Structure and biological properties of mixed-ligand Cu(II) Schiff base complexes as potential anticancer agents

Article abstract of DOI:10.1016/j.ejmech.2017.04.026

We synthesized two mixed-ligand Cu(II) complexes containing different aroylhydrazone ligands and a pyridine co-ligand, namely, [Cu(L1)(Py)] (C1) and [Cu(L2)(Py)(Br)] (C2) (L1?= (E)-2-hydroxy-N′-((2-hydroxynaphthalen-1-yl)methylene)benzohydrazide, Py?=?pyr

Full text of DOI:10.1016/j.ejmech.2017.04.026

Accepted Manuscript
Structure and biological properties of mixed-ligand Cu(II) Schiff base complexes as
potential anticancer agents
Yi Gou, Jinlong Li, Boyi Fan, Bohui Xu, Min Zhou, Feng Yang
PII:
S0223-5234(17)30283-0
10.1016/j.ejmech.2017.04.026
EJMECH 9373
DOI:
Reference:
To appear in: European Journal of Medicinal Chemistry
Received Date: 25 January 2017
Revised Date: 10 April 2017
Accepted Date: 11 April 2017
Please cite this article as: Y. Gou, J. Li, B. Fan, B. Xu, M. Zhou, F. Yang, Structure and biological
properties of mixed-ligand Cu(II) Schiff base complexes as potential anticancer agents, European
Journal of Medicinal Chemistry (2017), doi: 10.1016/j.ejmech.2017.04.026.
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Structure and biological properties of mixed-ligand Cu(II) Schiff base complexes
as potential anticancer agents
Yi Gou1*, Jinlong Li1, Boyi Fan1, Bohui Xu1, Min Zhou,1 Feng Yang1, 2*
1 School of Pharmacy, Nantong University, Nantong, Jiangsu, China.
2
State Key Laboratory for the Chemistry and Molecular Engineering of Medicinal
Resources, Ministry of Science and Technology of China. Guangxi Normal University,
Guilin, Guangxi, China.
*Corresponding authors: Yi Gou and Feng Yang
Email: sanchuanok@126.com
Phone/Fax : 86-773-584-8836
Address: 9 Seyuan Road, Nantong, Jiangsu, China. Zip code: 226019
Keywords: aroylhydrazone Cu(II) complexes; anti-metastatic activity; anticancer
activity; anticancer mechanism.
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Abstract
We synthesized two mixed-ligand Cu(II) complexes containing different
aroylhydrazone ligands and a pyridine co-ligand, namely, [Cu(L1)(Py)] (C1) and
[Cu(L2)(Py)(Br)]
(C2)
(L1
=
(E)-2-hydroxy-N'-((2-hydroxynaphthalen-1-
yl)methylene)benzohydrazide, Py = pyridine, L2 = (E)-2-hydroxy-N'-(phenyl(pyridin-
2-yl)methylene)benzohydrazide), and assessed their chemical and biological
properties to understand their marked activity. C2 showed better anticancer activity
than C1 in various human cancer cell lines, including the cisplatin-resistant lung
cancer cell line A549cisR. Both Cu(II) complexes, especially C2, displayed
promising anti-metastatic activity against HepG2 cells. Spectroscopic titration and
agarose gel electrophoresis experiments indicated that C2 exhibited binding affinity
toward calf-thymus DNA and efficient pBR322 DNA-cleaving ability. Further
mechanistic studies showed that C2 effectively induced DNA damage and thus led to
cell cycle arrest at the G2/M phase, and also stimulated mitochondrial dysfunction
mediated by reactive oxygen species and caspase-dependent apoptosis.
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1. Introduction
Cancer has been, and still remains, one of the leading causes of morbidity and
mortality worldwide [1,2]. Investigations performed by the American Cancer Society
showed that the global burden is expected to increase to 21.4 million new cancer
cases and approximately 13.2 million cancer deaths by 2030 [3]. Although traditional
cancer chemotherapy has been quite beneficial in improving survival rates in the past
few decades, it has also resulted in many adverse effects, including limited efficiency,
multidrug resistance, and undesirable side effects [3-5]. With the increase in
morbidity, medicinal chemistry is taking a growing interest in the development of
metal complexes for use as anticancer drugs [6-9]. Depending on the choice of metal,
its oxidation state and coordination geometry, and the types and number of
coordinated ligands, as well as kinetic properties, metal complexes have the potential
for unique mechanisms of drug action [9]. In recent years, a significant number of
metal complexes have been evaluated in vitro and in vivo, and some are presently
being clinically tested [9,10]. Among these, copper-based complexes have been
proposed as promising anticancer agents [7].
Copper is an essential micronutrient for all organisms and is employed as a
catalytic cofactor or a structural component of proteins, and consequently it is
involved in critical biological functions such as energy metabolism, oxygen transport,
enzyme activity, and cell signaling [11]. Not surprisingly, copper is involved in
cancer development and progression and can promote angiogenesis and cancer growth,
invasion, and metastasis [12]. Many studies have shown that numerous malignant
tissues (e.g., in breast, ovarian, cervical, and lung cancers and leukemia) have been
shown to take up greater amounts of copper than normal tissues [7,11]. On the other
hand, aroylhydrazones that possess a–CONHN=CH−azomethine group constitute an
important class of biologically active molecules that have attracted the interest of
medicinal chemists owing to their various biological and pharmaceutical activities,
including anticancer, anti-inflammatory, and anti-tubercular activities [13-15]. In
addition, aroylhydrazone derivatives are emerging as compounds of interest in
medical biotechnology because of the color intensification properties that accompany
their deprotonation [16]. In particular, aroylhydrazones, as an important class of
Schiff base ligands, can exhibit enol–keto tautomerism and can coordinate in a range
of deprotonated states such as neutral, monoanionic, dianionic, or tetraanionic forms,
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and thus can form a variety of complexes with different transition metals [16].
Interestingly, it has previously been reported that anticancer activity of a variety of
metals in complex with aroylhydrazones is higher than that of the ligand alone [17-
19]. Therefore, the development of aroylhydrazone complexes that chelate copper for
anticancer therapies has shown significant promise.
