Syntheses, characterizations and biophysical studies of Cu(II) diphenylphosphate complexes: Effect of co-ligands on their biological properties

Article abstract of DOI:10.1016/j.poly.2012.08.087

Using the same bridging ligand, diphenyl phosphate (dpp), and different co-ligands, 1,10-phenanthroline (phen)/2,2′-bipyridine (bpy), two structurally similar Cu(II) complexes, [Cu(μ-dpp)(phen)(NO₃)] ₂ (1) and [Cu(μ-dpp)(bpy)(NO₃)]₂ (2), were synthesized. These complexes were characterized by single crystal X-ray diffraction as well as other physico-chemical methods. Both complexes are dinuclear in nature, but complex 1 is extended into a 3D supramolecular architecture by π-π interactions, whereas complex 2 forms a 2D supramolecular layered structure by π-π and C-H...π interactions. We further explored the DNA binding ability of these complexes by an agarose gel electrophoresis study. Although the coordination structures of these two complexes are very similar, complex 1 was found to be more effective at DNA binding. Complex 1 showed a greater cytotoxic effect on the human hepatocellular carcinoma cell line HepG2 compared to complex 2. Moreover, complex 1 induced more S-phase arrest and apoptosis in HepG2 cells than complex 2, as determined by fluorescence activated cell sorters (FACs). Thus our results indicated that changing the co-ligands in copper complexes may rendered an overall change in the supramolecular structure as well as significant variations in the biological activities.

Full text of DOI:10.1016/j.poly.2012.08.087

Polyhedron 48 (2012) 157–166
Contents lists available at SciVerse ScienceDirect
Polyhedron
journal homepage: www.elsevier.com/locate/poly
Syntheses, characterizations and biophysical studies of Cu(II) diphenylphosphate
complexes: Effect of co-ligands on their biological properties
a,
Rajdip Dey a,1, Debalina Bhattacharya b,1, Parimal Karmakar b, , Debajyoti Ghoshal
⇑
⇑
a Department of Chemistry, Jadavpur University, Jadavpur, Kolkata 700 032, India
b Department of Life Science and Bio-technology, Jadavpur University, Jadavpur, Kolkata 700 032, India
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 2 July 2012
Accepted 28 August 2012
Available online 21 September 2012
Using the same bridging ligand, diphenyl phosphate (dpp), and different co-ligands, 1,10-phenanthroline
(phen)/2,20-bipyridine (bpy), two structurally similar Cu(II) complexes, [Cu(
-dpp)(phen)(NO3)]2 (1) and
[Cu( -dpp)(bpy)(NO3)]2 (2), were synthesized. These complexes were characterized by single crystal X-
ray diffraction as well as other physico-chemical methods. Both complexes are dinuclear in nature, but
complex 1 is extended into a 3D supramolecular architecture by interactions, whereas complex 2
forms a 2D supramolecular layered structure by and C–H. . . interactions. We further explored
the DNA binding ability of these complexes by an agarose gel electrophoresis study. Although the coor-
dination structures of these two complexes are very similar, complex 1 was found to be more effective at
DNA binding. Complex 1 showed a greater cytotoxic effect on the human hepatocellular carcinoma cell
line HepG2 compared to complex 2. Moreover, complex 1 induced more S-phase arrest and apoptosis
in HepG2 cells than complex 2, as determined by fluorescence activated cell sorters (FACs). Thus our
results indicated that changing the co-ligands in copper complexes may rendered an overall change in
the supramolecular structure as well as significant variations in the biological activities.
Ó 2012 Elsevier Ltd. All rights reserved.
l
l
p–p
p
Keywords:
p
–
p
Copper(II) complex
X-ray structure
Biophysical study
DNA binding
Bridging ligand
1. Introduction
cis-platin, one of the most popular metal-based anticancer
drugs, shows its activity by covalent linkage to DNA [12–17], but
Molecules able to irreversibly modify nucleic acids have re-
ceived considerable attention due to their potential applications
in molecular biology as well as in drug design [1–4]. Thus, several
intriguing efforts have been observed in the last two decades to
study the DNA binding ability of metal complexes to facilitate
the development of metal-based anticancer drugs. Metal com-
plexes can be associated with DNA by three distinct modes, namely
external binding, groove binding and intercalation. The coordi-
nated metal complexes can bind to DNA by either covalent link-
ages or being intercalated within the base pairs by means of
non-covalent supramolecular forces, and both may be used to de-
stroy malignant cells. Such metallo-intercalators formed by a cova-
lent attachment of organic intercalators to transition metal
coordination complexes can interact with DNA, and consequently
modulate biological activity. Metal complexes having aromatic
side arms can bind to DNA by metal coordination as well as
through intercalation of the attached aromatic ligand [5–11].
at the same time some severe side effects of this drug have been
reported [18–21]. On the other hand, the energy involvements in
supramolecular interactions are quite low [22–29], and thus sev-
eral other interactive weak forces can also be negotiated during
the modification of the structures of anticancer agents. As a result,
the design of non-covalently DNA binding anticancer drugs has be-
come important due to their lesser toxicity and target specific nat-
ure in comparison to drugs which creates covalent linkages to DNA
[30–32]. Thus, continuous efforts are being made to design such
non-covalent DNA intercalators using transition metals, particu-
larly Cu(II), for their possible use as potential anticancer drugs
[10,33–36].
