Interaction of copper(II) (3 + 2) N-chelating complexes with DNA and BSA: hydrolytic DNA cleavage and ROS-mediated apoptosis

Abstract

Mixed-ligand copper(II) complexes of the form [Cu(L)(phen)](ClO₄)₂, where L is (6-methylpyridin-2-ylmethylene)-(pyridin-2-ylmethyl)amine (L1) and (6-methylpyridin-2-ylmethylene)-(pyridin-2-ylethyl)amine (L2), are obtained and characterized using analytical, spectral, and electrochemical methods. A distorted square pyramidal coordination geometry is observed in the single-crystal X-ray structure of [Cu(L2)(phen)](ClO₄)₂. DFT calculations of copper(II) complexes in solution offered insights into the molecular geometry, electronic properties, and quantum chemical descriptors. Complexes interact hydrophobically with calf thymus (CT) DNA through partial intercalation, as demonstrated by absorption spectral titration, EthBr displacement assay, and circular dichroic spectral and electrochemical investigations. They undergo a static process of quenching when exposed to bovine serum albumin (BSA) protein. Both complexes cleave pUC19 supercoiled (SC) DNA in the absence of an activator at concentrations of 130–140 μM, degrading SC DNA into nicked circular (NC) DNA and thereby indicating that they are effective hydrolytic chemical nucleases. They are notable for exhibiting cytotoxicity against human cervical carcinoma (HeLa) and human lung epithelial adenocarcinoma (A549) cell lines with potency greater than cisplatin, demonstrating that they have the potential to be effective anticancer drugs. They preferentially target cancer cells because they are non-toxic to human lung epithelial (L132) cells and normal mouse embryonic fibroblasts (NIH 3T3). AO/EB and DAPI staining, annexin-V-FITC staining assay, propidium iodide assay, and metal-assisted reactive oxygen species generation indicated an apoptotic mode of cell death, possibly by causing membrane instability and initiating apoptosis. These complexes can also stop the proliferation of A549 cancer cells by causing cell cycle arrest at the S-phase. Their efficiency as promising drugs for targeted cancer treatment is highlighted in this study.

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Introduction

Many transition-metal complexes have been identified as potential chemotherapeutic drugs, and some of them are in various stages of clinical trials.¹ Among them, certain dinuclear platinum complexes have been found to be more effective anticancer drugs than cisplatin.^2–4 However, the widespread use of platinum-based drugs in medical applications has resulted in a variety of side effects, including nephrotoxicity, ototoxicity, and neurotoxicity, as well as intrinsic or acquired drug resistance.^5,6 As a result, the research community has been inspired to develop new low-toxicity compounds with improved pharmacological properties.^7,8 Among them, the metal ion selected also plays an outstanding role in chemotherapeutic drug development. Here, we opted for one of the most abundant transition metals, specifically the less toxic and biocompatible copper. Copper, a bio-essential metal ion, is involved in many biological processes, most notably the regular functioning of cells. Copper(II) complexes with well-defined redox activity can be effective antioxidants, antimicrobials, antiparasitics, and anticancer agents.^9,10 Some families of copper complexes have been studied as prospective anticancer agents in recent years,⁸ and our new copper complexes are considered better alternatives to platinum complexes.