Very recently, the introduction of co-ligands (such as pyridine, imidazole, diimine,
and quinolone ligands) into copper complexes has been found to enhance their DNA
binding affinity and give rise to prominent anticancer activity [20-22]. In the present
study, we have prepared two mixed-ligand Cu(II) complexes containing different
derivatives of aroylhydrazone and pyridine (Scheme 1), namely, [Cu(L1)(Py)] (C1)
and [Cu(L2)(Py)(Br)] (C2) (L1 = (E)-2-hydroxy-N'-((2-hydroxynaphthalen-1-
yl)methylene)benzohydrazide, Py = pyridine, L2 = (E)-2-hydroxy-N'-(phenyl(pyridin-
2-yl)methylene)benzohydrazide). Their cytotoxic activity against a large panel of
human cancer cell lines, including cisplatin-resistant and –sensitive cells, was
investigated. Although numerous cytotoxic aroylhydrazone copper complexes have
been reported to date, few data have so far been reported on the mechanisms
underlying their antiproliferative activity [7]. Thus, further experiments have been
carried out to determine the mechanisms by which aroylhydrazone Cu(II) complexes
inhibit cell proliferation and induce cell death.
2. Experimental section
All the chemicals and solvents used for the syntheses were of analytical grade.
The plasmid DNA pBR322, Calf thymus (CT)-DNA and loading buffer (10×) were
purchased from Fermentas (Lithuania). Elemental analyses of C, N and H were
performed on a PE 2400 series II CHNS/O analyzer. Infrared (IR) pectra (KBr pellets)
of all samples were recorded on an Interspec 2020 FTIR spectrophotometer in the
frequency of 400−4000 cm−1. Electronic spectra were measured in DMSO-buffer
solution on a Cary 1E UV−Visible spectrophotometer.
2.1 Synthesis and structures of Cu(II) complexes
2.1.1 Synthesis of L ligand Cu(II) complexes
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Syntheses of the Schiff base ligands: L1 and L2. (E)-2-hydroxy-N'-((2-
hydroxynaphthalen-1-yl)methylene)benzohydrazide (L1) was synthesized according
to a literature procedure [23]. In brief, 2-hydroxy-1-naphthaldehyde (1.72 g, 10
mmol), 2-hydroxybenzohydrazide (1.52 g, 10 mmol) and 50 ml methanol were mixed.
After stirring 15 min at 373 K, the resulting mixture was kept in air for 1 h, affording
the title compound as a yellow colored solid. Yield: 76%. Anal. Calcd for C18H14N2O3
(306.32): C, 70.58; H, 4.61 and N, 9.15. Found: C, 70.83; H, 4.77 and N, 9.36. IR
(KBr, cm−1): 3071m, 2539m, 1618vs, 1590s, 1498vs, 1435s, 1358s, 1290s, 1211vs,
1145m, 1112m, 1077m, 1042m, 899m, 846m, 817m, 762s, 647w, 586w, 555w, 471w.
1
MS (ESI) m/z 305 ([M – H]–). H NMR (400 MHz, DMSO-d6) δ 12.75 (1H, s), 9.55
(1H, s), 8.32 (1H, d, J = 8.6 Hz), 7.94 (2H, m), 7.89 (1H, brd, J = 8.2 Hz), 7.60 (1H,
m), 7.48 (1H, m), 7.41 (1H, t, J = 7.4 Hz), 7.24 (1H, d, J = 9.0 Hz), 7.02 (2H, m); 13C
NMR (100 MHz, DMSO-d6) δ 164.0, 158.8, 158.1, 147.7, 134.0, 132.9, 131.7, 128.9,
128.8, 127.8, 127.7, 123.6, 121.0, 119.2, 118.9, 117.3, 115.7, 108.6 (Figs. S1 and S2).
The (E)-2-hydroxy-N'-(phenyl(pyridin-2-yl)methylene)benzohydrazide ligand (L2)
was synthesized by mixing equimolar amounts of 2-hydroxybenzohydrazide (1.52 g,
10 mmol) with phenyl(pyridin-2-yl)methanone (1.83 g, 10 mmol) in methanol (50
mL). The reaction mixture was refluxed on a water bath for 6 h. Then the mixture was
kept in air for a week and light yellow colored solid was obtained. Yield: 69%. Anal.
Calcd for C19H15N3O2 (317.74): C, 71.91; H, 4.76 and N, 13.24. Found: C, 71.65; H,
4.77 and N, 13.02. IR (KBr, cm−1): 3175w, 3051m, 2833w, 1645vs, 1602s, 1576s,
1485m, 1412m, 1391m, 1301w, 1331s, 1287vs, 1188s, 1161m, 957w, 818w, 793m,
1
739m, 691m, 538w, 419w. MS (ESI) m/z 316 ([M – H]–). H NMR (400 MHz,
DMSO-d6) δ 13.14 (1H, s), 8.88 (1H, s), 8.04 (1H, m), 7.98 (1H, m), 7.63 (1H, m),
7.56 (2H, m), 7.46 (3H, m), 7.38 (2H, m), 6.96 (2H, m); 13C NMR (100 MHz,
DMSO-d6) δ 162.0, 156.2, 151.6, 149.6, 149.6, 138.0, 137.1, 133.5, 131.0, 129.4,
128.4 × 2 , 128.1 × 2, 125.0, 124.7, 119.7, 117.8, 116.7 (Figs. S3 and S4).
Synthesis of [Cu(L1)(Py)] (C1). This complex was prepared by adding a
methanolic solution of L1 ligand (2 mmol) and pyridine (Py, 2 mmol) to a solution of
CuBr2 (0.44 g, 2 mmol) in anhydrous methanol, and the mixture was heated to reflux
for 2 h. The greenish-blue solution was cooled to room temperature and then filtered.
Bright green crystals suitable for X-ray crystallography were obtained on slow
evaporation of the reaction mixture over a period of 3–7 days. Yield: 73%. Anal.
