medium
contains DMEM and F12K (Gibco, invitrogen, USA) supplemented with 10% heat inactivated
fetal bovine serum (FBS; Invitrogen.) and 1% penicillin G and streptomycin (Life Technologies,

Rockville, MD, USA). The cells, initially seeded at the concentration of 104 cells/mm2, were
incubated at 37 °C with humidity saturated controlled atmosphere and 5% CO2.
2.10.1. Hemolytic assay. Fresh human blood was centrifuged at 4000Xg for 10 min and the cell
pellet was washed thrice and re-suspended in 10 mM PBS at pH 7.4 to obtain a final
concentration of 1.6×109 erythrocytes/mL. Equal volumes of erythrocytes were incubated with
varying concentrations of capsaicin and eugenol and with shaking at 37 ºC for 1 h. Samples were
then subjected to centrifugation at 3500Xg for 10 min at 4 °C. RBC lysis was measured at
different peptide concentrations by taking absorbance at an OD of 540 nm. Complete hemolysis
(100%) was determined using 1% Triton X 100 as a control. Hemolytic activity of the spice
active components was calculated in percentage using the following equation:
H=100 × (Op-Ob)/(Om-Ob)
where Op is the optical density of given peptide concentration, Ob is the optical density of buffer
and Om is the optical density of Triton X 100.
2.10.2. Cell viability assay on PBMC. The Trypan Blue dye exclusion test is used to determine
the number of viable cells present in a cell suspension. It is based on the principle that live cells
possess intact cell membranes that exclude trypan blue, whereas dead cells do not. The effect of
complex 1 on the viability of PBMC cells was determined by Trypan blue exclusion test. Cells
were treated with varying of these molecules and at definite time point (24 h) the cells that could
exclude the Trypan blue dye were counted in haemocytometer as viable cells.
2.10.3. MTT assay. This colorimetric assay is based on the activity of mitochondrial succinate
dehydrogenase enzyme in live cells to reduce the yellow water soluble substrate MTT into an
insoluble, colored formazan product which is measured spectrophotometrically at 595 nm. Since
reduction of MTT can only occur in metabolically active cells, the level of activity is a measure
of the viability of cells. The cytotoxicity of these synthetic molecules was tested on AGS and
A549 cells by MTT-assay. Briefly, 2×104 cells were seeded in a 96-well microtitre plate and
incubated with different concentrations of these synthetic molecules. After 12 h of exposure to
these molecules, 10 µL of MTT (5 mg/mL) was added to each well, and the cells were incubated

in the dark at 37 °C for an additional 4 h. Thereafter, the medium was removed, the formazan
crystals were dissolved in 100 µL of dissolving solution (10% SDS in isopropanol), and the
absorbance measured at 595 nm.
2.10.4. Cell cycle distribution analysis. Cell cycle analysis is a method in that employs flow
cytometry to distinguish cells in different phases of the cell cycle based on fluorescence intensity
of the stained cells at certain wavelength. For the determination of cell cycle phase distribution
of nuclear DNA, in vitro AGS, A549 cells (1×106 cells) were harvested. After making a single
cell suspension, cells were fixed with 3% p-formaldehyde, permeabilized with 0.5% Triton
X-100, and nuclear DNA was labeled with propidium iodide (PI, 125 μg/mL) after RNase
(40 µg/mL) treatment. Cell cycle phase distribution of nuclear DNA determined on FACS
Caliber using Cell Quest Software (Becton-Dickinson Histogram display of DNA content
(x-axis, PI fluorescence) versus counts (y-axis) has been displayed.
2.10.5. Annexin V assay. Apoptosis assays were carried out based on the instruction from the
Annexin V Apoptosis Kit (eBioscience). Briefly, PI and Annexin V were added directly to AGS
and A549 cells treated with complex 1 (0.5 µg/mL). The mixture was incubated for 15 min at
37 °C. Cells were fixed and then analyzed on FACS VERSE (equipped with 488 nm Argon laser
light source; 515 nm band pass filter, FL1-H, and 623 nm band pass filter, FL2-H) (Becton
Dickinson). Electronic compensation of the instrument was done to exclude overlapping of the
emission spectra. Total 10,000 events were acquired, the cells were properly gated and dual
parameter dot plot of FL1-H (x-axis; Fluos-fluorescence) versus FL2-H (y-axis; PI-fluorescence)
shown in logarithmic fluorescence intensity.
2.10.6. Western blot analysis. Western blot (also called protein immunoblotting, because an
antibody is used to specifically detect its antigen) is a widely accepted analytical technique used
to detect specific proteins in the given sample. By analyzing location and intensity of the band
formed, expression details of the target proteins in the given cells could be obtained. AGS and
A549 cell lysates were obtained and equal amounts of protein from each sample were diluted
with loading buffer, denatured, and separated by 10% sodium dodecyl sulfate-polyacrylamide
gel electrophoresis (SDS-PAGE) followed by protein transfer to nitrocellulose membrane. The