Different complexes of Cu(II) have been extensively studied for
their DNA binding ability as well as DNA cleavage activity
[3,31,32,37]. Due to the redox potential value of Cu(II), it can cleave
DNA both in an oxidative way [38,39] as well as in a hydrolytic way
[34–36]. In addition, some Cu(II) complexes can cleave DNA upon
photo-activation, which may be potentially important for their
use in photodynamic therapy (PDT) [40–44]. The Cu(II) center,
along with some aromatic N, N0 donor chelating ligands, e.g.,
1,10-phenanthroline and 2,20-bipyridine, have proven efficient in
this regard, due to their planar character [3,31,37]. Thus the
⇑
Corresponding authors.
E-mail addresses: pkarmakar_28@yahoo.co.in (P. Karmakar), dghoshal@chemistry.
jdvu.ac.in (D. Ghoshal).
1
Authors have made an equal contribution.
0277-5387/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved.
http://dx.doi.org/10.1016/j.poly.2012.08.087

158
R. Dey et al. / Polyhedron 48 (2012) 157–166
composition and specific structure of such complexes are impor-
tant for therapeutic purposes.
In this work, we have synthesized two complexes of Cu(II) using
the 1,10-phenanthroline and 2,20-bipyridine ligands. Along with
these, we have used a phospho-diester, diphenyl phosphate, con-
sidering the well-known biological importance of these com-
1,10-phenanthroline. Blue colored single crystals suitable for X-
ray diffraction analysis were obtained after 20 days (yield 83%).
Anal. Calc. for C44H36N6 O14P2 Cu2: C, 49.76; H, 3.41; N, 7.91. Found:
C, 49.74; H, 3.39; N, 7.90%.
2.4. Crystallographic data collection and refinement
pounds [45]. Phosphate esters are present in
a variety of
naturally occurring substances and man-made derivatives which
can be employed for several biological applications [45]. Moreover,
phosphodiesters form the backbone of DNA and RNA molecules,
and that is why the recognition of these types of molecules are per-
haps important when associated with Cu(II) species [46].
Suitable single crystals of 1 and 2 were mounted on a thin glass
fiber with commercially available super glue. X-ray single crystal
data collection of the two crystals were performed at room
temperature (25 °C) using a Bruker APEX II diffractometer,
equipped with a normal focus, sealed tube X-ray source with
Here we wish to report the synthesis, characterization, single
crystal X-ray structure and biological activity of two compounds,
[Cu(l-dpp)(phen)(NO3)]2 (1) and [Cu(l-dpp)(bpy)(NO3)]2 (2). These
complexes have been synthesized with the same bridging ligand,
diphenyl phosphate (dpp), and two different co-ligands, 1,10-
phenanthroline (phen) and 2,20-bipyridine (bpy), In both cases the
compounds are dinuclear, but complex 1 is extended into a 3D
graphite monochromated Mo-Ka radiation (k = 0.71073 Å). The
data were integrated using the SAINT [47] program and the absorp-
tion corrections were made with SADABS. All the structures were
solved by SHELXS-97 [48] using the Patterson method and followed
by successive Fourier and difference Fourier synthesis. Full matrix
least-squares refinements were performed on F2 using SHELXL-97
[49] with anisotropic displacement parameters for all non-hydro-
gen atoms. All calculations were carried out using SHELXL-97,
SHELXS-97, PLATON v1.15 [50], ORTEP-3v2 [51] and WinGX system
Ver-1.80 [52].
supramolecular architecture by
plex 2 forms a 2D supramolecular sheet structure by
H. . . interactions. In order to explore the biological activity of
p–p
interactions, whereas com-
p–p
and C–
p
these complexes, their DNA binding abilities have been investi-
gated using different biophysical methods and also their cytotoxic
effects have been evaluated. Complex 1 is more effective than com-
plex 2 in binding with DNA and induces a nick in the super coiled
DNA. Additionally, the cytotoxic activity of complex 1 is more than
that of complex 2 on human hepatocellular carcinoma cell lines
HepG2 and the cytotoxicity is mediated by S-phase arrest and sub-
sequent induction of apoptosis.
2.5. Preparations of stock solutions for biophysical and biological
studies
Both the copper complexes were dissolved in DMSO at a con-
centration of 10 mM. The stability of the complexes in DMSO
was checked by comparing the electronic spectra of the complexes
in the solid state as well as in DMSO (Supplementary Fig. 1). Calf
thymus (CT) DNA and pUC19 plasmid DNA were dissolved sepa-
rately at a concentration of 3 mg/ml stock in 10 mM citrate phos-
phate buffer (pH 7.4) at room temperature. Powdered EtBr and
Hoechst 33258 were dissolved in double distilled water at concen-
trations of 10 mM and 1 mM, respectively. Stocks were stored at
4 °C in the dark and diluted freshly by double distilled water before
the experiments.