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Calcd for C23H17CuN3O3 (446.94): C, 61.81; H, 3.83 and N, 9.40. Found: C, 61.57; H,
3.91 and N, 9.55. IR (KBr, cm−1): 3110w, 3040w, 1604vs, 1531vs, 1509vs, 1480s,
1452s, 1394s, 1370s, 1334s, 1304m, 1255m, 1139m, 1196w, 1040w, 971w, 939w,
822m, 747vs, 695m, 676w, 667w, 605w, 525w, 474w, 462w.
Synthesis of [Cu(L2)(Py)(Br)] (C2). The procedure of C2 synthesis was the same
as that of C1 except that L1 ligand was replaced by L2 ligand. Bright green crystals
suitable for X-ray crystallography were obtained on slow evaporation of the reaction
mixture after several days. Yield: 65%. Anal. Calcd for C24H19BrCuN4O2 (538.88): C,
53.49; H, 3.55 and N, 10.38. Found: C, 53.21; H, 3.53 and N, 10.27. IR (KBr, cm−1):
3110w, 3093w, 3067w, 3003w, 1604m, 1573w, 1489m, 1446vs, 1391w, 1362w,
1219m, 1153w, 1077m, 1015w, 944w, 866w, 755vs, 682vs, 643w, 441m.
2.1.2 Crystal structures determination of Cu(II) complexes
The data collections on C1 and C2 were performed on a Bruker SMART Apex II
CCD equipped with a graphite−monochromated Mo−Kα (λ = 0.071073 nm) radiation.
Empirical absorption correction was applied by the SADABS program. The structures
of C1 and C2 were solved by direct methods and refined with full−matrix
least−squares methods using the SHELXTL version 5.1 software package [24].
Anisotropic thermal parameters were applied to all non−hydrogen atoms. H atoms
were added theoretically, and refined as riding atoms with a common fixed isotropic
thermal parameter. The selected crystallographic information of the Cu(II) complexes
are summarized in Table 1 and Table 2. Crystallographic data for C1 and C2 have
been deposited at the CCDC (Cambridge Crystallographic Data Centre), reference
numbers 1511939 for C1 and 1511940 for C2.
2.1.3. Computational details
Full geometry optimization of all structures was carried out at the density
functional theory (DFT) level of theory using the gradient-corrected correlation
functional of Lee, Yang, and Parr in combination with Becke’s three-parameter
hybrid exchange functional (B3LYP) with the help of the GAMESS suite of codes
[25-27]. The calculations were performed using the 6−31+G(d,p) basis set for all
atoms. Solvation effects were incorporated in the calculations using the integral
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equation formalism model (IEF−PCM) with methanol as a solvent [28]. The
experimental x-ray structure was chosen as the starting atomic coordinates for the
optimizations.
2.2. Cu(II)−ligand binding studies in solution
Cu(II)-ligand binding studies were assessed as method described previously [29].
Briefly, Cu(II)-ligand binding studies were carried out in PBS buffer having 10%
DMSO. To the ligand (50 µM) solution of each sample, increasing concentration of
CuBr2 was added gradually. For every addition, the sample was incubated for 5 min,
and then the absorption spectrum was recorded.
2.3. Cell lines
Culture medium DMEM (with L−glutamin), Foetal bovine serum (FBS),
phosphate buffered saline (PBS) and Antibiotic−Antimycotic were from E.U. Gibco
BRL. Human breast cancer cell lines (MCF-7), human liver hepatocellular carcinoma
cell lines (HepG2), human hepatocellular cell line (Bel-7402), human lung carcinoma
cell lines (A549), cisplatin-resistant A549 (A549cisR) and normal lung fibroblast
cells (WI-38) were maintained in DMEM supplemented with 10% FBS, penicillin (50
U/mL), streptomycin (50 mg/mL) at 37 °C and 5% CO2.
2.4. 3−(4,5−Dimethylthiazol−2−yl)−2,5−diphenyltetrazolium bromide (MTT) assays
The cytotoxicities of the compounds toward several human cancer cell lines were
investigated using an MTT assay. Briefly, the 100 µL of cell suspensions (5 × 104
cells/mL) was seeded in triplicate in 96−well plates and incubated for 24 h at 37 °C in
5% CO2. Then various concentrations of test compound (0.1-40 µM) were added and
incubated for 48 h at 37 °C under conditions of 5% CO2. The medium not containing
the compounds served as the control. After 48 h, each well was treated with 10 µL of
a 5 mg/mL MTT saline solution at 37 °C. After 4 h of incubation, DMSO (100 µL)
was added to each well. Then the absorbance of each well was read by enzyme
labeling instrument with 570/630 nm double wavelength measurement. All the tests
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were repeated in at least three independent experiments, and the mean values were
taken.
2.5. Scratch wound healing assay
To check the effect of the Cu(II) complexes on cancer cells migration, wound
healing assay was performed in HepG2 cell line. In brief, cells were seeded in a 6-
well plate and allowed to recover for a confluent cell monolayer formed in each well
(>90% confluence). The cell monolayer was wounded with a sterile 200 µL pipette tip
to generate a clean wound area. Subsequently, cell monolayers were washed once
with PBS, and then incubated in medium with or without Cu(II) complexes. The
closing of the gaps was observed during 24 h using a bright field microscope.
2.6. Adhesion assay
The cell adhesion assay was carried out as previously described [30]. Briefly,
fibronectin (5 mg/mL) was coated on 96-well plates at 37 °C for overnight and then
the wells were washed and blocked in 1% bovine serum albumin for 1 h. Serum-
starved HepG2 cells were exposed to Cu(II) complexes (0.75, 1.5, or 3 µM) for 24 h
before seeding. A control without treatment of Cu(II) complexes was also performed.
Target cells were harvested and seeded to fibronectin coated 96-well plates (2 × 105
cells/mL) and then incubated for 1 h at 37 °C. Wells were then washed and then
colorimetric MTT assay was employed to analyze the adhesion ability of HepG2 cells.