effect of complex 1 on PCNA expression was checked on AGS and A549 cancer cells. The blot
was incubated with anti-PCNA antibody, followed by blotting with HRP-conjugated secondary
antibody. The blots were then detected by using a chemiluminescent kit from Thermofisher. This
analysis was performed two times.
2.10.7. Statistical analysis. The experiments were repeated three times and the data were
analyzed statistically. Values have been represented as mean ± SD. Data were analyzed and
Student’s t-test was used to evaluate the statistical differences. Statistical significance was
considered when P < 0.05.
3. Results and discussion
3.1. IR spectra
IR spectrum of complex 1 is shown in figure S1. Complex 1 exhibits a strong band at 1595 cm-1
which can be assigned to the C=N stretching frequency of azomethine group. The band observed
at 489 cm-1 can be attributed to M-N stretching frequency, indicating the participation of
azomethine nitrogen in the coordination with the metal ion. Moreover a broad band at 3467 cm-1
appeared which is the characteristic stretching of N-H bond [48]. Pyrrole ring in-plane and out-
of-plane vibrations are observed in the region 1275-1436 cm-1.
3.2. Electronic absorption spectra
Absorption spectrum of the complex was recorded in Tris-HCl medium in the range 200-800 nm
at 300 K and the spectrum is shown in figure S2. A very low intensity band appeared at 521 nm
which corresponds to the d→d transition of Ni(II) having square planar stereochemistry [12]. A
high-intensity band at 314 nm and a low-intensity band at 272 nm were observed due to
intraligand →* transitions of the coordinated imines of the ligand. Complex 1 shows two low-
energy bands at 433 nm and at 389 nm with a shoulder which can be attributed to n→*
transition [49].
3.3. Cyclic voltammetry
The electrochemical property of mononuclear Ni(II) complex was investigated by cyclic
voltammetry at a scan rate of 50 mV sec-1 having potential range between +2.0 to -2.0 V in Tris-

HCl buffer solution containing TBAP as supporting electrolyte. Figure S3 shows two oxidative
peaks at potentials +0.77 to +1.30 V in the anodic region and one reductive peak at -0.86 V in
the cathodic region. The oxidation peak at +0.77 V (versus SCE) is due to the oxidation of Ni(II)
to Ni(III) and the other oxidative wave at +1.30 V can be attributed for the Schiff base oxidation.
Complex 1 also showed one irreversible reduction peak at -0.86 (versus SCE) which is believed
to be due to conversion of Ni(II) to Ni(I) [10].
3.4. X-ray crystal structure description
An ORTEP diagram of complex 1 with atom numbering scheme is given in figure 1 and packing
diagram is shown in figure S4. Crystal data and details of the structure determination for
complex 1 are shown in table 1. The asymmetric unit consists of two molecular units, Ni1 and
Ni2, which are crystallographically independent but chemically similar, with slightly different
bond lengths and bond angles. Each of the two mononuclear units contains Ni(II) as the central
atom having square planar geometry. Each Ni(II) atom (Ni1 and Ni2) is coordinated to two
N-atoms coming from 2-pyrolle moiety (N1 and N4 for Ni1; N6 and N8 for Ni2) and another
two nitrogen atoms from the imine part of the Schiff base (N2 and N3 for Ni1; N5 and N7 for
Ni2). Ni-N bond lengths are in the range 1.8854(19)-1.9119(19) Å, which is similar to the
compounds found in the literature [50-53].
The distortion in the Ni coordination environment is clearly shown by the deviations in
the cisoid and transoid N-Ni-N bond angles from their ideal values (table 2). These variations of
bond lengths and bond angles clearly indicate the Ni centers (Ni1 and Ni2) are slightly distorted
from ideal square planar geometry [52-54].
3.5. DNA-binding experiments
3.5.1. Electronic absorption titration. Before addition of CT-DNA to complex 1, its behavior
in the Tris-buffer solution (pH ~7.2) at room temperature was observed by UV-vis spectroscopy
for 24 h. No dissociation of the ligand (both H2L) was observed under these conditions. The
UV-vis spectra of complex 1 in DMSO, Tris-buffer medium and acetonitrile are also compared
in figure S5 (Supplementary Information). No significant differences were observed in the three
spectra showed (refer to table T1 of Supplementary Information). All these data indicate that