The significance of copper complexes in anticancer research has grown since the discovery of their antiproliferative activity. Researchers have examined the structure–activity relationship of known copper complexes to develop therapeutic molecules for cancer therapy.¹¹ Due to its importance in tumour growth, angiogenesis, and apoptosis, Cu has become increasingly essential in cancer treatment.¹² In chemotherapeutic applications, the anticancer activity of copper complexes is greatly affected by the ligand environment, which also determines the reduction potential of the Cu(II)/Cu(I) system.^13,14 According to Ruiz-Azuara and coworkers, Casiopeínas®, a copper(II) compound, exhibits selective cytotoxic, genotoxic, and antineoplastic effects. It interacts with DNA to promote mitochondrial toxicity and the production of reactive oxygen species (ROS), which damage cells and trigger apoptosis via the intrinsic pathway.^15,16 Based on recent reports, copper(II) complexes containing Schiff bases^9c have shown substantial antiproliferative activity and selectivity for lung adenocarcinoma cells, while copper(II) complexes containing 1,10-phenanthroline¹⁷ have demonstrated excellent inhibition activity on A549 tumour cells and induced cell death through apoptosis. Cubisbel (copper-bis-belinostat), as described by Marmion and coworkers,¹⁸ dramatically slowed the development of colon cancer cells by inhibiting histone deacetylase (HDAC) and inducing apoptosis. Interfering with important master regulators (MRs) in colon cancer improves metabolic stability in vitro and targets anticancer pathways. Tsvetkov et al. demonstrated that the mechanism of copper toxicity is different from all other known methods of controlled cell death, such as ferroptosis, pyroptosis, necroptosis, and apoptosis. Cuproptosis, a copper-dependent form of apoptosis, is induced by the direct binding of copper to the lipoylated components of the tricarboxylic acid (TCA) cycle. This results in the loss of iron–sulfur cluster proteins and the aggregation of lipoylated proteins, which ultimately leads to proteotoxic stress and cell death.¹⁹ Because their mechanisms of action are different from those of other metal complexes (especially platinum drugs), copper complexes have garnered attention and raised hopeful expectations.^20,21 Due to their strong DNA-binding/cleaving ability, mixed-ligand copper(II) complexes have recently been discovered to be good anticancer agents via inducing apoptosis.^22–24 Therefore, considering the advantages of processes that produce fragments similar to those formed by restriction enzymes,^25,26 designing suitable copper complexes for DNA binding and cleavage under both oxidative and hydrolytic conditions, depending on the recognition elements in the ligand, is of importance.

We synthesized and characterized two mixed ligand copper(II) complexes of the type [Cu(L1/L2)(phen)](ClO₄)₂, where L1 or L2 is a tridentate 3N Schiff base ligand derived from the condensation of 6-methylpyridine-2-carboxaldehyde with 2-aminomethylpyridine (L1) or 2-aminoethylpyridine (L2) (Scheme 1). Different degrees of flexibility can be imposed upon these complexes by alterations in the spacers between the pyridine and imine nitrogen of the ligands. The L1 was modified in order to investigate the impact of the number of chelate ring members on biological activity when the methyl spacer (L1, five-membered) was substituted with an ethyl spacer (L2, six-membered). Copper(II) complexes with the simple or methylated 3N tridentate ligand have attracted the most attention among the various coordination molecules due to their strong hydrolytic cleavage of genomic DNA and potent anticancer properties.^27,28 As a result, the biological activity of 3N Schiff base is linked to its capacity to chelate transition metal ions; coordinating with the metal ion via nitrogen atoms may increase biological activity by acting as a strong DNA cleaver and efficient cytotoxic substance.^25,29 It has been reported that combining a single phen moiety can increase activity; however, combining two phen moieties in the same complex can retard biological processes, especially under physiological conditions.³⁰ The interactions of these complexes with CT DNA and BSA were studied using various spectral and electrochemical techniques, and the intent was to explore binding properties. Further, the cleavage activities of the complexes with pUC19 supercoiled plasmid DNA was investigated. The chemotherapeutic efficacy of the complexes was studied in terms of cytotoxicity against human cervical carcinoma (HeLa) and human lung epithelial adenocarcinoma (A549) cells and non-cancerous mouse embryonic fibroblast (NIH 3T3) and human lung epithelial (L132) cell lines. The copper(II) complexes are more sensitive to cancerous A549 cells; hence, this cell line was selected to detect the mode of cell death, such as (i) cell cycle arrest, (ii) the production of reactive oxygen species (ROS), and (iii) the induction of apoptosis. Interestingly, the present mixed ligand copper(II) complexes of tridentate 3N Schiff base ligands with phen are involved in partial intercalative binding with DNA, display hydrolytic DNA cleavage, exhibit greater anticancer activities, induce apoptosis, and are predominantly non-toxic to normal cells.