2.7. Anticancer mechanism of C2
2.7.1. DNA-binding
UV spectroscopy. Electronic absorption spectra were determined by keeping the
Cu(II) complex' concentration constant (5 µM), and varying the concentration of calf-
thymus DNA (CT-DNA) in 5 mM Tris-HCl/50 mM NaCl buffer (pH 7.2). For each
addition, the sample was shaken and kept for 10 min at equilibrium before recording
its spectrum, and then the absorption spectra were recorded in the range of 200–600
nm. The binding constant (Kb) of the Cu(II) complex was measured using the
following Wolfe-Shimer equation [31]:
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[DNA]/(εa − εf) = [DNA]/(εb − εf) + 1/Kb(εb − εf)
(1)
where the apparent molar extinction coefficients εa = Aobs/[complex], εb = the
extinction coefficient for the copper(II) complex in the fully bound form and εf = the
extinction coefficient of free (unbound), respectively. [DNA] is the concentration of
CT-DNA in base pairs.
Fluorescence spectroscopy. The competitive binding study was carried out by
maintaining the ethidium bromide (EB, 8 µM) and CT-DNA (10 µM) in Tris-HCl
buffer (pH 7.2), and increasing the concentrations of the Cu(II) complex. Then
emission spectra were recorded from 530 to 700 nm with an excitation wavelength set
at 510 nm. Further, the apparent binding constant (Kapp) of the copper(II) complex has
been calculated from equation (2) [32]:
KEtBr [EB] = Kapp [complex]
(2)
where KEtBr is 1 × 107 M−1 and [complex] is the concentration of the complex causing
50% reduction of the fluorescence intensity of EB-DNA.
Circular dichroism (CD) spectral. The CD spectra were run on a JASCO J-810
CD spectropolarimeter. The concentration of CT-DNA was kept constant (100 µM)
and the concentration of the copper(II) complex varied from 10 to 20 µM. The
working solution was incubated for 15 min after each addition.
Viscometric determination. The relative change in viscosity was measured using
an Ubbelodhe viscometer, thermostated in a water bath maintained at 30 ± 0.5°C. The
concentration of DNA was 200 µM and increasing concentrations of the copper(II)
complex and EB were used. Each complex solution was measured three times for
accuracy and an average flow time was calculated. The results were presented as
(η/ηo)1/3 versus the ratio of the concentration of the complexes to CT-DNA, where ηo
is the viscosity of CT-DNA alone and η is the viscosity of CT-DNA in the presence of
the copper(II) complex and EB in Tris-HCl buffer (pH 7.2).
2.7.2. DNA cleavage experiments
The DNA cleavage activity of the Cu(II) complex was studied by agarose gel
electrophoresis using supercoiled pBR322 plasmid DNA. Reactions using supercoiled
pBR322 plasmid DNA in 1 × TBE (Tris−borate−EDTA) buffer (pH 7.26) was treated
with the Cu(II) complex (12.5−50 µM) by dilution with the 1 × TBE buffer to a total
volume of 16 µL and incubated at 37 °C. After 5 h, the loading buffer was added. The
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reaction mixture was then loaded on a 1% agarose gel containing 1% (v/v) goldview,
and analyzed with gel electrophoresis in 1 × TBE buffer at 80 V.
In order to investigate the DNA cleavage mechanism induced by the Cu(II)
complex, the agarose gel electrophoresis was also carried out in the presence of some
radical scavengers. Each sample was treated and analyzed according to the above
methods.
2.7.3. Cell cycle distribution analysis
Cell cycle distribution was carried out in HepG2 cells and analyzed by PI
(propidium iodide) staining and flow cytometry. In brief, HepG2 cells were exposed
to the tested Cu(II) complex at the indicated concentrations. The cells without the
treatment were used as control. After 24 h, cells were collected and fixed with 70%
ethanol and kept at −20 °C for 24 h. Next, the cells were again washed with PBS and
stained with PI solution (50 µg/mL) containing RnaseA (100 µg/mL) and and
analyzed by flow cytometry (FACScan, Bection Dickinson, San Jose, CA).
2.7.4. Apoptosis by Flow Cytometry
HepG2 cells seeded into 6-well plates were exposed to the tested Cu(II) complex
(1.5 and 3 µM) for 24 h. The cells without the treatment were used as control. After
that, the cells were extensively washed with PBS and subjected to flow cytometer
(FACScan, Bection Dickinson, San Jose, CA) for apoptosis analysis using the
Annexin V FITC kit following manufacturer’s protocol. Data were analyzed with
FlowJo Software (Tree Star, OR, USA).
2.7.5. Acridine orange/ethidium bromide (AO/EB) double staining
Characteristic apoptotic cell changes were observed using AO/EB double staining
in HepG2 cancer cell line. HepG2 cells were cultured on 6-well plates and were
incubated with and without the test compounds for 24 h at 37 ºC. Cells were washed
with ice-cold PBS twice, stained with a medium containing AO (100 µg/mL) and EB
(100 µg/mL) solution in the dark for 10 min. Afterward, the cells were washed with
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PBS twice and replaced with 1 mL dye-free DMEM for being examination by a
fluorescence microscopy (OLYMPUS Co., Japan).
2.7.6. Intracellular reactive oxygen species (ROS) measurements
Intracellular ROS generation was measured using the 2’,7’-dichlorofluorescein
diacetate (H2DCF-DA) fluorescent probe. Oxidation of the non-fluorescent molecule
H2DCF-DA by cellular ROS converts it into the fluorescent product, 2’,7’-
dichlorofluorescein (DCF). HepG2 cells were treated with and without the tested
Cu(II) complex at 37 ºC. After 24 h, cells were collected by and incubated for 30 min
at 37 ºC with 10 µM H2DCF-DA probe in serum-free DMEM. Intracellular ROS was
measured as an increase in green cytosolic DCF fluorescence using a flow cytometer
(FACScan, Bection Dickinson, San Jose, CA).
2.7.7. The change of mitochondrial membrane potential assay
Mitochondrial membrane potential was analyzed by the 5,5’,6,6’-tetrachloro-
1,1’,3,3’-tetraethyl-benzimidazolylcarbocyanine iodide (JC-1) probe (Beyotime
Jiangsu China). HepG2 cells were incubated with or without the tested Cu(II)
complex for 24 h at 37 ºC and were then washed 3 times with warm PBS. The cells
were then collected and resuspended in 0.5 mL of PBS buffer containing 10 mg/mL
of JC-1 dye. After 20 min’ incubation at 37 ºC in the dark, the cells were washed
twice with PBS, and were analyzed immediately by flow cytometry (FACScan,
Bection Dickinson, San Jose, CA). 10,000 events were acquired for each sample.