complex 1 was stable under the conditions examined (for a detailed discussion about the integrity
of complex 1 in aqueous medium, see figure S5).
The most important evidence of the integrity of complex 1 was obtained by ESI mass
spectra in positive mode in aqueous medium. The molecular ion peak appeared at m/z 286,
which could be attributed to the molecular ion [Ni(L)] peak with ~100% abundance (figure 2).
This proved that complex 1 did not undergo any kind of dissociation in aqueous medium under
ordinary conditions.
Electronic spectral study is a kind of reliable measurement which is commonly used for
investigation about the binding mode of metal complexes with DNA. Due to the cellular receptor
property of DNA, the metal complex-DNA interaction is one of the versatile areas of modern
research. Generally, metal complexes interact with DNA resulting in either hyperchromism or
hypochromism [55a-b]. Usually, during intercalative binding mode, a significant stacking
interaction occurs between DNA base pairs and planar aromatic ring of the investigating
complex. On the other hand, a prominent increase in absorption of the metal complex with the
addition of CT-DNA solution is an effective area of non-intercalative binding. In the case of non-
intercalative binding, a reasonable damage of CT-DNA double helix is observed. On the other
hand, during groove binding, the metal complex interacts with the external surface of DNA
either by van der Waals forces or hydrogen bonding [56a-b]. In the present study, complex 1
displayed two absorption bands in the range of 272-314 nm due to →* transition and a band at
389 nm with a shoulder due to n→* transition (figure 3). To determine the binding nature of
complex 1 with CT-DNA, fixed concentration of metal complex solution (50 µM) was titrated
with the incremental increase of CT-DNA solution (0-90 µM). Before obtaining the spectral
data, the mixed solution of metal complex and CT-DNA in Tris-HCl-buffer was allowed to
equilibrate for 10 min for each titration. After successive addition of CT-DNA to the fixed
concentration of metal complex, the →* transition energy increases along with
hyperchromism with slow blue-shift and also a steady decrease in n→* transition energy. The
observed hyperchromism with blue-shift is an indication of hydrogen bonding or van der Waals
interaction [57]. So, it is a kind of interaction that has occurred between the external surface of
CT-DNA molecule and complex 1 [57-59]. The spectral data of complex 1 in the absence or
presence of CT-DNA are shown in figure 3. The binding affinities of the nickel(II) complex with

CT-DNA can be determined by measuring intrinsic binding constant (Kb) using equation (1)
[60],
[DNA] / [a-f] = [DNA] / [b-f]+1/ Kb[b-f]
(1)
where [DNA] is the CT-DNA concentration in base pairs, a is the apparent extinction coefficient
corresponding to Aobsd/[Ni], b is extinction coefficient of the complex in fully bound form and
f is extinction coefficient of the complex in unbound form. For complex 1, when spectral data
are fitted in equation (1), we obtained a straight line with a positive slope 1/([b-f]) and an
intercept of 1/Kb [b-f], which are shown in figure 3 (inset). The intrinsic binding constant (Kb)
for complex 1 can be determined from the ratio of the slope of the Y-intercept [57, 61, 62]. The
observed Kb was 7.582(±0.016)×103 M-1, which is very much lower than that of a classical
intercalator such as EB having Kb value 1.4×105 M-1 [63a]. The observed intrinsic binding
constant (Kb) of nickel(II) complex is similar to previously reported related Schiff base metal
complexes, which are in the range of 103 M-1 [24] to 104 M-1 [63b-h]. This Kb value confirms that
complex 1 moderately bound to CT-DNA by non-intercalative binding mode.
3.5.2. Competitive binding experiments. To further investigate the binding mode of complex 1
with CT-DNA, fluorescence spectral titrations were performed. No emission spectrum was found
for complex 1 itself or in the presence CT-DNA solution at room temperature. Therefore to
determine the binding nature of complex 1, a competitive binding study has been performed
using EB. EB is a well-known planar organic cationic dye whose fluorescence intensity is low
but its emission intensity can be enhanced by intercalating into the base pairs of CT-DNA
[64, 65]. Therefore, EB is identified as a typical standard of intercalation [66, 67]. Fluorescence
binding experiments were carried out for complex 1 (0-50 µM) in the presence of CT-DNA
bound EB in Tris-HCl buffer medium. At 607 nm (figure 4) prominent decrease in the emission
intensity of EB-DNA [68, 69] has been observed due to the displacement of EB from CT-DNA–
EB mixture. This can be explained due to the fact that the number of binding sites on the
CT-DNA was reduced for EB. The EB-displacement study is explained on the basis of Stern-
Volmer equation [16],
I0/I = 1+Ksv [Q]
(2)