2. Experimental
2.1. Materials
High purity Cu(II) nitrate trihydrate, 1,10-phenanthroline
(phen), 2,20-bipyridine (bpy), diphenylphosphate (dpp), propidium
iodide and calf thymus DNA were purchased from Aldrich Chemi-
cal Company Inc. Ethidium bromide (EtBr) and pUC19 plasmid
DNA were purchased from Bangalore Genei, India. RNAse A was
obtained from SRL, India and Hoechst 33258 was purchased from
Polysciences, USA. All other chemicals were of AR grade. All the
chemicals are used as received without further purification.
2.6. Absorption spectroscopic study of plasmid DNA with the
complexes
Absorption spectroscopic studies were done on a HITACHI
U-2800 spectrophotometer, using pUC19 plasmid DNA (35
mL) with increasing concentrations of the complexes (10–
70 M). After each addition, the DNA and complex mixtures were
lg/
l
2.2. Physical measurements
incubated at room temperature for 15 min and scanned from 210
to 305 nm. Each sample was scanned for a cycle number of 2, cycle
time of 5 s at a scan speed of 100 nm/min. The modified Benesi–
Hildebrand equation [53] was used for the determination of the
ground state binding constant between pUC19 DNA and the com-
plex. The binding constant ‘‘K’’ was determined using the following
relation:
Elemental analysis (carbon, hydrogen and nitrogen) were per-
formed using a Perkin–Elmer 240C elemental analyzer. Infrared
spectra (4000–400 cmꢀ1) of powder samples were taken in KBr
pellets, using a Perkin–Elmer Spectrum BX-II IR spectrometer.
2.3. Syntheses of the complexes
A0=D
A ¼ A0=
D
Amax þ ðA0=
D
AmaxÞ ꢁ 1=K ꢁ 1=Lt
An aqueous solution of copper(II) nitrate trihydrate (1 mmol,
0.241 g) was mixed with a methanolic solution (10 ml) of diphenyl
phosphate (1 mmol, 0.250 g) with constant stirring for 15 min.
After that, a methanolic solution (10 ml) of 1,10-phenanthroline
(1 mmol, 0.198 g) was added to the above mixture. After stirring
the mixture for 3 h, it was filtered and the filtrate was kept in a
CaCl2-desiccator. Deep blue single crystals suitable for X-ray dif-
fraction analysis were obtained after 14 days (yield 78%). Anal.
Calc. for C48H36N6 O14P2 Cu2: C, 51.94; H, 3.26; N, 7.57. Found: C,
51.92; H, 3.25; N, 7.54%.
where
D
A = A0 ꢀ A,
D
Amax = maximum reduced absorbance, A0 -
= maximum absorbance of DNA (without any ligand), A = reduced
absorbance of DNA (in the presence of ligand) and Lt = ligand
concentration.
2.7. Fluorescence spectroscopic studies of complexes with CT DNA
Fluorescence spectroscopic studies of CT DNA (35 lg/ml) with
varying concentrations of the complexes were done by using a HIT-
ACHI F3010 spectrofluorimeter. In this experiment, EtBr solution
was gradually added to the said concentration of CT DNA and at
each time the fluorescence pattern was scanned. The fluorescence
Complex 2 was synthesized using the same procedure as
that for 1 using 2,20-bipyridine (1 mmol, 0.156 g) instead of

R. Dey et al. / Polyhedron 48 (2012) 157–166
159
intensity was saturated at 100
saturation level, the respective complexes were added gradually
(up to 90 M) until the fluorescence pattern decreased up to the
l
M concentration of EtBr. At this
were incubated at room temperature for 15 min in the dark. Final-
ly, the fluorescence of cells was immediately determined by a Bec-
ton Dickinson (FACS Calibur) flow cytometer [55].
l
starting value again. The excitation wavelength was 546 nm and
the emission spectra were scanned from 550 to 640 nm, spectral
response of 2 s, along with a scanning speed of 60 nm/min. The
same experiment was done using Hoechst 33258 instead of EtBr.
For Hoechst 33258, the excitation wavelength was 351 nm and
emission spectra were scanned from 400 to 550 nm. Saturation
of fluorescence intensity was observed at a concentration of
3. Results and discussion
3.1. IR spectroscopy
Both the complexes shows very similar bands in their IR spec-
trum due to their similar structures. Compounds 1 and 2 shows a
medium intensity band at 1208 and 1200 cmꢀ1 respectively, which
5.3
lM of Hoechst 33258 and then the complexes were added up
to 170
lM.
may be assigned to the stretching frequency (m) of the P@O bond
[56]. The mas and ms for the P–OR bond in 1 are observed at 1018
and 918 cmꢀ1, whereas those for 2 are at 1024 and 925 cmꢀ1
respectively. The three medium intensity bands present at 1466,
1361 and 1097 cmꢀ1 (for complex 1) and 1472, 1384 and
2.8. Study of the binding activity of the complexes by agarose gel
electrophoresis
pUC19 plasmid DNA (500 ng) was incubated separately with
different concentrations of both complexes and the pure phen li-
gand (0.5–5 mM) separately at 37 °C for 1 h, and then the samples
were run in 2% agarose gel in 1ꢁ Tris–acetate–EDTA buffer (pH 7.4)
at 2 V/cm for 12 h.