2.7.8. Western blot analysis
Western blotting was performed as previously described [33,34]. Primary
antibodies: rabbit anti-β-actin (Cell Signaling), mouse anti-CDK1(Cell Signaling),
mouse anti-Cyclin B1 (Cell Signaling), mouse anti-cleaved caspase-3 (Abcam),
mouse anti- cleaved PARP-1 (Abcam), rabbit anti-Bax (Abcam), mouse anti-Bcl-2
(Abcam), and mouse anti-Bcl-xl (Abcam). Secondary antibodies: horseradish
peroxidase-conjugated donkey anti-rabbit and horseradish peroxidase-conjugated
sheep anti-mouse (Abcam).
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2.8. Statistical analysis
All experiments were repeated 3 to 5 times. Data were compared using Student's t
test. Results were expressed as mean ± SD (standard deviations) and considered to be
significant when p < 0.05.
3. Results and Discussion
3.1. Characterization of Cu(II) complexes
Complex C1 crystallizes in the monoclinic space group P21/c, and its molecular
structure consists of a Cu(II) ion chelated by one Schiff base ligand L1 and one
pyridine molecule. As shown in Fig. 1A, the Cu(II) metal center, which is coordinated
by one nitrogen atom [Cu1−N3 = 2.011 Å] of pyridine and two oxygen atoms
[Cu1−O1 = 1.928 Å and Cu1−O2 = 1.887 Å] and one nitrogen atom [Cu1−N2 =
1.912 Å] of L1, adopts a square-planar coordination geometry. The Cu1−N2 distance
is significantly shorter than the Cu1−N3 distance, which can be attributed to stronger
π-back-bonding in the Cu−N(imine) bond than in the Cu−N(pyridine) bond. In the
solid state, there exist two weak interactions: π···π stacking arising from the aromatic
rings of the L1 ligands (face-to-face distance=3.352 Å; center-to-center distance =
3.695 Å) and strong intermolecular hydrogen bonding involving the hydroxy oxygen
atom (O3) and the nitrogen atom (N1) in the L1 ligand (O3···N1 = 2.602 Å and the
O3−H3···N1 angle is 146.5º).
Single-crystal x-ray diffraction reveals that complex C2 crystallizes in the
orthorhombic system with the space group P212121. As shown in Fig. 1B, the Cu(II)
center adopts a square-pyramidal geometry with the apical position occupied by a Br
atom. The Cu−N, Cu−Br, and Cu−O bond lengths observed here are within the range
reported for other similar Cu(II) complexes [34,35]. The coordination polyhedron
around the Cu1 center could be described as a slightly distorted square pyramid (τ =
0.17) [36], with the metal displaced from the N1/N2/O1/N4 basal plane (maximum
displacement = 0.038 Å for the N1 atom), the Br atom at the apex, and the Cu atom
displaced by 0.295 Å toward the Br atom from the mean basal plane. The
intermolecular hydrogen bonding in this complex (O2···N3 = 2.602 Å and the
O2−H2A···N3 angle is 144.0º) is very similar to that in C1.
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Interestingly, the L1 and L2 ligands can be represented by two tautomeric forms,
namely, the enol form (C=N−N=C−OH) and the keto form (C=N−NH−C=O). Here
the C7−O1 bond distances in C1 (1.290 Å) and C2 (1.274 Å) are longer than the
corresponding values in the keto forms of L1 (1.243 Å) [23] and L2 (1.228 Å) [35],
respectively, which supports the formation of alkoxide moieties in the ligands on
complexation. To investigate the energies involved in the process of tautomerization
of the L1 and L2 ligands in methanol, quantum chemical calculations were performed
(Fig. 1C and 1D). As expected, the keto forms of L1 and L2 were lower in energy by
1.3 and 3.5 kcal/mol in methanol, respectively, which implies that the L1 and L2
ligands exist mainly in their keto forms in methanol solution and that conversion to
the enol forms is necessary prior to complexation. Moreover, the barriers in methanol
for the conversion of the enol forms to the keto forms are predicted from theory to be
about 62.2 and 65.9 kcal/mol, respectively.
3.2. Studies of Cu(II)−ligand binding in solution
The formation processes of complexation of L1 or L2 with Cu(II) were studied by
UV-visible spectroscopy in PBS buffer containing 10% DMSO. As shown in Fig. 2,
L1 exhibited a strong absorption band at 376 nm in the buffer. Notably, upon addition
of increasing amounts of Cu(II) to L1, a new peak at 438 nm was observed. Upon
incremental addition of Cu(II) (0−1.0 equiv), the intensity of a new peak at 438 nm
increased with a simultaneous decrease in absorption at 376 nm but saturated at
approximately 0.5 equiv with respect to L1, indicating that L1 formed 1:2 Cu(II):L1
complexes with Cu(II). Similar results were also observed for L2 (Fig. 2). These
binding studies demonstrate that L1 and L2 preferred to form 1:2 (Cu(II)/ligand)
complexes in an aqueous solution. However, the formation of 1:2 Cu(II):L1/L2
complexes can be inhibited by some small molecule ligands [29]. For example, the
coordination of pyridine, as a coordinating ligand, with Cu(II) was confirmed by
single-crystal x-ray diffraction in the solid state.
3.3. Cytotoxic activity of the Cu(II) complexes
The in vitro anticancer activity of the Cu(II) complexes was assessed using the
MTT assay. The widely used metal-containing drug cisplatin was included as a
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control. The IC50 values obtained are shown in Table 3. L1, L2, and Cu2+ displayed
relatively low activity against cancer cells, but C1 and C2 exhibited high cytotoxicity
against cancer cells, which implies that the chelation of the Cu(II) ion by L1 or L2 is
probably responsible for the observed highly cytotoxic properties of C1 and C2. In
addition, the introduction of a pyridine moiety as a co-ligand may affect the
hydrophobicity and coordination geometry of the Cu(II) complexes and possibly
enhance the anticancer activity of C1 and C2. Both Cu(II) complexes displayed a
broad spectrum of inhibition against human cancer cells, with IC50 values ranging
from 1.57 to 5.23 µM, which were lower than those of cisplatin. Notably, neither of
the Cu(II) complexes exhibited cross-resistance with cisplatin, as typified by their
ability to indiscriminately kill the cisplatin-resistant and -sensitive lung cancer cell
lines A549cisR and A549. C2 exhibited higher in vitro cytotoxicity against the
selected tumor cell lines than C1, as evidenced by its lower IC50 values. As a measure
of therapeutic potential, we further determined the cytotoxicity of the Cu(II)
complexes against the normal lung fibroblast cell line WI-38. In general, both Cu(II)
complexes were less potent toward WI-38.