where I0 and I are the emission intensities in the absence and presence of the quencher,
respectively, Ksv is the linear Stern-Volmer quenching constant, and [Q] is the quencher
concentration. From the plot of I0/I versus [Q] is shown in figure 4 (inset). The calculated Ksv
value is 3.534(±0.078)×104 M-1. The quenching rate constant (Kq) can be determined with the
help of the equation Ksv = Kq×τ0, where τ0 is the lifetime of fluorophore in the absence of
quencher molecule (CT-DNA). Since the fluorescence lifetime is in the order of 10-8 s and using
this value, quenching rate constant (Kq) of complex 1 is 3.534(±0.078)×1012 M-1s-1. The observed
Kq value of complex 1 is higher than the limiting diffusion rate constant for a biomacromolecule
(2.0×1010 M-1s-1) [24]. The above observation illustrates that there is static quenching
phenomenon between the fluorophore and the quencher molecule and they form complex in the
ground state [24, 70]. The apparent binding constant (Kapp) can be estimated using equation (3),
KEB [EB] = Kapp [complex]
(3)
where KEB = 1.0×105 M-1 which is DNA binding constant of EB, [EB] is the concentration of EB
and [complex] is the concentration of the quencher used to determine 50% decrease in the
emission intensity of CT-DNA pretreated with EB [71, 72]. The apparent binding constant of
complex 1 is 2.86×104 M-1. Therefore, analyzing the difference in the Kapp value of complex 1 to
the KEB value of EB it is clear that complex 1 is not a good classical intercalator like EB [57].
The number of binding sites (n) were also determined using Scatchard equation [1, 73],
log [(I0-I)/I] = logKb + n log [Q]
(4)
where n is the number of binding sites. A straight line was obtained by plotting log [(I0-I)/I]
versus log [Q] and from the slope of the plot (figure S6, table 3); the number of binding sites was
obtained as 1.09079±0.1488 (table 3).
3.5.3. Electrochemical study. Cyclic voltammetry can be regarded as a very sensitive analytical
method for the determination about the change of the redox property of metal complex under
investigation [24, 74, 75]. Complex 1 displayed two oxidative responses at 0.77 and 1.30 V
(vs SCE) and one reductive response at -0.86 V (vs SCE) at a scan rate of 50 mV sec-1 in
Tris-HCl buffer medium containing 0.1 M TBAP as supporting electrolyte. After incremental

