Cadmium(II) complexes with a 4-acyl pyrazolone derivative and co-ligands: crystal structures and antitumor activity

Abstract

Three cadmium(II) complexes, [Cd(HL)₂(CH₃OH)₂]·(CH₃OH) (1), [Cd(HL)₂(bpy)]·(CH₃OH)₂ (2), and [Cd(HL)₂(phen)]·(CH₃OH)_1.5 (3), (where H₂L = N-(1-phenyl-3-methyl-4-(4-chlorobenzoyl)-5-pyrazolone)-2-thiophenecarboxylic acid hydrazide, bpy = 2,2′-bipyridine, phen = 1,10-phenanthroline) have been synthesized and characterized. Single crystal X-ray diffraction analyses indicated that complexes 1–3 exhibit mononuclear octahedral geometry. The spectrophotometric analyses showed that these complexes could bind with herring sperm DNA and bovine serum albumin (BSA). The intrinsic binding constants to HS-DNA were 2.46 × 10⁴ M⁻¹, 2.64 × 10⁴ M⁻¹, and 4.41 × 10⁴ M⁻¹, and the quenching constants to BSA were 1.23 × 10⁶ M⁻¹, 1.85 × 10⁶ M⁻¹, and 2.17 × 10⁶ M⁻¹ for the complexes 1–3, respectively. The complexes have higher cytotoxic activities against HeLa and Eca-109 tumor cells than that of the ligand and cisplatin. Complex 3 shows the highest cytotoxicity for both HeLa and Eca-109, and the IC₅₀ values are 3.35 ± 0.2 μg mL⁻¹ and 7.41 ± 0.07 μg mL⁻¹, respectively. When compared with our previous work, IC₅₀ value of the reported complex CdC₂₀H₁₈N₄O₃ to Eca-109 cells is 14.18 μg mL⁻¹. We further found that complex 3 inhibits the growth of HeLa cells by inducing apoptosis and arresting the cell cycle during the G0/G1 phase. These results suggest that complex 3 is a potential antitumor drug.

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1. Introduction

Although the strategies for the prevention and treatment of cancer have been greatly improved, the disease is still a major problem for human beings. Chemotherapy is the first-line treatment for various kinds of cancers. It is well known that the platinum metal complexes including cisplatin, carboplatin, and oxaliplatin have been widely used in the clinical treatment of cancer. However, these drugs have severe side effects, poor selectivity, cross resistance, and other defects.^1,2 Therefore, it is urgently required to explore new chemotherapy drugs with higher efficacy and minimal side effects.³ Great attention has been paid to the transition metal complexes of Schiff bases due to their advantages, including stability, easy synthesis, plasticity, and various bioactivities such as antibacterial,^4,5 antiviral,⁶ and anticancer activity.^7,8

In recent years, transition metal complexes of pyrazolone derivatives, especially 4-acyl pyrazolone Schiff base derivatives, have been extensively studied due to their broad spectrum of biological activities, such as antibacterial, antiviral, antioxidative, and antitumor activities,^9–15 which represent these as promising anticancer drug candidates. In addition, 4-acyl pyrazolone Schiff base derivatives have potential to form different types of complexes due to multiple coordination atoms and diversiform coordination modes.^16,17 X. H. Wang reported that the copper complex [CuL_a(EtOH)] (H₂L_a = N-(1-phenyl-3-methyl-4-propenylidene-5-pyrazolone)-salicylidene hydrazide) was able to induce apoptosis in the KB cells and KBv200 cells via a reactive oxygen species (ROS)-independent mitochondrial pathway.¹⁸ F. P. Fanizzi et al. synthesized and characterized new platinum(II) complexes of acylpyrazolone ligands. Note that, the water-soluble complex trans-[PtCl₂(DMSO)(HQ^Ph)] (HQ^Ph = 1-phenyl-3-methyl-4-benzoyl-5-pyrazolone) showed higher cytotoxicity towards the HeLa cells.¹⁹ Our recent study showed that the manganese complex [Mn(HL_b)(L_b)]·(CH₃CN)_1.5·H₂O (H₂L_b = N-(1-phenyl-3-methyl-4-benzoyl-5-pyrazolone)-2-thiophenecarboxylic acid hydrazide) inhibited the growth of HeLa cells by inducing apoptosis and arresting the cell cycle during the S phase.²⁰ R. Pettinari synthesized ruthenium(II) arene complexes containing a pyrazolone-based β-ketoamine ligand and investigated the antitumor activity of the ligand and ruthenium(II) complexes on the human ovarian cancer cell line A2780 and A2780cisR cells. The studies indicated that most of the complexes inhibited cell proliferation and their cell toxicity was better than that of the ligand and cisplatin.²¹

Accumulated evidence has shown that cadmium(II) can affect cell proliferation, differentiation, and apoptosis.^22–25 K. Sinha reported that cadmium(II) induces apoptosis in human cells depending on the exposure concentration and conditions. This study also explained the molecular mechanisms of cadmium(II) induced oxidative stress and apoptosis through a mitochondrial-dependent pathway in the SK-RC-45 cells.²⁶ Our study showed that the cadmium(II) complex [CdL_a]_n of a pyrazolone-based derivative greatly inhibits the growth of human esophageal cancer (Eca-109) cells. In addition, complex [CdL_a]_n displays a much lower inhibitory effect on the normal human gastric epithelial (GES) cells. The IC₅₀ value for GES cells was 142.6 μM, which is about five times higher than that for Eca-109 cells. The studies also suggested that [CdL_a]_n may induce apoptosis through the generation of ROS and a caspase-dependent mitochondria-mediated pathway.²⁷ These results motivated us to further investigate the antitumor activities of cadmium(II) complexes comprising 4-acyl pyrazolone Schiff base derivatives. Herein, a new Schiff base ligand (H₂L) was synthesized through the interaction of 4-acyl pyrazolone with 2-thiophenecarboxylic acid hydrazide. Three mixed ligand cadmium(II) complexes [Cd(HL)₂(CH₃OH)₂]·(CH₃OH) (1), [Cd(HL)₂(bpy)]·(CH₃OH)₂ (2), and [Cd(HL)₂(phen)]·(CH₃OH)_1.5 (3) were prepared. The interactions of H₂L and the complexes 1–3 with HS-DNA and BSA were investigated using spectrophotometric methods. Their antitumor activities in Eca-109 and HeLa cells were determined using an MTT assay (MTT = 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide). The effect of complex 3 on the cell cycle distribution and apoptosis of HeLa cells was analyzed by flow cytometry and Hoechst 33258 staining.