3.4. Assay of anti-metastatic effect
Because HepG2 cells exhibited the highest sensitivity to C1 and C2, this cell line
was used for further investigation of the anti-metastatic effect of C1 and C2. Cell
migration takes place in several diseases, especially in cancer, leading to metastasis
and invasion [37]. Therefore, we investigated the effect of both Cu(II) complexes on
cancer cell migration in HepG2 cells via a scratch wound healing assay. Fig. 3A
shows that untreated control cells migrated rapidly, whereas cells treated with C1 and
C2 did not migrate at such a high rate. This result indicates that both Cu(II)
complexes may have great potential to display anti-metastatic activity via the
inhibition of cancer cell migration. To further assess whether C1 and C2 regulate the
adhesion of cells to extracellular matrix components, HepG2 cells were pretreated
with different concentrations of the Cu(II) complexes prior to an adhesion assay on
fibronectin. As shown in Fig. 3B, in comparison with the controls C1 and C2, in
particular C2, decreased the adhesion capability of HepG2 cells more significantly in
the adhesion assay. Upon confirmation of the excellent cytotoxicity and anti-
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metastatic properties of C2, further biophysical and cellular (in HepG2 cells) studies
were performed to understand its mechanism of action.
3.5. Possible anticancer mechanism of C2
DNA is the primary intracellular target of many metallodrugs [7,38]. Therefore,
metallodrug–DNA interactions are of paramount importance for understanding their
mechanisms. The character of the binding of C2 to CT-DNA has been investigated by
using multiple biophysical techniques such as emission and absorption spectral
studies. In this study, the UV spectra of C2 in the absence and presence of CT-DNA
were recorded (Fig. 4A). In the absence of DNA, the UV spectra of C2 exhibited two
absorption bands at around 296 nm and 378 nm owing to the presence of the phenyl
ring and pyridine scaffold. Upon the addition of CT-DNA to the complex, there was
an increase in the absorption intensity or “hyperchromism” (296→292 nm), which
may indicate that there were interactions between DNA and C2 via electrostatic
intercalation and/or groove binding modes, which led to small perturbations [39]. The
value of the binding constant (1.86 ± 0.11 × 105 M−1) obtained for C2 was higher than
that for the anticancer drug cisplatin (3.20 ± 0.15 × 104 M−1) [40]. A competitive
binding assay was carried out using ethidium bromide (EB) as a probe to further
clarify the DNA binding mechanism of C2. The emission spectra of the EB–CT-DNA
system in the absence and presence of C2 are shown in Fig. 4B. The addition of C2
resulted in a decrease in the fluorescence intensity of the emission band of the
EB−CT-DNA system, which indicated that the Cu(II) complex efficiently competed
with EB for intercalative binding sites on CT-DNA by replacing EB, or that groove
binding of the Cu(II) complex to CT-DNA led to the transfer of energy/electrons from
the guanine bases of DNA via metal-to-ligand charge transfer in the Cu(II) complex
[41]. According to eqn. (2), the Kapp value (3.76 ± 0.21 × 106 M−1) of C2 is less than
the binding constant of classical intercalators and metallointercalators (107 M−1),
which suggested that the interaction between the Cu(II) complex and DNA occurred
via a moderate intercalative mode. Circular dichroism (CD) spectroscopy is a useful
method for monitoring changes in the structure of DNA during small molecule–DNA
interactions. As shown in Fig. 4C, the addition of C2 to a solution of CT-DNA
induced an increase in the intensity of the positive band at ca. 275 nm and a decrease
in the intensity of the negative-ellipticity band at ca. 245 nm, which suggested that the
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stacking mode and the orientation of base pairs in DNA were disturbed. To further
investigate the nature of the binding of the Cu(II) complex to DNA, measurements of
the viscosity of solutions of DNA incubated with EB and C2 were carried out. As
shown in Fig. 4D, upon continuous addition of the Cu(II) complex the relative
viscosity of DNA increased consistently, which suggested that the complex bound to
DNA in an intercalative mode.
It has been reported that many metal complexes are capable of catalyzing the
cleavage of DNA [42]. We therefore examined whether C2 exhibited DNA-cleaving
activities by an agarose gel electrophoresis assay using pBR322 DNA. As shown in
Fig. 4E, C2 could efficiently relax Form I (supercoiled form) of pBR322 DNA into
Form II (open circular form) in a concentration-dependent fashion. It is known that
DNA can be cleaved via an oxidative pathway. To confirm whether reactive oxygen
species (ROS) were, at least in part, responsible for the cleavage of DNA,
comparative cleavage reactions were carried out to determine the mechanism of the
cleavage of pBR322 plasmid DNA by C2 in the presence of various radical
scavengers such as DMSO and EtOH as hydroxyl radical (HO•) scavengers, NaN3 as a
singlet oxygen (1O2) scavenger, and KI as a hydrogen peroxide (H2O2) scavenger.
Furthermore, we conducted the cleavage reaction of C2 in the presence of the metal
ion-chelating agent EDTA. It can be seen from Fig. 4F that DMSO (Lane 3), EtOH
(Lane 4), and NaN3 (Lane 5) did not significantly inhibit DNA cleavage, which ruled
1
out the possibility that HO• and/or O2 were involved in DNA cleavage. However, in
the presence of KI, DNA cleavage was significantly inhibited (Lane 6), which
suggested that the cleavage of DNA by C2 probably proceeded via an oxidative
mechanism involving H2O2 species. Interestingly, EDTA efficiently inhibited DNA
cleavage (Lane 7), which suggested that the complexation of ligands with copper ions
was crucial for cleavage. Taken together, these results suggest that C2 can induce
DNA damage via interacting with DNA [43].