Scheme 1 Schematic of the copper(II) complex (a and b) and structure of ligands (L1, L2, and phen) and the proton numbering scheme (L1 and L2).

Experimental

Materials

Copper(II) acetate monohydrate, 1,10-phenanthroline (Merck), 2-aminomethylpyridine, 2-aminoethylpyridine, 6-methyl-pyridine-2-carboxaldehyde, ethidium bromide (EthBr), sodium azide (NaN₃), potassium iodide (KI), sodium hydrogen phosphate monohydrate, di-sodium hydrogen phosphate dehydrate, (Sigma-Aldrich), L-ascorbic acid (Fisher Scientific), hydrogen peroxide solution (30% w/v), tetra-N-butylammonium bromide, perchloric acid, sodium perchlorate (Sigma Aldrich), pUC19 supercoiled DNA and agarose (Genei). Calf thymus (CT) DNA (highly polymerized, stored at 4 °C), superoxide dismutase (SOD) (stored at 4 °C), bovine serum albumin (BSA) (stored at 4 °C), catalase (stored at 4 °C), and T4 DNA ligase enzyme (stored at 4 °C) (Sigma Aldrich) were used as received without further purification. Methanol, ethanol, diethyl ether, acetone, HPLC-grade dimethylformamide (DMF), dimethyl sulfoxide (DMSO), and acetonitrile were purchased from Merck. Ultrapure Milli Q water (18.2 μΩ) was used for all the experiments.

Physical measurements

The elemental analyses (C, H, N) were carried out using a PerkinElmer 2400 series II analyzer. The electrical conductivity was obtained with a Systronic 305 conductivity bridge using 1 × 10⁻³ M solution of the complex in DMF. FTIR spectra were recorded using a PerkinElmer Paragon 1000 FTIR spectrophotometer, equipped with a Golden Gate Diamond ATR device, applying the reflectance technique (4000–300 cm⁻¹), and the peaks are reported in cm⁻¹. PerkinElmer Lambda 35 and PerkinElmer Lambda 365 were used to record diffuse reflectance spectra, and a UV-VIS spectrophotometer was attached to a refrigerated circulating water bath (25 ± 0.2 °C) using cuvettes (path length, 1 cm) to capture solution electronic absorption spectra in the 200–1100 nm range. The X-band EPR spectra of the solid at ambient temperature and the solution at 77 K were acquired using a JEOL JES-FA200 ESR spectrometer at an X-band frequency (9–10 GHz) with 100 kHz field modulation. ¹H NMR spectra were recorded using a Bruker DPX 300 (300 MHz) spectrometer. Chemical shifts are reported as δ (ppm) values (multiplicity, integration, and assignment) relative to the DMSO-d₆ solvent peak. Emission intensity measurements were carried out using a Shimadzu RF-5301PC spectrofluorophotometer equipped with a thermostatic bath. A circular dichroic spectrum of DNA or BSA was obtained using a JASCO J-716 spectropolarimeter. A CHI 620C electrochemical analyzer operating at 25 ± 0.2 °C was used to evaluate the redox potentials of copper(II) complexes 1 and 2 (0.001 M) in methanol with TBAP (0.1 M) as the supporting electrolyte. Cyclic voltammetry (CV) and differential pulse voltammetry (DPV) were employed. The standard configurations of three electrodes, such as the working electrode (glassy carbon; A, 0.0707 cm²), counter electrode (platinum wire), and reference electrode (saturated calomel), were employed. The potential varied between −1 and +1 V at scan speeds of 0.01–0.1 V s⁻¹. Before measurements, N₂ was quickly bubbled into the solution to eliminate oxygen. Using the formula E_1/2 = (E_pc + E_pa)/2 and ΔE_p = E_pa − E_pc, redox data were calculated.