1096 cmꢀ1 (for complex 2) correspond to
mas(NO2), ms(NO2) and
m(N@O) respectively [56]. Both the complexes display medium
intensity bands in the range 3100–3000 cmꢀ1 which corresponds
to the stretching vibration of aromatic C–H groups. The next group
of bands appears for both the complexes at around 1400–
1600 cmꢀ1, corresponding to the stretching vibration of aromatic
C@C groups [56].
2.9. Cell culture and MTT assay
Human liver carcinoma cells (HepG2) were cultured in Dul-
becco’s modified eagle medium (DMEM) supplemented with 10%
fetal bovine serum (FBS) and 100 U/ml penicillin–streptomycin at
5% CO2 and 37 °C. Viability of the HepG2 cells after exposure to
various concentrations of the complexes and phen ligand were
assayed by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium
bromide (MTT). Briefly, cells were seeded in 96 well plates at
1 ꢁ 104 cells/well and these exponentially growing cells were ex-
posed to the complexes at concentrations of 0, 0.1, 0.2, 0.5, 1, 2.5,
3.2. Crystal structures
3.2.1. Single crystal X-ray structure of complex 1
ꢀ
Complex 1 crystallizes in the triclinic centrosymmetric P1 space
group. Data collection, structure refinement parameters and crys-
tallographic data for complex 1 are given in Table 1. The complex
is dinuclear in nature, where the phenyl phosphate acts as a bridg-
ing ligand (Fig. 1). Here each Cu(II) center exhibits a distorted octa-
hedral geometry with an elongated Cu–O bond [Cu1–O7
2.6255(18) Å], which is characteristic for a Jahn Teller distorted
Cu(II) ion. The Cu(II) center is chelated by two nitrogen atoms of
the 1,10-phenanthroline (N1,N2), with Cu–N distances of
2.028(15) and 2.014(18) Å (Table 2). The other four coordination
sites of the metal atom are occupied by two oxygen atoms of the
nitrate ligand (O5, O7), having Cu–O distances of 1.995(16) and
5 and 10
washed with phosphate buffer saline (PBS) twice and incubated
with MTT solution (450 g/ml) for 3–4 h at 37 °C. The resulting
lM for 24 and 48 h. After incubation, the cells were
l
formazan crystals were dissolved in an MTT solubilization buffer,
the absorbance was measured at 570 nm by using a Biorad micro-
plate reader and the values were compared to the mock-treated
cells.
2.10. Cell cycle analysis
Table 1
About 1 ꢁ 106 cells of HepG2 were seeded in a 90 mm tissue
Crystallographic and structural refinement parameters for complexes 1 and 2.
culture plate for 24 h. The exponentially growing cells were treated
1
2
separately with 1 lM complex 1 and 4 lM complex 2. After 24 h,
Formula
C
48H36N6O14P2Cu2
C44H36N6O14P2Cu2
1061.83
monoclinic
P21/c
the cell pellets were washed with 1ꢁ PBS and fixed with chilled
Formula weight
Crystal system
Space group
Unit cell dimensions
a (Å)
1109.87
triclinic
P1
80% ethanol and kept at 4 °C for overnight. Prior to staining with
ꢀ
50
lg/ml propidium iodide, the cells were incubated for 1 h with
100
l
g/ml of DNAse free RNAse A at 37 °C. The cell cycle was ana-
9.7573(4)
11.0256(4)
12.0534(4)
88.628(2)
68.447(1)
75.615(1)
1164.96(8)
1
1.582
1.058
566
1.8–27.6
19732
11.182(5
18.720(5)
11.163(5)
90
101.425(5)
90
2290.4(16)
2
1.540
lyzed with a Becton Dickinson fluorescence activated cell sorter
(FACS) Calibur flow cytometer and 25000 events were counted at
each data point [54].
b (Å)
c (Å)
a
(°)
b (°)
c
(°)
2.11. Apoptosis by annexin V-FITC
V (Å3)
Z
Dcalc (g cm–3
)
Exponentially growing HepG2 cells were checked for apoptosis
by addition of either complex 1 or complex 2 at the dose men-
tioned in the previous section, and untreated HepG2 was kept as
a control. Both control and test samples were incubated for 24 h.
Following the incubation, the cells were washed twice with PBS.
After that, the cells at a concentration of about 1 ꢁ 106 cells/ml
were resuspended in 1ꢁ annexin binding buffer. Five hundred
microliters of each cell suspension was added to a FACS tube
l
(mm–1
)
1.072
1084
F (000)
hrange (°)
Reflections collected
Unique reflections
1.9–27.7
33127
5339
4022 (0.075)
1.06
5353
4954 (0.083)
1.09
0.0379
0.1073
Reflections (Rint
)
Goodness-of-fit (GOF) (F2)
R1 [I > 2
wR2
r
(I)]a
0.0541
0.1669
a
(12 ꢁ 75 mm), and 2
lg/ml of annexin V-FITC and 0.5 lg/ml of pro-
pidium iodide were added to each cell suspension. Then the tubes
P
P
P
P
||Fo| ꢀ |Fc||/ |Fo|, wR2 = [ (w(F2o ꢀ Fc2)2)/ w(Fo2)2]
.
a
½
R1
=

160
R. Dey et al. / Polyhedron 48 (2012) 157–166
Fig. 1. Atom labeling diagram of complex 1.
phen and the phenyl rings of dpp are involved in p–p interactions
Table 2
exclusively within the rings of phen and dpp respectively. This
may be attributed to the self-recognition phenomena portrayed
by two different aromatic rings present in a complex. One of the
Selected bond lengths (Å) and bond angles (°) for complex 1.