As DNA damage is thought to affect the cell cycle and/or apoptosis, we therefore
monitored features related to these pathways. In this study, the effect of C2 on the cell
cycle was monitored by measuring the DNA content with the help of flow cytometry.
Flow cytometric analysis of DNA (Fig. 5A) revealed that C2 arrested the cell cycle at
the G2/M phase in a dose-dependent manner (ca. threefold increase in the G2/M
population after incubation at a concentration of 3 µM). Cyclin complexes, together
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with cyclin-dependent kinases, are major control switches in progression of the cell
cycle [44]. To further elucidate the mechanisms involved in the anticancer activity of
C2, the expression of cyclin-dependent kinase 1 (CDK1) and cyclin B1 was examined.
The results in Fig. 5B show that after treatment with C2 the expression levels of
CDK1 and cyclin B1 were markedly suppressed in a concentration-dependent manner.
These results indicate that the inhibition of CDK1 and cyclin B1 may represent a
mechanism by which the Cu(II) complex exerts its potent anticancer activity.
It is well known that most small molecule anticancer drugs exert their cytotoxic
effects via apoptosis, and we therefore monitored features related to this pathway.
Here, the annexin V/propidium iodide dual staining method was used to determine
whether the death of cancer cells incubated with C2 was induced by apoptosis. As
shown in Fig. 6A, C2 efficiently induced apoptosis in HepG2 cells in a concentration-
dependent manner. Apoptosis is commonly characterized by distinct morphological
features such as blebbing, chromatin condensation, nuclear fragmentation, or the
formation of apoptotic bodies [45]. To gain more evidence of the induction of
apoptotic cell death by the Cu(II) complex, the apoptosis-inducing properties of C2
were further examined by fluorescence microscopy using AO/EB staining. HepG2
cells treated with C2 displayed a dose-dependent morphological change. As shown in
Fig. 6B, chromatin condensation, nuclear fragmentation, and the formation of
apoptotic bodies occurred following the treatment of HepG2 cells with 3 µM C2 for
24 h. The results of the cell cycle and apoptosis experiments demonstrate that C2 can
effectively induce DNA damage and thus lead to cell cycle arrest and apoptosis in
HepG2 cells.
Mitochondrial dysfunction plays a vital role in triggering various apoptotic
pathways [46]. To establish whether apoptosis induced by C2 is related to
mitochondrial dysfunction, the mitochondrial membrane potential (∆ψm) was
measured by flow cytometry using staining with JC-1 (5,5',6,6'-tetrachloro-1,1',3,3'-
tetraethyl-benzimidazolyl-carbocyanine iodide). The results in Fig. 7A show that the
treatment of HepG2 cells with C2 caused a decrease in ∆ψm to varying degrees, which
indicates the activation of mitochondrially mediated apoptosis.
Mitochondrial dysfunction and the elevation of reactive oxygen species (ROS)
levels are two closely related critical events in cell apoptosis [47]. Most previously
reported metal compounds induce apoptosis in the respective cancer cells via the
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generation of ROS [48-50]. The formation of ROS was investigated in HepG2 cells in
the presence of C2 using the DCFH-DA (2',7'-dichlorofluorescein diacetate)
fluorescent probe. As shown in Fig. 7B, the incubation of cells with C2 led to a shift
in the fluorescence peak to the right relative to the control cells, which demonstrated
that C2 significantly increased intracellular ROS levels in a concentration-dependent
manner. Furthermore, the intracellular formation of ROS was quantified and the
results are shown in Fig. 7C. At a concentration of 3 µM, treatment with C2 for 24 h
stimulated an approximately fourfold increase in ROS levels.
The Bcl-2 family of proteins have been described as key regulators of apoptosis
and consist of both anti-apoptotic proteins such as Bcl-2 and Bcl-xL and pro-apoptotic
proteins such as Bad, Bak, and Bax [51]. Many studies of metal complexes that
induce cell apoptosis have shown that these proteins play a critical role in
mitochondrially mediated apoptosis [52,53]. To investigate the effect of the Cu(II)
complex on the expression levels of these proteins, HepG2 cells were treated with
different concentrations of C2 for 24 h. It was found that the exposure of HepG2 cells
to C2 resulted in a decrease in the expression of the anti-apoptotic proteins Bcl-2 and
Bcl-xl and a slight increase in the expression of the pro-apoptotic protein Bax (Fig. 8).
As a result of these changes, the ratio of Bax to Bcl-2 increased, which led to the
mitochondrial release of apoptogenic factors. In addition, upregulation of the cleavage
of caspase-3 and the subsequent cleavage of PARP-1 were detected (Fig. 8). These
results provide confirmation of the mitochondrially based intrinsic apoptosis pathway.
4. Conclusions
In this study, two aroylhydrazone ligands and their copper(II) complexes were
synthesized and their anticancer activity was evaluated. Among all these compounds,
C2 displayed high activities against various cancer cell lines, including the cisplatin-
resistant lung cancer cell line A549cisR. C2 can bind to DNA and was capable of
efficiently converting supercoiled pBR322 DNA into an open circular form, which
most probably occurred via an oxidative cleavage mechanism. Further mechanistic
studies showed that C2 can effectively induce caspase-dependent apoptosis via
mitochondrially based pathways and the elevation of intracellular ROS levels. In
addition, C2 can induce the suppression of the expression levels of two key regulatory
genes, namely, CDK1 and cyclin B1, and inhibit cell growth by triggering cell cycle
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arrest at the G2/M phase. The results are helpful for the design and synthesis of new
Cu(II) complexes of aroylhydrazones as potent anticancer drugs.
Acknowledgements
The authors are grateful to the Guangxi Normal University for partial support of
this work. This work was supported by the Natural Science Foundation of China
(31460232) and Technology Division of Nantong (MS12016022).
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Figure captions
Scheme 1. Synthesis of copper(II) complexes.