addition of CT-DNA solution, the oxidative response at 0.77 V was shifted to 0.63 V (more
negative potential value) and the reductive response was also shifted to -0.79 V (more positive
potential value). Further addition of CT-DNA solution results in the disappearance of the
oxidative response at 0.77 V (figure S7). The observed changes in both anodic and cathodic
responses clearly indicate that complex 1 binds with CT-DNA [24, 76]. So, finally it can be
assumed that there is a kind of interaction between complex 1 and CT-DNA through which the
electron cloud of complex 1 could be “pulled” towards the CT-DNA strands [77].
3.5.4. Viscosity properties. Viscosity measurements are a very useful technique to investigate
the binding modes between intercalating molecules and DNA. Generally, in classical
intercalative binding modes, prominent increase in DNA viscosity is generally observed due to
the increase of the length of DNA double helix and also because of accommodation of molecules
into the base pairs [78]. On the other hand, in the case of groove binding, DNA viscosity is little
influenced by small molecules [79] due to bending or kinking of DNA double helix, which
results in reduced effective length of DNA [68, 80]. The effect of complex 1 on CT-DNA
viscosity is shown in figure S8. In this figure, it is clear that CT-DNA viscosity in Tris-HCl
buffer medium does not show any significant changes with the successive addition of increasing
concentration of complex 1. From this, it is confirmed that the Ni(II) complex does not
intercalate into the base pair of CT-DNA double helix. So, the kind of interaction that occurs
between complex 1 and the external surface of CT-DNA is either by van der Waals forces or by
hydrophobic interaction.
3.6. Protein binding studies
3.6.1. Electronic absorption titration. Electronic absorption measurement is a useful method to
determine the structural changes and mechanism of complex formation in donor-acceptor
systems. To investigate the protein-metal complex molecule relationship, BSA as well as HSA
are widely used for this purpose due to their similar structural environment [42, 81-83]. Serum
albumin plays an important role in the transport of drugs and metal ions throughout the blood
circulatory system. Generally based on the mechanism, quenching process is classified in two
categories, one is static and another is dynamic quenching. In the case of static quenching,
fluorophore-quencher complex formation occurs in ground state and for dynamic quenching,

quencher molecule and fluorophore come into contact during the short-lived excited state [84].
To confirm whether the titled nickel(II) complex acts as either static or dynamic quencher,
UV-vis absorption measurements were carried out. The changes in absorption spectra of both
BSA and HSA after successive addition of complex 1 are shown in figures S9A and S9B,
respectively. From figure S9, it is clear that the absorption bands of both BSA and HSA
increased after the addition of complex 1. There is a significant change of the absorption band of
both the fluorophore molecules BSA and HSA which indicates static quenching process, whereas
in dynamic quenching, the absorption spectra of fluorophore molecule is not changed. So, there
is a ground-state complex formation between the nickel complex and BSA as well as HSA
[81, 84].
3.6.2. Fluorescence quenching study. Fluorescence quenching measurements are widely
studied for further confirmation regarding the quenching pattern of metal complexes with protein
molecules like BSA and HSA [12, 85-87]. Basically protein fluorescence is responsible for three
residues of protein, i.e. tryptophan, tyrosine and phenylalanine [8]. Binding with these protein
molecules can result in either loss or enhancement of the biological properties of the drug
molecule. Fluorescence quenching is a phenomenon which reduces the fluorescence intensity of
a fluorophore molecule because of the variety of processes involved [88]. In our current study,
both BSA and HSA were selected as target protein molecules. In figures 5 and S10, there is a
prominent decrease in emission intensity of BSA and HSA at 345 and 341 nm, respectively, after
incremental addition of complex 1. This is due to formation of complex-protein system and
alteration of the local environment around tryptophan residue. A significant red-shift was
observed for both proteins which indicates a conformational change of tertiary structure of
protein molecules. The above quenching effect was analyzed with the help of the Stern-Volmer
equation [22, 70],
F0/F = 1+Ksv [Q]
(5)
where F0 and F are the fluorescence intensity in the absence and presence of quencher molecule,
respectively. Ksv denotes linear Stern-Volmer quenching constant and [Q] is the quencher
concentration. A plot of F0/F versus [Q] for both BSA and HSA results in a linear plot (insets in
figures 5 and S10). The value of Ksv is 4.83(±0.034)×104 and 3.59(±0.066)×104 M-1 for BSA and