2. Experimental section

2.1 Materials and methods

1-Phenyl-3-methyl-4-(4-chlorobenzoyl)-5-pyrazolone (PM4ClBP) was synthesized according to a reported procedure.²⁸ 2-Thiophenecarboxylic acid hydrazide (TAH), 2,2′-bipyridine, 1,10-phenanthroline, Cd(CH₃COO)₂·2H₂O, glacial acetic acid, ethanol, and acetonitrile were all analytical grade and used without further purification. Dioxane was used after purification.

BSA, ethidium bromide (EB), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), dimethyl sulfoxide (DMSO), and Hoechst 33258 were obtained from Sigma-Aldrich. Herring sperm DNA (HS-DNA) was purchased from Beijing Solarbio Science & Technology Co., Ltd. An Annexin V-FITC Apoptosis Detection Kit was obtained from Calbiochem Co. Other biological routine laboratory reagents were obtained from the commercial sources and were of analytical or HPLC grade.

IR spectra were obtained using a Bruker Vertex-70 spectrophotometer within 400–4000 cm⁻¹ with the samples prepared as pellets using KBr. The UV spectra were obtained using a Hitachi U-3900H spectrophotometer. The fluorescence behavior of the ligands and complexes were studied using a Hitachi F-4500 fluorescence spectrophotometer for DNA and Horiba Jobin-Yvon Fluorolog-3 fluorescence spectrophotometer for BSA with a Xe arc lamp as the light source at room temperature.

All experiments involving the interaction of the complexes with HS-DNA and BSA were carried out in Tris–HCl buffer (pH = 7.2). A solution of HS-DNA in Tris–HCl buffer gave the ratio of 1.8–1.9 : 1 for the UV absorbance at 260 and 280 nm (A₂₆₀/A₂₈₀), indicating that the DNA was sufficiently free from protein contamination.²⁹ The HS-DNA concentration was determined by employing an extinction coefficient of 6600 M⁻¹ cm⁻¹ at 260 nm.^30–32

2.2 Synthesis

2.2.1 Preparation of the H₂L ligand. PM4ClBP (5 mmol, 1.564 g) was dissolved in a 40 mL of hot anhydrous ethanol to which an ethanol solution of TAH (5 mmol, 0.711 g) was slowly added with constant stirring. After adding a few drops of pure acetic acid as catalyst, the mixture was refluxed for 4 h. After cooling down to room temperature, the yellow product was filtered and recrystallized using ethanol. Anal. calc. for C₂₂H₁₇ClN₄O₂S (F.W.: 436.91): C, 60.47; H, 3.89; N, 12.89. Found: C, 60.32; H, 3.95; N, 13.05%. Selected IR bands (cm⁻¹): 2979 (w), 1594 (s), 1499 (s), 1368 (m), 1242 (w), 1030 (m), 753 (m). ¹H NMR (CHCl₃-d₆, 400 MHz) δ ppm: 15.93 (s, 1H), 8.43 (s, 1H), 8.02 (d, J = 8 Hz, 2H), 7.86 (d, J = 4 Hz, 1H), 7.58 (m, 5H), 7.46 (m, 2H), 7.24 (m, 1H), 7.16 (m, 1H), 1.59 (s, 3H).

2.2.2 Preparation of the complexes.
[Cd(HL)₂(CH₃OH)₂]·(CH₃OH) (1). The ligand H₂L (0.0306 g, 0.07 mmol) was dissolved in a 8 mL of methanol and 2 mL of DMF. Then, a methanol solution of Cd(CH₃COO)₂·2H₂O (0.0187 g, 0.07 mmol) was added dropwise with constant stirring. The mixture was stirred for 2 h at room temperature and then filtered. Yellow single crystals suitable for X-ray diffraction analysis were obtained by slow evaporation of the filtrate. Anal. calc. for C₄₇H₄₄CdCl₂N₈O₇S₂ (F.W.: 1080.32): C, 52.21; H, 4.07; N, 10.27. Found: C, 52.39; H, 3.85; N, 10.18%. Selected IR bands (cm⁻¹): 2974 (w), 1598 (s), 1415 (s), 1384 (m), 1245 (w), 1015 (m), 755 (m). ¹H NMR (CHCl₃-d₆, 400 MHz) δ ppm: 8.72 (s, 2H), 8.09 (s, 2H), 7.85 (s, 2H), 7.65 (d, J = 8 Hz, 4H), 7.50 (d, J = 4 Hz, 2H), 7.43–7.40 (m, 6H), 7.24–7.22 (m, 6H), 7.08–7.03 (m, 2H), 1.60 (s, 6H), 1.44 (s, 6H), 1.00 (s, 2H).
[Cd(HL)₂(bpy)]·(CH₃OH)₂ (2). A methanolic solution of Cd(CH₃COO)₂·2H₂O (0.0187 g, 0.07 mmol) was slowly added to a solution of H₂L ligand (0.0306 g, 0.07 mmol) in a 10 mL of MeOH/CH₂Cl₂ (7/3, v/v) and the mixture was stirred for 30 min. Then, a methanolic solution of 2,2′-bipyridine (0.0109 g, 0.07 mmol) was added dropwise. The resulting mixture was stirred for 2 h and filtered. Yellow single crystals suitable for X-ray analysis were obtained by slow evaporation of the filtrate at room temperature. Anal. calc. for C₅₆H₄₈CdCl₂N₁₀O₆S₂ (F.W.: 1204.46): C, 55.79; H, 3.99; N, 11.62. Found: C, 55.63; H, 3.93; N, 11.73%. Selected IR bands (cm⁻¹): 2922 (w), 1598 (s), 1418 (s), 1375 (m), 1246 (w), 1014 (m), 759 (m). ¹H NMR (CHCl₃-d₆, 400 MHz) δ ppm: 18.21 (t, J = 16 Hz, 2H), 8.23 (d, J = 4 Hz, 2H), 8.00 (m, 8H), 7.91 (d, J = 4 Hz, 2H), 7.65 (d, J = 4 Hz, 2H), 7.46 (m, 2H), 7.39 (m, 4H), 7.25 (m, 2H), 7.15 (m, 2H), 7.05 (s, 4H), 6.77 (d, J = 8 Hz, 4H), 1.17 (s, 6H).
[Cd(HL)₂(phen)]₂·(CH₃OH)₃ (3). The complex 3 was synthesized via the same procedure that was used to prepare complex 2 with an exception of using 1,10-phenanthroline instead of 2,2′-bipyridine. Yellow crystals of 3 were obtained. Anal. calc. for C₁₁₅H₉₂Cd₂Cl₄N₂₀O₁₁S₄ (F.W.: 2424.93): C, 56.91; H, 3.79; N, 11.55. Found: C, 57.08; H, 3.65; N, 11.61%. Selected IR bands (cm⁻¹): 2972 (w), 1591 (s), 1420 (s), 1376 (m), 1248 (w), 1013 (m), 757 (m). ¹H NMR (CHCl₃-d₆, 400 MHz) δ ppm: 18.18 (t, J = 16 Hz, 2H), 8.46 (d, J = 8 Hz, 2H), 8.25 (d, J = 4 Hz, 2H), 8.21 (m, 2H), 7.99 (d, J = 8 Hz, 4H), 7.95 (s, 2H), 7.76 (m, 2H), 7.67 (d, J = 4 Hz, 2H), 7.38 (t, J = 16 Hz, 4H), 7.27 (m, 2H), 7.14 (t, J = 16 Hz, 2H), 6.90 (s, 4H), 6.47 (d, J = 8 Hz, 4H), 1.09 (s, 6H).