Cu1–O1
Cu1–O7
Cu1–N2
O1–Cu1–O5
O1–Cu1–N1
O1–Cu1–O2a
O5–Cu1–N1
O2a–Cu1–O5
O7–Cu1–N2
1.9342(14)
2.6255(18)
2.0149(18)
95.00(6)
169.62(7)
97.29(5)
92.15(6)
95.55(6)
113.67(6)
Cu1–O5
1.9959(16)
2.0280(15)
2.2103(12)
88.14(6)
90.07(7)
53.60(6)
165.96(6)
90.05(6)
149.11(6)
Cu1–N1
aromatic rings of phen, which is not involved in intra-sheet
p–p
Cu1–O2a
interactions, creates inter-sheet interactions to get an overall
p–p
O1–Cu1–O7
O1–Cu1–N2
O5–Cu1–O7
O5–Cu1–N2
O7–Cu1–N1
O2a–Cu1–O7
3D supramolecular structure (Supplementary Fig. 2 and Table 3).
3.2.2. Single crystal X-ray structure of complex 2
Complex 2 crystallizes in the monoclinic system with the P21/c
space group. Data collection, structure refinement parameters and
crystallographic data for complex 2 are given in Table 1. Like 1 this
complex is also dinuclear in nature, where the phenyl phosphate
acts as a bridging ligand (Fig. 3). Here also each Cu(II) center
exhibits a distorted octahedral geometry with a elongated Cu–O
bond [Cu1–O 2.731(4) Å] (Table 4), which is characteristic of a Jahn
Teller distorted Cu(II) ion. Each Cu(II) is chelated by two nitrogen
atoms [N1,N2; Cu–N distances 2.007(2) and 2.000(2) Å
respectively] of the 2,20-bipyridine, which along with two oxygen
atoms, one from the nitrate group (O5) and the other from the
diphenyl phosphate [O1a; Cu–O distance 1.927(2) Å], create the
equatorial plane of the octahedron. The axial sites are occupied
by the oxygen atoms (O2) of dpp and the O atom (O6) of the nitrate
ligand with bond distances of 2.154 (2) and 2.731(4) Å respectively
(Table 4).
Symmetry code: a = 1 ꢀ x, 2 ꢀ y, 1 – z.
2.6255(18) Å, and two O atoms from the phenyl phosphate (O1 and
O2) with Cu–O distances of 1.934(14) and 2.210(12) Å (Table 2).
The two nitrogen atoms (N1,N2) of the 1,10-phenanthroline and
two oxygen atoms, one from the nitrate group (O5) and the other
from the phenyl phosphate (O1), creates the equatorial plane.
The axial sites are occupied by another oxygen atom (O2a) of sym-
metry generated diphenyl phosphate (dpp) and the O atom (O7)
from the nitrate.
In the crystal packing, two adjacent dimeric units are locked by
face-to-face
p–p interactions created between the aromatic rings
of one phen moiety, which produces a one dimensional supramo-
lecular chain along the b axis. In that 1D pattern, the floppy phenyl
The crystal packing of complex 2 shows a strong face to face
rings of dpp are further locked by face-to-face
create a supramolecular sheet structure in the bc plane (Fig. 2 and
Table 3). Here it is interesting to note that the aromatic rings of
p
–
p
interactions to
p
–
p
interaction between the aromatic rings of bpy along the bc
plane and the phenyl rings of the dpp are engaged in a C–H. . .
interaction with the H atom of another phenyl ring of dpp. These
p

R. Dey et al. / Polyhedron 48 (2012) 157–166
161
Fig. 2. Projection along a-axis of 1 showing the interlocking of the dimeric units.
Table 3
and C–H. . .p interactions in 1.
p
–p
Ring(i) ? ring(j)
Distance of
centroid(i) from
ring(j) (Å)
Dihedral Distance between the
angle(i,j) (i,j) ring centroids (Å)
(°)
R(1) ? R(1)i
R(1) ? R(2)i
R(2) ? R(2)ii
R(3) ? R(1)i
R(4) ? R(4)iii
C–H ? ring(j)
3.7657(12)
3.7227(11)
3.5282(11)
3.7226(11)
4.1829(16)
0
3.5316(8)
3.5184(8)
3.2752(8)
3.5413(8)
3.4629(11)
C. . .R distance (Å)
1.52(9)
0
1.52(9)
0
H. . .R distance (Å) C–H. . .R
angle (°)
128
C(14)–H(14) ? R(1)iv 2.98
3.626(2)
Symmetry code: (i) = 2 ꢀ x, 1 ꢀ y, 1 ꢀ z; (ii) = 2 ꢀ x, 2 ꢀ y, 1 ꢀ z; (iii) = ꢀx, 2 ꢀ y,
2 ꢀ z; (iv) = 1 ꢀ x, 2 ꢀ y, 1 ꢀ z. R(i)/R(j) denotes the ith/jth rings: R(1) = N(1)/C(10)/
C(9)/C(8)/C(7)/ C(11); R(2) = N(2)/C(1)/C(2)/C(3)/C(4)/ C(12); R(3) = C(4)/C(5)/C(6)/
C(7)/C(11)/C(12); R(4) = C(13)/C(14)/C(15)/C(16)/C(17)/C(18).
interactions create a 2D single layer structure (Fig. 4 and Table 5).