Synthesis of ligands

Synthesis of (6-methylpyridin-2-ylmethylene)-(pyridin-2-ylmethyl)amine, L1. 2-Aminomethylpyridine (0.55 g, 5 mmol) in ethanol (15 mL) was added dropwise to 6-methyl-pyridine-2-carboxaldehyde (0.60 g, 5 mmol) in ethanol (15 mL). The mixture was refluxed for 3 h with constant stirring. After the solvent was removed under reduced pressure, 20 mL of CH₂Cl₂ was then added, and it was rotaevaporated to obtain (6-methylpyridin-2-ylmethylene)-(pyridin-2-ylmethyl)amine as a bright yellow oil. Yield: ∼0.76 g, 72%. Anal. calcd for C₁₃H₁₃N₃: C, 73.91; H, 6.20; N, 19.89. Found: C, 73.99; H, 6.15; N, 19.83%. Selected FT-IR (cm⁻¹) bands in KBr: 1610 ν_imine(C=N), 1525 ν(C=C), 982 ν_C–H(H–C=N), 726, and 782 ν(C–H). HRMS value in CH₂Cl₂: m/z, 211.98 (M + H)⁺. ¹H NMR (300 MHz, DMSO-d₆) δ/ppm: 7.45 (d, 1H, H_d), 7.06 (t, 1H, H_e), 7.21 (d, 1H, H_f), 8.91 (s, 1H, H_g), 3.79 (s, 2H, H_h), 7.36 (d, 1H, H₃), 7.56 (t, 1H, H₄), 7.14 (t, 1H, H₅), 8.53 (d, 1H, H₆), 2.46 (s, 3H, Py-CH₃).

Synthesis of (6-methylpyridin-2-ylmethylene)-(pyridin-2-ylethyl)amine, L2. Ligand L2 was prepared using the method adopted for the preparation of L1, except that 2-aminoethylpyridine (0.11 g, 1 mmol) was used instead of 2-aminomethylpyridine. Yield: ∼0.90 g, 78%. Anal. calcd for C₁₄H₁₅N₃: C, 74.64; H, 6.71; N, 18.65. Found: C, 74.61; H, 6.79; N, 18.71%. Selected FT-IR (cm⁻¹) bands in KBr: 1611 ν_imine(C=N), 1529 ν(C=C), 981 ν_C–H(H–C=N), 730, and 783 ν(C–H). HRMS value in CH₂Cl₂: m/z, 226.22 (M + H)⁺. ¹H NMR (300 MHz, DMSO-d₆) δ/ppm: 7.43 (d, 1H, H_d), 7.07 (t, 1H, H_e), 7.25 (d, 1H, H_f), 8.95 (s, 1H, H_g), 3.81 (t, 2H, H_h), 3.62 (t, 2H, H_i), 7.41 (d, 1H, H₃), 7.63 (t, 1H, H₄), 6.98 (t, 1H, H₅), 8.62 (d, 1H, H₆), 2.48 (s, 3H, Py-CH₃).

Synthesis of copper(II) complexes

The copper(II) complexes were prepared using the general procedure as follows: copper(II) acetate monohydrate (0.2 g, 1 mmol) in ethanol (15 mL) was slowly added to 1,10-phenanthroline monohydrate (0.20 g, 1 mmol) in ethanol (15 mL), and the mixture was stirred for 1 h to ensure complete dissolution. To this mixture, ligand L1 (0.21 g, 1 mmol) or L2 (0.23 g, 1 mmol) was added with stirring to the reaction mixture. The resulting green solution was refluxed at 70 °C for 3 h. After the complexation, it was filtered while hot, and an ethanolic solution of NaClO₄ (0.12 g, 1 mmol) was added. The solution was cooled to room temperature, and green colored crystalline solids of 1 or 2 were formed after three days. They were filtered off, washed with a small amount of cold ethanol, and dried under a vacuum over P₄O₁₀.