HSA, respectively. The quenching rate constant (Kq) of complex 1 can be obtained using the
equation Ksv = Kqτ0, where τ0 is the average lifetime of protein molecule in the absence of
quencher. The τ0 value for tryptophan fluorescence in proteins is 10-8 s [55, 89]. Complex 1
shows higher quenching rate constant values (for both BSA and HSA) than that of biopolymers
(2.0×1010 M-1 s-1) [89]. The quenching rate constant values of nickel(II) complex with both BSA
and HSA are given in table 4. The quenching rate constant (Kq) of complex 1 determined by
protein quenching procedure is 100 times higher than scatter procedure. The above observation is
an indication of static quenching mechanism [90, 91]. Moreover for dynamic quenching, the
quenching rate constant (Kq) value is normally in the range of 1010 M-1s-1 [1]. This behavior is
the indication of static interaction between BSA or HSA with complex 1. When the nickel(II)
complex binds to the active site of the protein molecules, the binding constant (Kb) as well as
number of binding sites (n) can be determined using the Scatchard equation [84],
log [(F0-F)/F] = logKb + n log[Q]
(6)
By plotting of log [(F0-F)/F] versus log[Q] for both proteins (figure S11), the number of binding
sites (n) and the binding constant (Kb) values were determined from the slope and intercept of the
plot, respectively. The quenching constant (Kq), binding constant (Kb) and number of binding
site (n) values are shown in table 4. By comparing all the experimental data of both BSA and
HSA, it is clear that Kb, Kq and n values of BSA are slightly higher than those of HSA. So,
finally it can be concluded that complex 1 binds more strongly to BSA than HSA and so more
strongly quenches the emission intensity of BSA than HSA.
3.7. Molecular docking of complex 1 with DNA
In order to gain better insight into the nature of DNA-complex binding, molecular docking study
of nickel complex with DNA duplex of sequence d(CGCGAATTCGCG)2 was performed.
Docked complex conformation was examined in terms of interaction energy and hydrophobic
interaction between synthesized nickel complex and duplex DNA. In silico molecular docking
study revealed that the nickel complex fits into the minor groove of duplex DNA by hydrophobic
interaction with functional groups of B-DNA (figure 6). From the interaction energy analysis, it

is clear that van der Waals interactions play a predominant role relative to electrostatic
interactions in the binding process (table 5).
3.8. Cytotoxic profile study
3.8.1. Cytotoxicity of complex 1 in normal cells. Percentage haemolytic activity of complex 1
on human red blood cells is shown in figure S12. Data from hemolytic assay reveals that
complex 1 does not show significant haemolysis up to a concentration of 10 µg/mL. Percentage
of haemolysis increases quite significantly beyond 10 µg/mL. However, the cytotoxic activity of
the compound was found to increase considerably in nucleated cells as determined by Trypan
blue exclusion method in human PBMC. Percentage PBMC viability measured by Trypan Blue
exclusion method of complex 1 is shown in figure 7. Results suggest that there was significant
loss in cell viability beyond a concentration of 0.5 µg/mL.
Haemolytic assay has been done to test the haemolytic effect (lysis of RBC) of the
particular compound on human erythrocytes [92] whereas cytotoxicity test on PBMC indicates
compound toxicity towards normal cell [93].
Several reports suggest that many drugs have toxic effects on red-blood cell membrane
and causes haemolytic anaemia [92, 94]. Therefore, haemolytic assay is needed for a particular
drug before use. In our experiment, we have found that compound 1 shows its lytic effect
significantly from 10-15 µg/mL.
Peripheral blood mononuclear cell (PBMC) is considered as a normal cell for studying
cytotoxic effects of a drug. Several reports indicate that a killing dose of anticancer drug
sometimes shows more toxicity to normal cells than cancer cells. So it is necessary to select
particular drug concentration that would not kill normal cell but has cytotoxic effects (killing not
lysis as in haemolysis) on cancer cell. Our experimental result shows that at 0.5 µg/mL (1/20 of
the haemolytic dose), compound 1 shows more killing effect on cancer cells (A549, AGS) than
normal cell (PBMC).
3.8.2. Determination of IC50 in cancer cell lines by MTT assay. A549 and AGS cell viability
measured by MTT assay at various concentration of complex 1 is shown in figure 8. MTT assay
revealed the cytotoxicity of complex 1 in A549 and AGS cells in a concentration-dependent