The disposition of the 2D sheets shows an almost parallel arrange-
ment along the b-axis (Supplementary Fig. 3)
3.3. Bio-physical study
Fig. 3. Atom labeling diagram of complex 2.
The in vitro binding of these two complexes with pUC 19 DNA was
studied by absorption spectrophotometry. The absorption maxima
at 258 nm of pUC 19 DNA was decreased gradually with increasing
constant of complexes 1 and 2 with DNA are calculated as
3.36 ꢁ 105 and 2.7 ꢁ 105 Mꢀ1 respectively from the Benesi–Hilde-
concentrations of the complexes (1.5–7.5 lM). The association

162
R. Dey et al. / Polyhedron 48 (2012) 157–166
Table 4
Table 5
Selected bond lengths (Å) and bond angles (°) for complex 2.
p–p and C–H. . .p interactions in 2.
Cu1–O2
Cu1–O6
Cu1–N2
2.154(2)
2.731(4)
2.000(3)
93.19(9)
106.96(9)
93.87(9)
159.01(9)
93.33(9)
91.73(12)
80.74(10)
171.58(10)
Cu1–O5
2.007(2)
2.007(2)
1.927(2)
143.85(10)
90.64(10)
50.65(10)
93.51(11)
109.03(10)
88.83(10)
91.14(9)
Ring(i) ? ring(j)
Distance of
centroid(i) from
ring(j) (Å)
Dihedral Distance between the
angle(i,j) (i,j) ring centroids (Å)
(°)
Cu1–N1
Cu1–O1a
O2–Cu1–O5
O2–Cu1–N1
O1a–Cu1–O2
O5–Cu1–N1
O1a–Cu1–O5
O6–Cu1–N2
N1–Cu1–N2
O1a–Cu1–N2
O2–Cu1–O6
O2–Cu1–N2
O5–Cu1–O6
O5–Cu1–N2
O6–Cu1–N1
O1a–Cu1–O6
O1a–Cu1–N1
R(1) ? R(2)i
R(2) ? R(1)ii
C–H ? ring(j)
3.631(3)
3.631(3)
H. . .R distance (Å) C–H. . .R
2.3(2)
2.3(2)
3.6105(19)
3.5953(16)
C. . .R distance (Å)
angle (°)
C(15)–H(15) ? R(3)i 2.91
149
3.739(4)
Symmetry code: (i) = x, 1/2 ꢀ y, 1/2 + z; (ii) = x, 1/2 ꢀ y, ꢀ1/2 + z. R(i)/R(j) denotes the
ith/jth rings: R(1) = N(1)/C(1)/C(2)/C(3)/C(4)/C(10); R(2) = N(2)/C(8)/C(7)/C(6)/C(5)/
C(9); R(3) = C(17)/C(18)/C(19)/C(20)/C(21)/C(22).
Symmetry code: a = 1 ꢀ x, ꢀy, 2 – z.
brand equation [53] (Supplementary Figs. 4 and 5). To study the abil-
ity of these compounds to interfere with the complex formed by
DNA intercalated with the dye EtBr, each of the complexes was
added gradually to EtBr saturated DNA and the fluorescence spectra
were taken after each addition. The fluorescence intensity of pUC 19
DNA (OD 258 = 0.7) and EtBr was measured by adding increasing
concentrations of EtBr (Supplementary Fig. 6). The saturation in
fluorescence intensity was observed at an EtBr concentration of
with increasing concentrations of complex 2, it was seen that the
fluorescence intensity slowly decreased and returned to the basal le-
vel after the addition of 80 lM solution of complex 2 (Supplemen-
tary Figs. 6c and 6d). Further, we explored the ability of the said
agents to interfere with the complex formed by DNA and the DNA
groove binder Hoechst 33258, a well-known DNA minor groove bin-
der. The fluorescence intensity of a solution containing 30 lg/ml of
pUC 19 plasmid DNA in 10 mM Tris–Cl [tris(hydroxymethyl)amino
methane and conc. HCl] of pH 7.4 was gradually increased with
the addition of increasing concentrations of Hoechst 33258. The
fluorescence intensity was saturated at a concentration of 1.7 lM
of Hoechst 33258 (Supplementary Fig. 8). The fluorescence intensity
100 lM, whichshowed maximum fluorescence intensity at an emis-
sion wavelength of 580 nm. Complex 1 was then added gradually to
the EtBr saturated DNA and the fluorescence spectra were taken in
each time. With the addition of complex 1, the fluorescence intensity
of EtBr saturated DNA was gradually decreased and finally returned
of the Hoechst 33258–DNA complex did not return to the basal value
to the basal level at a concentration of 20 lM of complex 1 (Supple-
after gradual addition of complex 1 up to 40 lM, indicating partial
mentary Figs. 6a and 6b). It is important to note that neither of the
complexes have any fluorescence property, either in the free form
or in the DNA bounded form. Moreover, the ligand phen did not
change the fluorescence intensity of EtBr saturated DNA (Supple-
mentary Fig. 7). Thus, the quenching of the fluorescence intensity
of EtBr saturated DNA was probably due to the binding of the
complexes with DNA. When EtBr saturated DNA was challenged
displacement of the DNA groove binding dye from DNA (Supple-
mentary Figs. 8a and 8b). It is also worth mentioning that the phen
ligand does not change the fluorescence intensity of the Hoechst
33258–DNA complex at all (Supplementary Fig. 9). The same exper-
iment has performed with complex 2 (Supplementary Figs. 8c and
8d), where the fluorescence intensity of the Hoechst 33258–DNA
complex was slightly quenched. Thus the spectrofluorimetric data
Fig. 4. 2D sheet formation in 2 along the a-axis by p–p and C–H. . .p interactions.