2.3 X-ray crystallography

X-ray single crystal diffraction data for the H₂L ligand and complexes 1–3 were obtained using a Bruker Apex-II CCD area-detector diffractometer equipped with graphite monochromated Mo Kα radiation (λ = 0.71073 Å) at 120 K. All the structures were solved by direct methods using SHELXS-97 and refined by full-matrix least-squares methods on F² using the SHELXL-97 program package. Non-H atoms were anisotropically refined using all the reflections with I > 2σ(I). The details of the crystal parameters, data collection, and refinements for H₂L and complexes 1–3 are summarized in Table 1. Selected bond lengths and bond angles are listed in Table S1. CCDC 1506204–1506207 contains the supplementary crystallographic data for this study.†

Table 1 Crystal data and structural refinement for the H₂L ligand and complexes 1–3

Compound	H2L	1	2	3
a Maximum and minimum residual electron density.
Formula	C22H17ClN4O2S	C47H44CdCl2N8O7S2	C56H48CdCl2N10O6S2	C115H92Cd2Cl4N20O11S4
F.W.	436.91	1080.32	1204.46	2424.93
Crystal system	Monoclinic	Monoclinic	Monoclinic	Triclinic
Space group	P21/c	P21/c	P21/c	P1
a (Å)	11.1799(11)	28.215(2)	12.1579(14)	10.309(3)
b (Å)	22.336(2)	9.7235(8)	20.045(2)	12.277(3)
c (Å)	8.8940(9)	17.3731(14)	21.961(2)	21.891(6)
α (°)	90	90	90	85.849(3)
β (°)	112.803(2)	100.0690(10)	94.411(2)	88.224(3)
γ (°)	90	90	90	87.932(3)
V (Å^3)	2047.3(4)	4692.9(6)	5336.2(10)	2760.4(13)
Z	4	4	4	1
D calc (g cm^−3)	1.417	1.529	1.499	1.459
F(000)	904	2208	2464	1238
Reflections collected	10 276	11 025	24 632	21 599
Uniq. reflect. (Rint)	3638 (0.0505)	3981 (0.0307)	8926 (0.0796)	9736 (0.0324)
S	1.009	1.085	1.096	1.099
R indices [I > 2σ(I)]	R 1 = 0.0340, ωR2 = 0.0917	R 1 = 0.0656, ωR2 = 0.2104	R 1 = 0.0668, ωR2 = 0.1613	R 1 = 0.0649, ωR2 = 0.1842
R indices (all data)	R 1 = 0.0368, ωR2 = 0.0942	R 1 = 0.0709, ωR2 = 0.2213	R 1 = 0.1099, ωR2 = 0.1892	R 1 = 0.0764, ωR2 = 0.1946
Δρ^a (e Å^−3)	0.324, −0.320	3.063, −2.244	1.001, −1.601	2.278, −0.919

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2.4 DNA-binding studies

2.4.1 UV absorption titration. An electronic absorption titration experiment was performed with a fixed concentration of the ligand and complexes (50 μM) and the concentration of DNA (0–58.7 μM) was gradually increased. After each addition, the DNA and complex mixtures were incubated at room temperature for 5 min and scanned from 240 to 550 nm. The data were then fitted to the following equation to obtain the intrinsic binding constant K_b:³³

[DNA]/[ε_a − ε_f] = [DNA]/[ε_b − ε_f] + 1/K_b[ε_b − ε_f] (1)

where [DNA] is the concentration of DNA in base pairs, ε_a is the extinction coefficient of the complex at a given DNA concentration, ε_f is the extinction coefficient of the complex in free solution, and ε_b is the extinction coefficient of the complex when fully bound to DNA. A plot of [DNA]/[ε_a − ε_f] versus [DNA] gave a slope and intercept equal to 1/[ε_a − ε_f] and (1/K_b)[ε_b − ε_f], respectively. The intrinsic binding constant K_b was calculated from the ratio of the slope to the intercept.

2.4.2 Fluorescence spectroscopy studies. Competitive studies of the complexes with EB were investigated using fluorescence spectroscopy. The experiments were conducted by adding a solution of the complex step by step (0–1.6 μM) to the Tris–HCl buffer containing DNA-bound EB. The changes in the fluorescence intensity were obtained. The excitation wavelength of the ligand and complexes was 530 nm and the emission wavelength was 590 nm.

Further support for the binding mode was given by an emission quenching experiment. Quenching data were analyzed on the basis of the Stern–Volmer equation, which can be used to determine the fluorescence quenching mechanism:³⁴

F₀/F = 1 + K_q[Q] (2)

where, F₀ is the emission intensity in the absence of the quencher, F is the emission intensity in the presence of the quencher, K_q is the quenching constant, and [Q] is the quencher concentration. The K_q value was obtained as the slope of the plot of F₀/F versus [Q].

2.5 BSA binding studies

The BSA binding studies were performed at an excitation wavelength of 280 nm and the corresponding emission at 346 nm. Synchronous fluorescence spectra were also obtained with the same concentrations of BSA, and the complexes corresponding to the Δλ values (the difference between the excitation and emission wavelengths of BSA) 15 and 60 nm, respectively.

The fluorescence quenching was analyzed according to the Stern–Volmer quenching equation:³⁵

F₀/F = 1 + K_sv[Q] = 1 + κ_qτ₀[Q] (3)

where F₀ and F are the fluorescence intensities of the fluorophore in the absence and presence of the quencher, respectively, K_sv is the dynamic quenching constant, and [Q] is the concentration of the quencher. κ_q is the biomolecular quenching constant and τ₀ is the average lifetime of protein in the absence of quencher and its value was 10⁻⁸ s.

To further elucidate the quenching mechanism, fluorescence quenching data were analyzed using the Scatchard equation:^36,37

log[(F₀ − F)/F] = log K_bin + n log[Q] (4)

where K_bin is the apparent binding constant and n is the number of binding sites per albumin, which is calculated from the intercept and slope of the plot of log[(F₀ − F)/F] versus log[Q].