[Cu(L1)(phen)](ClO₄)₂1. Yield: ∼0.37 g (63%). Found, C, 50.84; H, 3.55; N, 11.91%. C₂₅H₂₁N₅O₈Cl₂Cu requires C, 50.86; H, 3.59; N, 11.86%. Λ_M in DMF solution, 203 Ω⁻¹ cm⁻¹ mol⁻¹. ESI-MS m/z = 391.25 [Cu(L1)(phen)]²⁺ (calcd 391.48). μ_eff, 1.86 μ_B (27 °C). Selected FT-IR (cm⁻¹) bands in KBr: 1585 ν_imine(C=N), 1519 ν(C=C), 852 ν_C–H(H–C=N), 724, 778 ν(C–H), 1090, 622 ν(ClO₄⁻), 530, and 562 ν(Cu–N). UV-Visible data in DMF [λ_max/nm (ε_max/dm³ mol⁻¹ cm⁻¹)]: 684 (100), 896 sh, 323 (4485), 294 (24 670), 268 (46 420). EPR data: in solid (RT) g_iso = 2.147; in DMF solution (77 K), g_∥ = 2.233, g_⊥ = 2.045, A_∥ = 188 × 10⁻⁴ cm⁻¹, g_∥/A_∥ = 119 cm, G = 5.2. Electrochemical data (DMF, 0.1 M TBAP): cyclic voltammetry, E_1/2 = −0.077 V, ΔE_p = 100 mV, i_pa/i_pc = 0.7, D = 7.7 × 10⁻⁶ cm² s⁻¹, and differential pulse voltammetry, E_1/2 = −0.063 V.

[Cu(L2)(phen)](ClO₄)₂2. Yield: ∼0.39 g (65%). Found, C, 51.71; H, 3.89; N, 11.64%. C₂₆H₂₃N₅O₈Cl₂Cu requires C, 51.67; H, 3.84; N, 11.59%. Λ_M in DMF solution, 205 Ω⁻¹ cm⁻¹ mol⁻¹. ESI-MS m/z = 405.27 [Cu(L2)(phen)]²⁺ (calcd 405.50). μ_eff, 1.88 μ_B (27 °C). Selected FT-IR (cm⁻¹) bands in KBr: 1585 ν_imine(C=N), 1519 ν(C=C), 852 ν_C–H(H–C=N), 724, 779 ν(C–H), 1090, 622 ν(ClO₄⁻), 538, 571 ν(Cu–N). UV-Visible data in DMF [λ_max/nm (ε_max/dm³ mol⁻¹ cm⁻¹)]: 675 (280), 856 sh, 321 (7895), 293 (32 420), 268 (49 330). EPR data: in solid (RT) g_iso = 2.172; in DMF solution (77 K), g_∥ = 2.233, g_⊥ = 2.042, A_∥ = 189 × 10⁻⁴ cm⁻¹, g_∥/A_∥ = 118 cm, G = 5.5. Electrochemical data (DMF, 0.1 M TBAP): cyclic voltammetry, E_1/2 = −0.072 V, ΔE_p = 86 mV, i_pa/i_pc = 0.9, D = 8.2 × 10⁻⁶ cm² s⁻¹, and differential pulse voltammetry, E_1/2 = −0.059 V.

X-ray structure crystallography

The single crystal of [Cu(L2)(phen)](ClO₄)₂ (2) (CCDC no. 2143125) in a green color suitable for the determination of X-ray structure was acquired upon slow evaporation at 5 °C after a couple of weeks in a MeOH : MeCN (2 : 1 v/v) mixture. A single crystal of the size 0.350 × 0.320 × 0.210 mm³ was chosen under the polarizing microscope and then mounted on the glass fiber. Diffraction data for the complex were collected at 298 K using a Bruker AXS-KAPPA APEX II diffractometer (Mo-K_α radiation, λ = 0.71073 Å) in the θ range 2–25°. Unit cell parameters were determined using the least-squares method based on all reflections with F² > 2σ(F²) with Bruker SMART software. Cell refinement, correction for Lorentz and polarization effects, integration of intensities, and absorption corrections were performed using Bruker SAINT and SADABS. The structure was solved using the direct method (SIR-92)³¹ and refined using a full-matrix least-squares method (SHELXL-2014).³² All the calculations were carried out using the programs in the WinGX module.³³ Non-hydrogen atoms were refined anisotropically. Cl1 and O1, O2, O3, and O4 were found to be disordered over two positions, and the occupancy factors were assigned to 0.775(10) and 0.225(10). O5, O6, O7, and O8 were also disordered over two positions, and the occupancy factors were 0.506(8) and 0.494(8). All the hydrogen atoms were placed geometrically and refined with the riding model. The final difference Fourier maps displayed no peaks of chemical importance. The graphic representation of the molecule was prepared using the ORTEP3 program for Windows³⁴ with 40% probability displacement ellipsoids.