manner. More than 50% killing was observed at and above a concentration of 0.5 µg/mL in both
cell lines.
As mentioned above, 0.5 µg/mL is comparatively less toxic to normal cells and is able to
induce more than 50% killing in cancer cells. For this reason we have selected 0.5 µg/mL
concentration for further studies on cancer cells.
Coordination of metal ions with ligands is the key-step for cytotoxicity of novel metal
complexes [31]. IC50 values of complex 1 against AGS and A549 cell lines are 15.09 and
35.09 µM, respectively. The cytotoxic activity of complex 1 is compared with other similar
complexes and shown in table 6 [92-104]. In the cytotoxity process, coordination of metal ions
and the nature of the ring substituents play an important role [31]. The cytotoxic activity of
complex 1 is somewhat superior to that of similar complexes reported in the literature (table 6)
[92-104]. A valid explanation is the difference in activity against the selected tumor cell lines
which can well be explained in terms of variable ligand occupancy with the central metal ion
[60]. When ligand:metal is 1:1, then it is a less planar tetrahedral geometry but when the
coordination is 2:1, then it must have more planar square planar geometry [60]. In this case,
monomeric complex 1 showed higher activity than non-planar complexes (dimer or polymer)
(table 6), but are nearly comparable in terms of activity with similar Ni-complexes having planar
geometry. Complex 1 has good potential as a cytotoxic agent due to the presence of a pyrolle
ring of the coordinated ligand [60]. As not many references available for cytotoxic study with
AGS cell line, in this case Ni- and Cu-complexes are cited. Although complex 1 is active against
AGS and A549 cell lines under in vitro cytotoxicity experiment, the complex is nearly
comparable as the effectiveness shown by the standard drug cis-platin (table 6).
3.8.3. Analysis of cell cycle phase distribution of cancer cells. Figure 9 shows cell-cycle
analysis of treated and untreated AGS and A549 cells by flow cytometer using PI as DNA-
binding fluorochrome. Treatment with complex 1 resulted in a significant increase in the sub-G0
region in both AGS and A549 cells (hypopoidy population). As compared to 4.03±0.058 % cells
in the hypoploidy region of the AGS control group, complex 1 at 0.5 µg/mL caused a significant
increase in the hypopoidy peak which is 10.57±1.08 %. In the case of A549 cells, hypoploidy
population was 13.7±1.1 % in complex 1 treated cells as compared to 5.3±0.43 % in the control

group. Bar diagram representation of cell cycle phase distribution of AGS with wild type and
p53 knock out cells is shown in figure S13.
3.8.4. Effect of complex 1 in AGS and A549 apoptosis. To confirm the increase in apoptotic
induction the complex 1, we performed annexin V/PI assay in all the groups by staining the AGS
and A549 cells with FITC tagged annexin V and PI and measuring the fluorescence intensity in a
flow cytometer. The results of annexin V assay in AGS and A549 cells are shown in figure 10.
The results suggest a significant increase in the percentage of annexin positive cells from
5.14±0.04 % in the AGS control group to 11.37±0.59 % in complex 1 treated groups. Percentage
of annexin positive cells increases from 3.88±0.04 % in the A549 control group to 12.83±0.89 %
in complex 1 groups.
3.8.5. Western Blot analysis of protein expressions. Bar diagram representation of percent
Annexin V positive AGS cells from different experimental groups are also given in figure S14.
Expression of cell proliferation associated marker, proliferating cell nuclear antigen (PCNA),
was found to reduce significantly upon complex 1 treatment in both AGS and A549 cells.
4. Conclusion
One nickel(II) complex with N4-chelation [Ni(L)] has been synthesized and characterized by
structural, analytical as well as spectroscopic methods. DNA-binding activity of complex 1 was
determined using UV-vis absorption, competitive fluorescence titration, cyclic voltammetry and
viscosity measurements. The low value (in the order 103 M-1) of the binding constant suggests
groove mode of binding. This fact was further established by in silico molecular docking study.
Biological profile of complex 1 as anticancer agent is also studied. Complex 1 has been found to
exhibit in vitro antiproliferative activity towards two cell lines, AGS and A549 cancer cells.
MTT assay, cell-cycle analysis and annexin-V assay have been performed to know the extent of
effect of complex 1 as an anticancer agent.
All the results demonstrate that complex 1 has very good anticancer activity.

Supplementary data
CCDC 1008381 contains the supplementary crystallographic data for complex 1. These data can
be obtained free of charge via http://www.ccdc.cam.ac.uk/conts/retrieving.html, or from the
Cambridge Crystallographic Data Centre, 12 Union Road, Cambridge CB2 1EZ, UK; Fax: (+44)
1223-336-033; or E-mail: deposit@ccdc.cam.ac.uk. Supplementary data associated with this
article can be found in the online version.
Acknowledgement
C. Roy Choudhury acknowledges DST-FIST (Project No. SR/FST/CSI-246/2012) New Delhi,
Govt. of India for instrumental support under capital heads.
Disclosure statement
No potential conflict of interest was reported by the authors.
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Graphical abstract

Figure 1. ORTEP view of complex 1 with displacement ellipsoids drawn at 50% probability
level (hydrogens are omitted for clarity).