Figure 1. (A) The local coordination environment of C1, hydrogen atoms are omitted
for clarity. (B) The local coordination environment of C2, hydrogen atoms are
omitted for clarity. (C) Optimized structures of ground (enol form and keto form) and
transition states (TS) in methanol for L1. (D) Optimized structures of ground (enol
form and keto form) and TS in CH3OH for L2.
Figure 2. UV-Vis spectra of L1/L2 (50 µM) with the addition of various
concentrations of Cu(II) in PBS buffer containing 10% (v/v) DMSO.
Figure 3. (A) Anti-migratory effect of C1 (3 µM) and C2 (3 µM) on HepG2 cells by
wound healing assay. (B) Anti-adhensive effect of C1 and C2 on HepG2 cells. (*p <
0.05, **p < 0.01, compared with control).
Figure 4. (A) Absorption spectra of C2 (5 µM) upon the titration of CT-DNA (0-15
µM). (B) Fluorescence quenching curves of ethidium bromide bound to DNA:C2.
[DNA] = 10 µM, [EB] = 8 µM, and C2 = 0-35 µM. (C) CD spectrum of CT‒DNA
with the addition of C2. (D) Effect of increasing amounts of EB and C2 on the
relative viscosity of CT-DNA. (E) Agarose gel electrophoresis patterns for the
cleavage of pBR322 DNA by C2 at pH 7.0 at 37 °C for 5 h. Lane 1: DNA alone;
Lanes 2−4: DNA with complexes at the concentrations of 12.5, 25, 50 µM,
respectively. (F) Agarose GE patterns for the cleavage of pBR322 DNA by C2 (50
µM) at pH 7.0 and 37 °C for 5 h, in the presence of DMSO (1 M, Lane 3), EtOH (1 M,
Lane 4), NaN3 (0.1 M, Lane 5), KI (0.1 M, Lane 6) and EDTA (0.1 M, Lane 7). Lane
1, DNA alone and Lane 2: DNA + C2.
Figure 5. (A) Cell cycle contributions resulting from treatment with C2 (0, 1.5 and 3
µM) for 24 h. (B) Representative Western blots of the effects of C2 on the protein
expression levels of CDK1 and cyclin B1. β−Actin was assessed as a loading control.
Figure 6. (A) Representative dot plots of annexin V and PI double staining on the
HepG2 cells after treatment with C2 (0, 1.5 and 3 µM) for 24 h. (B) Representative
images of AO/EB double stained HepG2 cells after treatment with C2 (0, 1.5 and 3
µM) for 24 h.
Figure 7. (A) Effects of C2 on mitochondrial membrane potential analyzed by JC-1
staining and flow cytometry. (B) Analysis of ROS levels by flow cytometry after
HepG2 cells were treated with vehicle or C2 at 1.5 and 3 µM for 24 h. (C)
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Quantification of the flow cytometric results in (B) showing the percentage of cells
with increased intracellular DCF oxidation compared to control cells. (*p < 0.05, **p
< 0.01, compared with control).
Figure 8. Western blot analysis of cleaved Caspase-3, cleaved PARP-1, Bcl-2, Bcl-xl,
and Bax in HepG2 cells treated with different concentrations of C2 for 24 h.
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Table 1 Crystal data for complexes C1 and C2.
Complex
Empirical formula
Molecular weight
Crystal system
Space group
a (Å)
C1
C2
C23H17CuN3O3 C24H19BrCuN4O2
446.94
monoclinic
P21/c
538.88
orthorhombic
P212121
7.7759(10)
16.670(2)
34.660(5)
90.00
11.787(3)
8.203(2)
20.478(6)
90.00
b (Å)
c (Å)
α (o)
β (o)
γ (o)
105.525(4)
90.00
90.00
90.00
T (K)
V (Å3)
296.15
1907.7(9)
4
296.15
4492.8(10)
8
Z
ρ
calc. (g·cm-3)
1.556
1.593
F(000)
916
2168
µ(Mo-Kα) (mm-1)
1.177
2.780
Data/restraint/parameters 3904/0/272
Goodness-of-fit on F2
1.055
9170/0/579
1.066
Final R1, wR2 [I > 2σ(I)] 0.0494, 0.0955 0.0673, 0.0935

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Table 2 Selected bond lengths [Å] and angles [º] in complexes C1 and C2.
C1
Cu1−N2
Cu1−O1
Cu1−O2
Cu1−N3
1.912(2)
O2−Cu1−N2 92.83(7)
1.9280(17) O2−Cu1−O1 173.88(7)
1.8876(16) N2−Cu1−N3 171.30(8)
2.011(2)
N2−Cu1−O1 81.60(7)
O1−Cu1−N3 92.07(8)
O2−Cu1−N3 93.74(8)
C3
Br1−Cu1
Cu1−N2
Cu1−O1
Cu1−N4
Cu1−N1
2.6356(8) N2−Cu1−O1 79.61(15)
1.948(4)
1.990(4)
1.975(4)
2.054(4)
N2−Cu1−N4 163.70(17)
N2−Cu1−N1 79.23(17)
O1−Cu1−Br1 108.75(12)
O1−Cu1−N1 153.41(16)
N2−Cu1−Br1 96.91(12) N4−Cu1−Br1 99.25(12)
N4−Cu1−O1 93.14(16) N4−Cu1−N1 102.91(18)
N1−Cu1−Br1 89.65(12)

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a
Table 3 Inhibition of human cancer cell lines growth (IC50 , µM) for C1 and C2.
compounds
cell growth inhibition, IC50 ± SD (µM)
HepG2
Bel-7402
MCF-7
A549
A549cisR
WI−38
L1
L2
C1
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
3.87 ± 0.24 5.23 ± 0.31 3.13 ± 0.18 4.69 ± 0.29
4.87 ±
0.27
5.33 ± 0.55
1.57 ± 0.08 1.86 ± 0.09 1.69 ± 0.18 2.86 ± 0.21
2.91 ±
0.17
4.19 ± 0.32
C2
CuBr2
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
> 40
Pyridine
Cisplatin
> 40
16.33 ±
1.58
14.27 ±
1.34
13.48 ±
1.12
23.17 ±
1.48
> 40
22.81 ±
1.16
a IC50 values are presented as the mean ± SD from three separated experiments.

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