2.6 Cytotoxicity evaluation

The effects of the cadmium(II) complexes on the Eca-109 and HeLa cell proliferation were investigated using the MTT assays.³⁸ Cells were maintained in a humidified atmosphere containing 5% CO₂ at 37 °C in RPMI 1640 medium supplemented with 100 units per mL penicillin, 100 μg mL⁻¹ streptomycin, and 10% fetal bovine serum. Briefly, cell lines of human esophageal cancer Eca-109 cells and cervical cancer HeLa cells with a density 2 × 10⁴ cells per well were precultured into the 96-well microtiter plates for 24 h.

Cells were treated with different concentrations of the cadmium(II) complexes and the H₂L ligand. After 24 h, MTT solution (0.5 mg mL⁻¹ PBS) was added to each well. After 3–4 h of incubation, DMSO was added to solubilize the MTT formazan. The optical density of each well was then measured on a microplate spectrophotometer at 570 nm. All experiments were performed in triplicate and the percentage of cell viability was calculated according to the following equation:

Cell growth inhibition (%) = [1 − A₅₇₀ (sample)/A₅₇₀ (control)] × 100% (5)

where A₅₇₀ (sample) refers to the wells treated with the complexes and A₅₇₀ (control) refers to the wells treated with medium containing 10% fetal bovine serum (FBS) only. The half-maximal inhibitory concentration (IC₅₀) was determined using GraphPad Prism 5.0.

2.7 Apoptosis evaluation

2.7.1 Hoechst 33258 staining. To examine whether complex 3 induced apoptosis in Hela cells, 5 × 10⁴ cells were incubated with complex 3 at 0, 2, 4, 8, and 16 μg mL⁻¹ for 6 h, respectively. Subsequently, the HeLa cells were washed twice with ice-cold PBS and incubated with 1 mL of Hoechst 33258 (0.5 μg mL⁻¹) for 10 min at 37 °C in the dark. After washing twice with PBS, the cells were visualized using fluorescence microscopy (Leica Dmirb, Germany).

2.7.2 Cell cycle assay. 5 × 10⁵ HeLa cells were treated with different concentrations (0, 1, 2, 4, and 8 μg mL⁻¹) of complex 3 at 37 °C for 24 h. The cells were harvested in cold PBS, centrifuged, resuspended, and fixed in 70% cold ethanol 30 min at 4 °C. After further washing steps with cold PBS, the cells were stained with PI (300 μL) in the dark at room temperature for 10 min. The distribution of the cell cycle was measured using a FACStar Plus flow cytometer, and the data analysis was carried out using ModFit LT software.

2.7.3 Measurement of apoptosis by Annexin V-FITC/PI analysis. 8 × 10⁵ HeLa cells treated with various concentrations (0, 2, 4, 8, and 16 μg mL⁻¹) of complex 3 for 24 h were harvested and washed with PBS. Thereafter, the cells were resuspended in the binding buffer (100 μL). Annexin V-FITC (3 μL) was added to the cells and incubated for 10 min in the dark at room temperature. The samples were incubated for 5 min by PI (3 μL) before adding an additional binding buffer (100 μL) and were examined using an Annexin V-FITC Apoptosis Detection Kit.

2.8 Statistical analysis

All experiments were repeated 3 times. The data were processed by test and the one-way analysis of variance (ANOVA) using GraphPad Prism 5.0. The results were expressed as the mean ± standard deviation (S.D.) and considered to be significant when p < 0.05.

3. Results and discussion

3.1 Description of the crystal structure

3.1.1 The structure of the H₂L ligand. A molecular view of the H₂L ligand using the atom-numbering scheme is shown in Fig. 1. X-ray crystallographic analysis revealed that the H₂L ligand crystallizes in the monoclinic space group P2₁/c. The bond lengths of O1–C7 and O2–C18 are 1.2619(19) and 1.2398(18) Å, respectively, which are shorter than 1.43 Å for the C–O single bond and slightly longer than 1.22 Å for the C=O double bond. Similarly, the bond length of C8–C11 is 1.406(2) Å, which is indicative of the conjugated C=C double bond character. This information indicates that the H₂L ligand adopted a ketone structure rather than the Schiff base form.

Fig. 1 The crystal structure of the H₂L ligand and its atom-numbering scheme.

3.1.2 The structure of complex 1. The crystal structure of complex 1 is shown in Fig. 2 with its atom-numbering scheme. Complex 1 features a mononuclear structure and its asymmetric unit contains one Cd(II) center, two ligand anions HL⁻, two coordinated methanol molecules, and one lattice methanol molecule. The Cd(II) atom is located at the inversion center and hexacoordinated by O2, N3, O2A, and N3A atoms from a pair of ligand anions HL⁻ and two O atoms (O3 and O3A) from the coordinated methanol molecules to form an octahedron geometry. Moreover, O2, O3, N3, and O2A atoms consist of an equatorial plane with a least square plane deviation of 0.1267 Å, and the Cd(II) atom strays from the equatorial plane by 0.1118 Å. The axial positions are occupied by O3A and N3A atoms. The bond lengths of Cd1–O2, Cd1–O3, and Cd1–N3 are 2.315(3), 2.313(3), and 2.340(4) Å, respectively. The bond angles of O2–Cd1–O2A and O3–Cd1–N3 are 161.75(17)° and 153.34(12)°, respectively, which shows a deviation from the theoretical value of 180°. Therefore, the coordination geometry of the Cd(II) center can be described as a highly distorted octahedron.

Fig. 2 The crystal structure of complex 1 with its atom-numbering scheme. The phenyl groups at 1-position of the pyrazolone ring and 4-chlorophenyl groups are omitted for clarity.

In addition, the ligand acts as a negative monovalent bidentate chelating agent that bonds with Cd(II) atom. Note that only azomethine N3 and enolic O2 atoms coordinate with the Cd(II) atom, whereas O1 atom of the pyrazolone ring is free of coordination. The unexpected monovalent bidentate coordination mode of the Schiff base ligand is very rare in the reported complexes of pyrazolone derivatives.

3.1.3 The structure of complex 2. X-ray crystallographic analysis revealed that complex 2 crystallizes in the monoclinic space group of P2₁/c and the asymmetric unit contains one Cd(II) center, two crystallographically independent ligand anions HL⁻, one 2,2′-bipyridine, and two lattice methanol molecules. The coordination environment around the Cd(II) center in complex 2 is presented in Fig. 3 with its atom-numbering scheme. The Cd1(II) atom also adopts a distorted octahedron geometry and is coordinated by O2 and N3 atoms from one ligand anion, O4 and N7 atoms from another ligand anion, and two N atoms (N9 and N10) from the 2,2′-bipyridine molecule. The basal plane is made up of O2, N3, N7, and N9 atoms and Cd(II) atom is displaced by 0.0932 Å from the basal plane. The Cd1–O and Cd1–N bond distances are in the range from 2.299(5) to 2.370(6) Å. The bond angles of O4–Cd1–N10, O2–Cd1–N9, and N3–Cd1–N7 are 158.19(18)°, 170.20(17)°, and 155.56(19)°, respectively, which are seriously deviated from the ideal value. All these observations indicate that the geometry around Cd(II) atom in 2 is a seriously distorted octahedron. The ligand anion HL⁻, also acts as a bidentate chelating agent, coordinates to Cd(II) atom.