Theoretical methods

The coordination geometries of copper(II) mixed ligand complexes in the ground state with doublet spin state were optimized using density functional theory (DFT) at the B3LYP level of theory by employing the Gaussian 09 program package.³⁵ The calculations were administered with a mixed basis set (B1) of LANL2DZ for the copper metal atom, which contains a relativistic effective core potential with a valence basis set and 6-31G* for the remaining atoms.^36–38 The normal mode analyses were performed to examine the minimal energy nature of the geometry. Copper(II) complexes 1 and 2 were considered mono-positive cations, as the copper metal was considered in the +2 oxidation state and doublet spin state, while the ligand was considered a positive ligand. The salvation of the complexes was achieved by employing the CPCM method using methanol as the solvent.

Charge distribution and molecular orbital analyses

We used the natural bond orbital (NBO)³⁹ approach to calculate the atomic charge distribution on each complex atom. From the NBO analysis, we can predict electron delocalization, atomic charge distribution, and intra-atomic interaction of atoms within a compound.³⁹ The energies of the frontier molecular orbitals (FMOs),⁴⁰ the highest and second-highest occupied molecular orbitals (E_HOMO and E_HOMO−1), and the lowest and second-lowest unoccupied molecular orbitals (E_LUMO and E_LUMO+1) were calculated. Some quantum chemical descriptors, including ionization potential (IP),⁴¹ electron affinity (EA),⁴¹ band gap (ΔE), chemical hardness (η),⁴² global softness (S),⁴² electrochemical potential (μ),⁴³ electrophilicity index (ω),⁴⁴ and electronegativity (χ),⁴³ were estimated as outlined in eqn (1)–(8).

IP = −E_HOMO (1)

EA = −E_LUMO (2)

ΔE = E_LUMO − E_HOMO (3)

S = 1/η (5)

χ = −μ (8)

DNA and protein binding experiments

Solutions of DNA in 5 mM Tris HCl/50 mM NaCl buffer gave a ratio of UV absorbances at 260 and 280 nm, A₂₆₀/A₂₈₀, of 1.9,⁴⁵ indicating that the DNA was sufficiently free of protein. The stock solution of protein (1.0 × 10⁻⁴ mol L⁻¹) was prepared by dissolving the solid BSA in 0.05 M phosphate buffer at pH 7.4, stored at 0–4 °C in the dark for about a week, and then diluted to 1.0 × 10⁻⁶ mol L⁻¹ using phosphate buffer (pH 7.4, 0.05 M) when used. The concentration of BSA was determined from optical density measurements using the value of molar absorptivity of ε₂₈₀ = 44 720 M⁻¹ cm⁻¹.⁴⁶ The detailed procedure for the spectroscopic experiments of DNA and protein binding, as well as the electrochemical experiments for DNA binding, is described in the SI.

DNA cleavage experiments

The cleavage of DNA in the absence of activating agents was monitored using agarose gel electrophoresis. A typical reaction mixture containing pUC19 supercoiled phage DNA (form I, 20 µM) and copper(II) complex in 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer (pH 7.1) was incubated at 37 °C for 1 h. After the incubation period, the reaction was quenched by keeping the samples at −20 °C, followed by the addition of loading buffer (0.025 mg bromophenol blue, 1 mL glycerol, and 1 mL MilliQ water). This was then loaded on a 1% agarose gel containing ethidium bromide (2.54 µM in the gel and buffer). The gels were run at a constant voltage of 40 V for 3 h in the 1 × TBE buffer (TBE = Tris-Borate-EDTA buffer) containing ethidium bromide. After washing with distilled water, the gels were visualized under a UV transilluminator, and the bands were documented and quantified using a BioRad Gel Doc 1000 apparatus interfaced with a computer. The cleavage efficiency was measured by determining the ability of the complex to convert the supercoiled DNA (SC) to a nicked circular form (NC) and linear form (LC). To identify the reactive oxygen species (ROS) involved in the cleavage reaction, radical scavengers such as hydroxyl radical (DMSO, 20 μM), singlet oxygen (NaN₃, 100 μM), superoxide (SOD, 0.5 units), and catalase (0.5 units) were introduced. Additionally, methyl green (100 μM) is used to determine the groove binding nature of copper(II) complexes.