R. Dey et al. / Polyhedron 48 (2012) 157–166
163
Fig. 5. Agarose gel (2%) electrophoresis study of pUC19 plasmid DNA (500 ng) with increasing concentrations of (a): ligand, (b) complex 1 and (c): complex 2. Lane 8 is marker
DNA for all figures. Relative intensity of RF1 and RF2 for respective concentrations of (a) ligand, (b) complex 1 and (c) complex 2 is plotted at the bottom.
suggest that both the complexes can interact with DNA by intercala-
tion as well possibly having a weak partial interaction with the DNA
minor groove.
seen in Fig. 6, complex 2 induces almost no effect on HepG2 cells
up to 5 M, but complex 1 shows significant cell killing ability
(Fig. 6a). In order to rationalize the cell killing ability of complex
1, we performed the same assay with the phen ligand only. The
respective ligand induced only a moderate effect on the cells. It
l
3.4. Gel electrophoresis
is interesting to note that for complex 1, 2.5
all the cells. The calculated LD50 values are 1
and 2.5 M for the free phen ligand.
We also tested the cytotoxic effect of the complexes, when the
cells were incubated for 48 h in the presences of the complexes.
l
M is sufficient to kill
The effect of the complexes was also studied by DNA migration
in agarose gel. pUC 19 plasmid DNA was incubated for 60 min at
37 °C in the presence of increasing concentrations of the respective
complexes or ligand (phen only), and the samples were run in 2%
agarose gel in 1ꢁ TAE (a mixture of Tris base, acetic acid and EDTA)
buffer of pH 7.4 at 2 V/cm for 12 h. The gel was stained with EtBr
and photographs were taken in a gel documentation system. As
seen in Fig. 5, the presence of phen did not change the DNA migra-
tion or intensity of the super coiled (RF1) DNA. But in the presence
of phen containing complex 1, the intensity of super coiled DNA de-
creased with increasing concentrations of the complex, resulting in
an increased in RF2 population and indicating nicking in the plas-
mid DNA by complex 1. In case of bpy containing complex 2, the
intensity of RF2 increased only at a higher concentration of the
complex.
lM for complex 1
l
Complex 2 had no effect on the HepG2 cells up to 10 lM, but com-
plex 1 induced significant cytotoxic effects on the cells (Fig. 6b). It
is evident from MTT assay that the toxic effect of complex 1 is
much more than that of complex 2 and the cytotoxic effect of com-
plex 1 is both time and concentration depended (Fig. 6). The cyto-
toxic effect is likely to be linked with cell cycle arrest, so we have
also performed experiments on cell cycle analysis. As complex 2
was effective in killing cells at a higher concentration, we took
4 lM of complex 2 and 1 lM of complex 1. As seen in Fig. 7, signif-
icant populations of the cells were arrested in the S phase when
they were treated with these complexes. Complex 2 induced more
S-phase arrest at a concentration 4
lM and complex 1 induced S-
3.5. MTT-assay
phase arrest at a much lower concentration (1
lM). The nature of
cell death was also determined by double staining the cells with
propidium iodide and annexin V-FITC. After incubation in the pres-
ence of either of the complexes for 24 h, the cells were stained
simultaneously with propidium iodide and annexin V-FITC and
then subjected to FACS analysis. Using same concentrations of
the complexes as was used in cell cycle analysis, we found complex
The in vitro DNA binding ability of the complexes prompted us
to investigate the effect of the complexes on human cell lines. Hu-
man hepatocellular carcinoma cell line HepG2 was taken and the
cells were incubated in the presence of either of the complexes
or phen for 24 h before they were subjected to an MTT assay. As

164
R. Dey et al. / Polyhedron 48 (2012) 157–166
found a marked difference between the two complexes. Both the
complexes bind with DNA and possibly by base intercalation or
weakly through the minor groove of DNA. Apart from these meth-
ods, the complexes may bind with DNA in such a manner which
distorts the DNA structure, rendering the release of EtBr or Hoechst
33258 from DNA. Both the complexes produced nicking in super
coiled pUC19 DNA and consequently increased RF2 population. In
the DNA migration assay it was found that complex 1 was more
effective in converting supercoiled DNA into RF2 population. This
might be due to the presence of the phen ring which can maintain
planarity much better than bpy, and as a result complex 1 may
have better chance to intercalate with DNA. Though phen interacts
with DNA, in vitro it cannot produce nicking in plasmid DNA, as ob-
served in the gel electrophoresis study. The nicking of the DNA by
the complexes may be due to the presence of the transition metal.