Fig. 3 The crystal structure of complex 2 with the coordination environment around the Cd(II) center. The phenyl groups at 1-position of the pyrazolone ring and the 4-chlorophenyl groups are omitted for clarity.

3.1.4 The structure of complex 3. The molecular structure of complex 3 is displayed in Fig. 4. The structure of complex 3 is similar to complex 2 except for the replacement of 2,2′-bipyridine with 1,10-phenanthroline. X-ray diffraction analysis revealed that complex 3 crystallizes in the triclinic space group of P1. The distorted octahedral coordination sphere of Cd1(II) atom is completed by four donor atoms (O2, O4, N3, N7) from two ligand anions HL⁻ and N9, N10 atoms from a single 1,10-phenanthroline. The O2, O4, N9, and N10 atoms comprise the equatorial plane with a least square plane deviation of 0.1784 Å, and the Cd(II) atom almost lies in the equatorial plane with a deviation of 0.0736 Å. N3 and N7 atoms occupy the remaining apical coordination sites. The Cd1–O and Cd1–N bond lengths are in the normal range and the bond angles of O2–Cd1–N10, O4–Cd1–N9, and N3–Cd1–N7 are 160.79(16)°, 168.19(17)°, and 158.92(16)°, respectively. Thus, the coordination geometry of the Cd(II) center in complex 3 is seriously distorted from a regular octahedron.

Fig. 4 The crystal structure of complex 3 with its atom-numbering scheme. The phenyl groups at 1-position of the pyrazolone ring and the 4-chlorophenyl groups are omitted for clarity.

3.2 DNA-binding studies

3.2.1 UV-visible absorption spectroscopy. The UV-visible absorption spectra of the ligand and complexes in the absence and presence of HS-DNA are shown in Fig. 5. The addition of DNA to the complexes caused significant changes in their absorption spectra accompanied by a weak red shift, which indicates that the metal complexes partially bind with DNA by insertion.^39–41 To quantitatively compare the binding strength of the ligand and complexes with HS-DNA, their intrinsic binding constants (K_b) were obtained from eqn (1), and the values of K_b were 2.31 × 10⁴ M⁻¹, 2.46 × 10⁴ M⁻¹, 2.64 × 10⁴ M⁻¹, and 4.41 × 10⁴ M⁻¹ corresponding to the ligand and complexes 1–3, respectively. The values of K_b revealed that the complex 3 has a higher binding affinity with HS-DNA as compared to that of the other complexes and the order of binding affinity was 3 > 2 > 1 > H₂L. The coordinated 1,10-phenanthroline molecule in complex 3 with a larger aromatic ring surface makes a deeper insertion into the base pairs of DNA as compared to the coordinated 2,2′-bipyridine molecule in complex 2, which might cause the higher DNA binding affinity of complex 3.⁴²

Fig. 5 The UV-visible absorption spectra of the H₂L ligand and complexes 1–3 (50 μM) in the presence of HS-DNA (0–58.7 μM). The arrow shows the absorbance changes upon increasing the DNA concentration. Inset shows the plots of [DNA]/[ε_a − ε_f] vs. [DNA].

3.2.2 Fluorescence spectroscopy studies. We further investigated the interaction of these complexes with DNA using a competitive binding assay of the EB-DNA system. EB is an intercalator that induces a significant increase in the fluorescence emission when bound to DNA, which can be quenched by the addition of another DNA binding molecule by either replacing the EB and/or by accepting the excited state electron of EB through an electron transfer mechanism.^43–46 As shown in Fig. S1,† the fluorescence intensity of the EB-DNA binding system at 599 nm decreased in the presence of the ligand and complexes, which indicates that the ligand and complexes competitively bind to DNA and displace EB. The quenching constant K_q obtained from the plot of [Q] versus F₀/F (shown as insets in Fig. S1†) was used to evaluate the quenching efficiency of the ligand and complexes according to the linear Stern–Volmer eqn (2). The K_q values obtained for the ligand and complexes 1–3 were 6.59 × 10⁴ M⁻¹, 7.14 × 10⁴ M⁻¹, 7.20 × 10⁴ M⁻¹, and 7.82 × 10⁴ M⁻¹, respectively. Similar with the abovementioned results, complex 3 has a higher binding affinity with DNA followed by 2, 1, and H₂L. Taken together, these results indicate that the H₂L ligand and complexes may bind to HS-DNA via an intercalative mode.

3.3 BSA binding studies

Fluorescence quenching measurements have been widely used to study the interaction of metal complexes or small molecules with proteins.^47,48 Increasing concentrations of the ligand and complexes were added into the BSA solution (5.1 μM) and the fluorescence spectra of the samples were obtained in the wavelength range of 300–540 nm upon excitation at 280 nm (Fig. 6). We observed that the H₂L ligand and complexes significantly decreased the fluorescence intensity of BSA at 346 nm up to 74.74%, 83.82%, 88.55%, and 89.50% of the initial fluorescence intensity of BSA for the H₂L ligand and complexes 1–3, respectively, with a slight hypsochromic shift. Commonly, fluorescence quenching can be described by the Stern–Volmer eqn (3). The K_sv value was obtained from the slope of the plot of F₀/F versus [Q] (shown as insets in Fig. 7). The K_sv values were 0.71 × 10⁶ M⁻¹, 1.23 × 10⁶ M⁻¹, 1.85 × 10⁶ M⁻¹, and 2.17 × 10⁶ M⁻¹ corresponding to the H₂L ligand and complexes 1–3, respectively, and suggest that complex 3 has a higher binding affinity with BSA followed by 2, 1, and H₂L.

Fig. 6 The emission spectra of BSA in the absence and presence of an increasing amount of the H₂L ligand and complexes 1–3. λ_ex = 280 nm, λ_em = 347 nm, [BSA] = 5.1 μM; [complex] = 0–4.55 μM from top to bottom, respectively. The arrow shows the emission intensities changes on increasing the complex concentration. Inset shows the Stern–Volmer plots of F₀/F vs. [Q].

Fig. 7 The plot of log[(F₀ − F)/F] vs. log[Q].