Religation studies

An enzymatic study was carried out using T4 DNA ligase to determine whether the cleaved products were consistent with the hydrolysis of the phosphodiester linkages in DNA. For the religation tests, the solution was incubated for 18 h at 16 °C prior to gel electrophoresis. The nicked circular (NC) DNA obtained from the hydrolytic cleavage reaction was recovered from the agarose gel using a phenol extraction method and purified by ethanol precipitation. This was followed by the addition of a 10× ligation buffer and T4 DNA ligase (4 units) to the purified NC DNA.

Cell line

The human cervical cancer cell line (HeLa), human lung epithelial adenocarcinoma cell line (A549), normal mouse embryonic fibroblast cells (NIH 3T3), and human lung epithelial normal cell line (L132) were acquired from the National Centre for Cell Science (NCCS), Pune. They were grown in Eagle's minimum essential medium containing 10% fetal bovine serum (FBS). The cells were preserved at 37 °C, 5% CO₂, 95% air, and 100% relative humidity.⁴⁷ Maintenance cultures were channelized weekly, and the culture medium was altered twice a week.

Cell culture

To make single-cell suspensions, the monolayer cells were detached with trypsin-ethylenediaminetetraacetic acid (EDTA), and viable cells were counted using a hemocytometer. They were diluted with a medium containing 5% FBS to give a final density of 1 × 10⁵ cells per mL. The cell suspension of one hundred microlitres per well was seeded into 96-well plates at a plating density of 10 000 cells per well. They were incubated to allow for cell attachment at 37 °C, 5% CO₂, 95% air, and 100% relative humidity. The cells were treated with serial concentrations of the test samples after 24 h. They were dissolved in dimethylsulfoxide (DMSO), and an aliquot of the sample solution was diluted twice to the desired final maximum test concentration with a serum-free medium. Additionally, four serial dilutions were made to give a total of five sample concentrations. Aliquots of 100 µl of different sample dilutions were added to the suitable wells already containing 100 µl of the medium, resulting in the required final sample concentrations. The plates were incubated, followed by sample addition at 37 °C, 5% CO₂, 95% air, and 100% relative humidity for an additional 48 h. The medium alone served as a control, and triplicate was maintained for all concentrations.

Cell viability assay

The cell viability was carried out using the MTT assay.⁴⁸ Complex in the concentration range of 0.25–10 μM (in HeLa), 0.2–1.0 μM (in A549), 25–500 μM (in NIH 3T3) or 25–250 μM (in L132) dissolved in 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1 was added to the wells 24 h after seeding of 1 × 10⁵ cells per well in 100 μL of fresh culture medium. After 48 h, 15 µL of 3-[4,5-dimethylthiazol-2-yl]2,5-diphenyltetrazolium bromide (MTT, 5 mg mL⁻¹) in phosphate-buffered saline (PBS) was added to each well and incubated at 37 °C for 4 h. The formed formazan crystals were solubilized in 100 µL of DMSO after the medium with MTT was flicked off. The microplate reader was used to measure the absorbance at 570 nm. Data were collected in three replicates, and the percentage of cell viability and the percentage of cell inhibition were calculated using the following formulas:

% Cell viability = [A_s]/[A_c] × 100

where A_s is the absorbance of the sample and A_c is the absorbance of the control,

% Cell inhibition = [100 − ([A_s]/[A_c])] × 100.

A nonlinear regression graph was plotted between % cell inhibition and log concentration, and IC₅₀ was calculated using GraphPad Prism software.