It is possible that free phen dissociates from the DNA or its pres-
ence may not produce any significant shift in the DNA under the
gel electrophoresis conditions. Moreover, the interaction of free
phen with DNA in cells may be responsible for the cytotoxicity of
phen in the MTT assay.
Binding of Cu(II) to DNA and subsequent induction of single
strand breaks was reported in several pieces of literature [47].
Actually it was demonstrated that copper–phen complexes and
their conjugates are useful reagents for studying nucleic acid inter-
actions [57]. Also, the nuclease activity of the copper–phen ion can
be targeted to specific DNA sequences by attachment of the ligand
to the 50 end of complementary deoxyoligonucleotides via a phos-
phoramidate linkage [58]. Moreover, Cu(II) ions, in the presence of
phen, O2 and a reducing agent, degraded DNA with the release of
thiobarbituric-acid-reactive material [59].
In present case, the chances for the availability of copper for
interaction with DNA are poor, if not impossible. Thus the apparent
activity of the complexes is due to their ligands, i.e. bpy or phen,
where we clearly demonstrated that the phen ligand was more ac-
tive than bpy when they were entrapped in same type of com-
plexes. In cell survival assay it was demonstrated that phen
containing complex 1 had a greater cytotoxic effect than bpy con-
taining complex 2. Both the complexes have the same lipophilic
part (phenyl rings of phosphate), but complex 1 might effectively
interfere with DNA metabolisms at much lower concentration
compared to complex 2. In vitro complex 1 produces a nick in
the super coiled DNA, but in vivo it may also interfere with differ-
ent cellular proteins, which are essential for replication. Also, nick-
ing of DNA may lead the cells to arrest in the G1 or G2 rather than
the S-phase, and the cell cycle data suggested that cells were ar-
rested in the S phase after treatment with the complexes. During
the S phase the chromosomal DNA is more accessible to the com-
plex due to its open conformation at the sites of replication and
after binding, it can induce DNA strand breaks, resulting in arrest
in the S-phase. Additionally, complex 1 may interact with cellular
proteins involved in DNA replication and thereby inducing S-phase
arrest. Whatever the reason is, stalled DNA replication may induce
genomic instability, resulting in apoptosis. In the annexin V-FITC
assay, we also noticed most of the dead cells were in early apopto-
sis. Cell cycle arrest and subsequent induction of apoptosis is a
hallmark of many DNA damaging agents. Here also, the observa-
tion that complex 1 induces apoptosis after S-phase arrest impli-
cates a possible therapeutic application.
Fig. 6. Cell survivability assay. (a) HepG2 cells were incubated with increasing
concentrations of the complex (1 or 2) or phen ligand for 24 h and the cells were
subjected to MTT assay. (b) MTT assay done with HepG2 cells when the complexes
were treated for 48 h.
Fig. 7. Cell cycle analysis by FACS along with the statistical analysis: HepG2 cells
were treated with either complex 1 or 2 for 24 h and then the cells were processed
for cell cycle analysis. (a) mock treated, (b) complex 1 treated, (c) complex 2 treated.
Percentage of cells in S-phase after treatment was plotted in the bottom panel.
1 induced 28.5% early apoptotic cells, whereas only 19.75% apopto-
tic cell were observed for complex 2 (Fig. 8).
4. Conclusion
3.6. Comments regarding the biophysical/biological properties
The complexes presented in this report have similar coordina-
tion structures as their compositions are similar, with the only dif-
ference being in the co-ligands phen and bpy. This difference in co-
ligand makes their supramolecular arrangements different. Both
Both complexes have similar coordination structures, only with
a difference with respect to the co-ligand used (phen and bpy), but
during a comparison of the biophysical and biological activity we

R. Dey et al. / Polyhedron 48 (2012) 157–166
165
Fig. 8. Apoptosis study of HepG2 cells (24 h) by annexin V-FITC and propidium iodide staining along with the percentage of apoptotic cells after treatment in the bottom
panel.
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the EtBr–DNA complex and partially displace Hoechst 33258 from
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the Hoechst–DNA complex. The DNA binding activity and cell kill-
ing ability of complex 1 is more effective than those of complex 2,
and the mechanism of cell killing is mediated by apoptosis. Thus it
can be concluded that the slight change in co-ligand may result in
the overall change in the supramolecular structure as well as the
significant variation in biological properties, and thus complex 1
may have some potential implications from a therapeutic point
of view. In summary, this work is a nice example of controlling
structure property relationships.
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Acknowledgements
The authors are gratefully acknowledged UGC for financial sup-
port under the major research scheme. The X-ray diffractometer
facility under the DST FIST program is also gratefully acknowledged.
Appendix A. Supplementary data
CCDC reference numbers 859416 and 859417 contain the sup-
plementary crystallographic data for complexes 1 and 2. These
data can be obtained free of charge via http://www.ccdc.cam.a-
c.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,
at http://dx.doi.org/10.1016/j.poly.2012.08.087.
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