Fluorescence quenching mechanisms are usually classified as either static or dynamic quenching. Static quenching refers to the combination of a quencher with the fluorophore to form complexes. Dynamic quenching refers to the contact of the fluorophore and quencher during the transient existence of the excited state.⁴⁹ As shown in Table 3, the κ_q values (10¹³ M⁻¹ s⁻¹) of the H₂L ligand and complexes were 0.71 × 10¹⁴, 1.23 × 10¹⁴, 1.85 × 10¹⁴, and 2.17 × 10¹⁴, respectively, which are much higher than the maximum scatter collision-quenching constant of the diverse kinds of quenchers for biopolymers fluorescence (2 × 10¹⁰ M⁻¹ s⁻¹) and indicate the existence of a static quenching mechanism.⁵⁰

Table 2 The quenching constant, binding constant, and number of binding sites obtained for the interactions of the H₂L ligand and complexes 1–3 with BSA

Compounds	K sv (M^−1)	κ q (M^−1 s^−1)	n	K bin
H2L	0.71 × 10^6	0.71 × 10^14	1.42	1.15 × 10^8
1	1.23 × 10^6	1.23 × 10^14	1.44	2.63 × 10^8
2	1.85 × 10^6	1.85 × 10^14	1.69	8.13 × 10^9
3	2.17 × 10^6	2.17 × 10^14	1.71	1.02 × 10^10

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Table 3 The cytotoxic activity of the H₂L ligand and complexes 1–3

	IC50 values (μg mL^−1)
Compound	HeLa	Eca-109
H2L	20.34 ± 1.1	—
1	10.21 ± 0.1	19.29 ± 0.6
2	8.11 ± 0.2	17.26 ± 0.5
3	3.35 ± 0.2	7.41 ± 0.07
DDP	29.26 ± 1.1	72.72 ± 0.3

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For the static quenching interaction, the apparent binding constant K_bin and the number of binding sites (n) can be calculated according to the Scatchard eqn (4) based on the slope and the intercept of the double logarithm regression curve of log[(F₀ − F)/F] versus log[Q] (Fig. 7). As shown in Table 2, the K_bin values of the H₂L ligand and complexes 1–3 are 1.15 × 10⁸ M⁻¹, 2.63 × 10⁸ M⁻¹, 8.31 × 10⁹ M⁻¹, and 1.02 × 10¹⁰ M⁻¹, respectively. The n values of the H₂L ligand and complexes 1–3 are 1.42, 1.44, 1.69, and 1.71, respectively. The results suggest that the complexes have a stronger interaction with BSA as compared to the H₂L ligand. Similarly, complex 3 has a higher binding affinity with BSA than that of the other complexes.

3.3.1 Synchronous fluorescence spectroscopy studies of BSA. BSA contains tryptophan, tyrosine, and phenylalanine, which are natural amino acids with fluorescing groups. Synchronization fluorescence spectrum was obtained to distinguish tryptophan (the wavelength interval: Δλ = 60 nm) and tyrosine (Δλ = 15 nm) residues in proteins and to evaluate the conformational changes of BSA.^51,52 The effects of the H₂L ligand and complexes 1–3 on the BSA synchronous fluorescence spectra with Δλ = 15 nm and Δλ = 60 nm are shown in Fig. S2 and S3,† respectively. We found that the fluorescence intensity of the emission corresponding to both tyrosine and tryptophan were dramatically decreased by the H₂L ligand and complexes 1–3 in a dose-dependent manner. In the synchronous fluorescence spectra of BSA at Δλ = 15 nm, the fluorescence intensity of BSA at 301 nm was decreased up to 53.43%, 61.95%, 73.84%, and 78.21% of the initial fluorescence intensity of BSA for the H₂L ligand and complexes 1–3, respectively (Fig. S2†). At the same time, in the synchronous fluorescence spectra of BSA at Δλ = 60, the fluorescence intensity at 347 nm was decreased up to 74.87%, 83.53%, 88.40%, and 90.28% of the initial fluorescence intensity of BSA for the ligand and complexes 1–3, respectively (Fig. S3†). The results further demonstrated that complex 3 has a higher binding affinity with BSA.

Fig. 8 The inhibition activities of complex 1, 2, and 3 towards HeLa and Eca-109 cells. The HeLa (A) and Eca-109 (B) cells were treated with various concentrations of complexes 1–3 for 24 h, and then the cell viability was measured using an MTT assay. (C) The HeLa and Eca-109 cells were treated with the number of moles of Cd(CH₃COO)₂·2H₂O similar to that of complex 3 for 24 h, and then the cell viability was determined using an MTT assay. Each data point is the mean ± standard error obtained from three independent experiments. Significant differences from the untreated control were indicated by p: *p < 0.05, **p < 0.01, and ***p < 0.001. (D) Images of the HeLa cells after treatment with the same number of moles of Cd(CH₃COO)₂·2H₂O (upper panel) or complex 3 (lower panel) for 24 h.

Fig. 9 Morphological changes in the HeLa cells observed under a fluorescent microscope after Hoechst 33258 staining.

Fig. 10 The effect of the complex 3-induced apoptosis on the Hela cells for 24 h. HeLa cells were treated with different concentrations of complex 3. The harvested apoptotic cells were stained with Annexin V-FITC/PI and analyzed using flow cytometry. The horizontal axis represents the Annexin V-FITC intensity and the vertical axis shows the PI staining. *p < 0.05, **p < 0.01, and ***p < 0.001 vs. control cells.

3.4 Cytotoxicity evaluation

HeLa and Eca-109 cells were treated with different concentrations of the complexes and the cytotoxicity of the complexes was determined using an MTT assay. To define the stability of the new complexes under solution conditions conventionally used for biological testing, the time-dependent UV-vis spectra were obtained at 37 °C and no changes were observed (Fig. S4†), suggesting that these complexes are stable in the testing solution. The MTT results indicate that all three complexes significantly inhibited the proliferation of HeLa and Eca-109 cells in a dose-dependent manner (Fig. 8). As shown in Table 3, complexes 1–3 have lower IC₅₀ values for both cell lines as compared to those of the H₂L ligand and cisplatin, indicating that the complexes have higher cytotoxic activities against Eca-109 and HeLa cells than that of H₂L ligand and cisplatin. Complex 3 has the highest cytotoxic activity towards both cell lines, followed by 2 and 1, which was consistent with the binding affinity of the complexes with DNA and BSA. The results indicate that the larger aromatic ring may afford the improved biological activity. Although the complexes are stable in the medium, we still detect the effects of Cd²⁺ on the tumor cell viability. The number of moles of Cd(CH₃COO)₂·2H₂O similar to that of complex 3 were used to treat HeLa and Eca-109 cells. We observed that Cd(CH₃COO)₂·2H₂O shows weak inhibitory activity, which was much lower than that observed for complex 3, indicating that the antitumor effect of complex 3 was not due to Cd²⁺ (Fig. 8C). Consistently, we observed dramatic changes in the HeLa cell morphology after treatment with complex 3 but not with Cd(CH₃COO)₂·2H₂O (Fig. 8D).

3.5 Apoptosis evaluation

We further investigated whether complex 3 could induce apoptosis and cell cycle arrest in the HeLa cells.

3.5.1 Hoechst 33285 staining. Changes in the cell morphology can be detected using Hoechst 33258 staining.⁵³ As shown in Fig. 9, the control HeLa cells without complex 3 treatment manifested round and homogeneous nuclei. However, complex 3-treated HeLa cells showed bright blue nuclei due to karyopyknosis and chromatin condensation, which are typical characteristics of apoptosis. Complex 3 dose-dependently induced karyopyknosis and chromatin condensation in the HeLa cells, suggesting that complex 3 can induce apoptosis in HeLa cells.

3.5.2 Annexin V-FITC/PI staining. The effect of complex 3 on the induction of apoptosis was further investigated using Annexin V-FITC/PI staining. HeLa cells were treated with different concentrations of complex 3. After 24 h, cells were stained with Annexin V-FITC/PI and the samples were analyzed by flow cytometry. The results show that complex 3 induced HeLa cell apoptosis in a dose-dependent manner (Fig. 10), which was in agreement with the results obtained from the Hoechst 33258 staining experiments. The percentage of the apoptotic cells including early (Annexin V⁺PI⁻) and late (Annexin V⁺PI⁺) apoptosis was significantly increased from 11.6% (control) to 59.6% (when treated with 16 μg mL⁻¹ of complex 3). Although the percentage of necrotic cells (Annexin V⁻PI⁺) was significantly increased from 0.3% to 9.0%, it was a small population as compared to that of the apoptotic cells. These results suggest that complex 3 can effectively induce the apoptosis of HeLa cells.

3.5.3 Complex 3 influenced the cell cycle distribution. The DNA intercalative binding properties of many metal complexes may block the replication of DNA to induce the cell cycle arrest in tumor cells.⁵⁴ After treatment with complex 3, the cell cycle distribution of HeLa cells was examined using flow cytometry. As shown in Fig. 11, complex 3 increased the percentage of HeLa cells at the G0/G1 phase and decreased the percentage of HeLa cells at the S phase, suggesting that complex 3 induced cell cycle arrest at the G0/G1 phase. This is consistent with the DNA binding ability of complex 3. In addition, high doses of complex 3 significantly increased the frequency of cells at the sub-G0 phase, which was consistent with the results observed for the apoptosis experiments (Scheme 1).

Fig. 11 The cell cycle changes observed in the HeLa cells after treatment with complex 3 for 24 h. *p < 0.05, **p < 0.01 and ***p < 0.001 vs. the untreated cells.

Scheme 1 Complex 3 inhibits the proliferation of HeLa cells.

4. Conclusions

In summary, three cadmium(II) complexes, [Cd(HL)₂(CH₃OH)₂]·(CH₃OH) (1), [Cd(HL)₂(bpy)]·(CH₃OH)₂ (2), and [Cd(HL)₂(phen)]·(CH₃OH)_1.5 (3), based on 4-acyl pyrazolone derivatives, have been synthesized and fully characterized. Structural analysis revealed that all of them have a distorted octahedral geometry around the cadmium ion. The H₂L ligand and complexes 1–3 can interact with DNA via intercalation and the DNA binding affinity followed the order of 3 > 2 > 1 > H₂L. The H₂L ligand and complexes 1–3 also bound to BSA via a static quenching mechanism through interactions with both tyrosine and tryptophan residues. The H₂L ligand and complexes 1–3 showed an inhibitory effect on the proliferation of Eca-109 and HeLa cells in vitro and the cytotoxicity followed the order of 3 > 2 > 1 > H₂L. Moreover, the inhibitory effect of complex 3 on the proliferation of HeLa cells may be mediated by the induction of apoptosis and cell cycle arrest. These results indicate that complex 3 might be a potential antitumor drug candidate.

Acknowledgements

This work was financially supported by the National Natural Science Foundation of China (No. 21161019, 21361025), Natural Science Fund for Distinguished Young Scholars of Xinjiang Uygur Autonomous (No. 2013711008), Technological Innovation Youth Training Project of Xinjiang Autonomous (No. 2013721017) and the Graduate Research Innovation Project of Xinjiang (XJGRI2015016).

References

1. B. Rosenberg, L. VanCamp, J. E. Trosko and V. H. Mansour, Nature, 1969, 222, 385–386.
2. A. Florea and D. Büsselberg, Cancers, 2011, 3, 1351–1371.
3. B. W. Stewart and C. P. Wild, World Cancer Report, 2014.
4. E. Sundaravadivel, S. Vedavalli, M. Kandaswamy, B. Varghese and P. Madankumar, RSC Adv., 2014, 4, 40763–40775.
5. M. Manjunath, A. D. Kulkarni, G. B. Bagihalli, S. Malladi and S. A. Patil, J. Mol. Struct., 2017, 1127, 314–321.
6. A. Kimata, H. Nakagawa, R. Ohyama, T. Fukuuchi, S. Ohta, T. Suzuki and N. Miyata, J. Med. Chem., 2007, 50, 5053–5056.
7. R. Loganathan, S. Ramakrishnan, E. Suresh, M. Palaniandavar, A. Riyasdeen and M. A. Akbarsha, Dalton Trans., 2014, 43, 6177–6194.
8. U. Golla, A. Adhikary, A. K. Mondal, R. S. Tomar and S. Konar, Dalton Trans., 2016, 45, 11849–11863.
9. N. Parmar, S. Teraiya, R. Patel, H. Patel, H. Jajda and V. Thakkar, J. Saudi Chem. Soc., 2015, 19, 36–41.
10. X. Y. Wang and Z. J. Guo, Chem. Soc. Rev., 2013, 42, 202–224.
11. S. Medici, M. Peana, V. M. Nurchi, J. I. Lachowicz, G. Criponi and M. A. Zoroddu, Coord. Chem. Rev., 2015, 284, 329–350.
12. G. C. Xu, L. Zhang, L. Liu, G. F. Liu and D. Z. Jia, Polyhedron, 2008, 27, 12–24.
13. R. N. Jadeja, M. Chhatrola and V. K. Gupta, Polyhedron, 2013, 63, 117–126.
14. K. M. Vyas, R. N. Jadeja, D. Patel, R. V. Devkar and V. K. Gupta, Polyhedron, 2014, 80, 20–33.
15. G. Mariappan, B. P. Saha, L. Sutharson, S. Ankit, S. Garg, L. Pandey and D. Kumar, J. Pharmacol. Res., 2010, 3, 2856–2859.
16. V. A. Joseph, M. V. Komal, H. P. Jignesh, K. G. Vivek and R. N. Jadeja, J. Coord. Chem., 2013, 66, 1094–1106.
17. J. S. Casas, M. S. García-Tasende and A. Sánchez, Coord. Chem. Rev., 2007, 251, 1561–1589.
18. X. H. Wang, D. Z. Jia, Y. J. Liang, S. L. Yan, Y. Ding, L. M. Chen, Z. Shi, M. S. Zeng, G. F. Liu and L. W. Fu, Cancer Lett., 2007, 249, 256–270.
19. S. A. D. Pascali, D. Migoni, M. Monari, C. Pettinari, F. Marchetti, A. Muscella and F. P. Fanizzi, Eur. J. Inorg. Chem., 2014, 2014, 1249–1259.
20. Y. Li, J. Zhao, C. C. He, L. Zhang, S. R. Sun and G. C. Xu, J. Inorg. Biochem., 2015, 150, 28–37.
21. R. Pettinari, F. Marchetti, C. Pettinari, A. Petrini, R. Scopelliti, C. M. Clavel and P. J. Dyson, Inorg. Chem., 2014, 53, 13105–13111.
22. L. Li, H. Yang, D. Chen, C. Cui and Q. P. Dou, Toxicol. Appl. Pharmacol., 2008, 229, 206–214.
23. X. P. Zhou, Y. Koizumi, M. X. Zhang, M. Natsui, S. Koyota, M. Yamada, Y. Kondo, F. Hamada and T. Sugiyama, Cancer Sci., 2015, 106, 635–641.
24. Y. X. Tai, Y. M. Ji, Y. L. Lu, M. X. Li, Y. Y. Wu and Q. X. Han, Synth. Met., 2016, 219, 109–114.
25. J. D. Henderson, F. P. Filice, M. S. M. Li and Z. F. Ding, J. Inorg. Biochem., 2016, 158, 92–98.
26. K. Sinha, P. B. Pal and P. C. Sil, Toxicol. In Vitro, 2014, 28, 307–318.
27. J. Zhao, L. Zhang, J. Y. Li, T. Wu, M. F. Wang, G. C. Xu, F. C. Zhang, L. Liu, J. H. Yang and S. R. Sun, Chem.-Biol. Interact., 2015, 231, 1–9.
28. B. S. Jensen, Acta Chem. Scand., 1959, 13, 1668–1670.
29. M. Alagesan, N. S. P. Bhuvanesh and N. Dharmaraj, Dalton Trans., 2013, 42, 7210–7223.
30. J. Marmur, J. Mol. Biol., 1961, 3, 585–594.
31. M. E. Reichmann, S. A. Rice, C. A. Thomas and P. Doty, J. Am. Chem. Soc., 1954, 76, 3047–3054.
32. F. M. Wang, Z. Y. Yang, Y. Li and H. G. Li, J. Coord. Chem., 2011, 64, 2974–2983.
33. A. Wolfe, G. H. Shimer Jr and T. Meehan, Biochemistry, 1987, 26, 6392–6396.
34. K. Jeyalakshmi, Y. Arun, N. S. P. Bhuvanesh, P. T. Perumal, A. Sreekantha and R. Karvembu, Inorg. Chem. Front., 2015, 2, 780–798.
35. J. R. Lakowicz and G. Webber, Biochemistry, 1973, 12, 4171–4179.
36. R. R. Kumar, M. K. M. Subarkhan and R. Ramesh, RSC Adv., 2015, 5, 46760–46773.
37. Y. J. Hu, Y. Liu, J. B. Wang, X. H. Xiao and S. S. Qu, J. Pharm. Biomed. Anal., 2004, 36, 915–919.
38. T. Mosmann, J. Immunol. Methods, 1983, 65, 55–63.
39. R. Pradhan, M. Banik, D. B. Cordes, A. M. Z. Slawin and N. C. Saha, Inorg. Chim. Acta, 2016, 442, 70–80.
40. K. Zheng, M. X. Yan, Y. T. Li, Z. Y. Wu and C. W. Yan, Eur. J. Med. Chem., 2016, 109, 47–58.
41. X. Q. Zhou, Y. Li, D. Y. Zhang, Y. Nie, Z. J. Li, W. Gu, X. Liu, J. L. Tian and S. P. Yan, Eur. J. Med. Chem., 2016, 114, 244–256.
42. M. Ganeshpandian, R. Loganathan, S. Ramakrishnan, A. Riyasdeen, M. A. Akbarsha and M. Palaniandavar, Polyhedron, 2013, 52, 924–938.
43. M. Alagesan, P. Sathyadevi, P. Krishnamoorthy, N. S. P. Bhuvanesh and N. Dharmara, Dalton Trans., 2014, 43, 15829–15840.
44. S. Jovanović, K. Obrenčević, Ž. D. Bugarčić, I. Popović, J. Žakula and B. Petrović, Dalton Trans., 2016, 45, 12444–12457.
45. D. Prieto, G. Aparicio, P. E. Morande and F. R. Zolessi, Histochem. Cell Biol., 2014, 142, 335–345.
46. A. K. Krey and F. E. Hahn, Biochemistry, 1975, 14, 5061–5067.
47. S. M. T. Shaikh, J. Seetharamappa, S. Ashoka and P. B. Kandagal, Dalton Trans., 2016, 45, 12518–12531.
48. K. Zheng, F. Liu, Y. T. Li, Z. Y. Wu and C. W. Yan, J. Inorg. Biochem., 2016, 156, 75–88.
49. M. R. Eftink and C. A. Ghiron, J. Phys. Chem., 1976, 80, 486–493.
50. W. R. Ware, J. Phys. Chem., 1962, 66, 455–458.
51. Y. H. Liu, A. Li, J. Shao, C. Z. Xie, X. Q. Song, W. G. Bao and J. Y. Xu, Dalton Trans., 2016, 45, 8036–8049.
52. A. Paul, R. K. Gupta, M. Dubey, G. Sharma, B. Koch, G. Hundal, M. S. Hundal and D. S. Pandey, RSC Adv., 2014, 4, 41228–41236.
53. Q. P. Qin, Y. H. Liu, H. L. Wang, J. L. Qin, F. J. Cheng, S. F. Tang and H. Liang, Metallomics, 2015, 7, 1124–1136.
54. X. Qiao, Z. Y. Ma, C. Z. Xie, F. Xue, Y. W. Zhang, J. Y. Xu, Z. Y. Qiang, J. S. Lou, G. J. Chen and S. P. Yan, J. Inorg. Biochem., 2011, 105, 728–738.

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Footnote

† Electronic supplementary information (ESI) available. CCDC 1506204–1506207. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c6ra23938b

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