Studies on intracellular reactive oxygen species (ROS)

Fluorescent probe method. The production of intracellular ROS was measured using the fluorescent probe 2′,7′-dichloro-fluorescein diacetate (DCFH-DA). DCFH-DA diffuses through the cell membrane and is enzymatically hydrolyzed by intracellular esterases to form the non-fluorescent compound DCFH, which is then rapidly oxidized to form the highly fluorescent 2′,7′-dichlorofluorescein (DCF) in the presence of ROS. The DCF fluorescence intensity is parallel to the amount of ROS formed intracellularly.⁴⁹ The cultured A549 cancerous cells were treated with two copper(II) complexes at their IC₅₀ concentrations, and the untreated cells were maintained as a control. After 24 h, the cells were incubated with 10 μM of DCFH-DA at 37 °C for 30 min. The levels of intracellular ROS were examined using an inverted fluorescence microscope (Zeiss Axio Observer D1) and a FACSVerse flow cytometer (Becton Dickinson).³¹ When the excitation wavelength was 485 nm, fluorescence was observed at 528 nm. ROS levels inside the cells were expressed using the average fluorescence intensity.

Flow cytometry method. After the cells were treated with positive control H₂O₂ (250 μM) and IC₅₀ concentration of 1 or 2 for 24 h, the A549 cells were seeded in six-well plates with high glucose DMEM for the treatment until cell confluency. The cells were trypsinized, and 10 μM 2′,7′-dichlorodihydrofluorescein diacetate (DCHF-DA, Sigma-Aldrich) dye was added to the pellet after the treatment. This underwent 10 minutes, 37 °C, and total darkness incubation. Afterwards, cells were examined to evaluate their ROS levels using an inverted fluorescence microscope (Zeiss Axio Observer D1) and a FACSVerse flow cytometer (Becton Dickinson).⁵⁰ The fluorescence was detected at 528 nm, while the excitation wavelength was 485 nm. The average fluorescence intensity was used to express the amount of intracellular ROS.

Studies on cell cycle arrest

The A549 cells were seeded in six-well plates with high glucose DMEM after the cells had grown, and the cells were treated with two copper(II) complexes in their respective IC₅₀ concentrations in the medium for 24 h. After 24 h of incubation, the cells were trypsinized and resuspended in complete media. The cells were collected and centrifuged at 1000 rpm for 5 min. The cell pellet was washed with phosphate-buffered saline (PBS) twice and subsequently fixed with 1 mL of 70% ice-cold ethanol overnight at 4 °C. Following the ice-cold ethanol, the cell pellet was washed twice with cold PBS and added to 10 μL of RNase A at 10 g mL⁻¹ of concentration, which was then incubated for 30 min and washed with PBS at the end. The cells were then incubated in 1 mL of PBS with 50 μL of propidium iodide (PI) (1 mg mL⁻¹ of stock) for 30 min in the dark. Then, the cells were analyzed to check the cell cycle phase using the FACSVerse flow cytometer (Becton Dickinson).⁵⁰

Studies on the induction of apoptosis

AO (acridine orange)/EB (ethidium bromide) staining method. Apoptosis studies were performed using a staining method with AO and EB. A monolayer of A549 cells was incubated in the absence or presence of copper(II) complexes at their IC₅₀ concentration at 37 °C and 5% CO₂ for 24 h. After 24 h, the cells were stained with AO/EB solution (100 μg mL⁻¹ AO, 100 μg mL⁻¹ EB). The samples were viewed under a fluorescence microscope (Zeiss Axio Observer D1).

DAPI staining method. DAPI (4′,6′-diamidino-2-phenylindole) staining was performed using the method described earlier with slight modifications.⁵¹ The 5 × 10⁵ A549 cells were treated with two copper(II) complexes at their IC₅₀ concentration for 24 h in a 6-well culture plate and were fixed with 4% paraformaldehyde, followed by permeabilization with 0.1% Triton X-100. Cells were then stained with 50 μg mL⁻¹ DAPI for 30 min at room temperature. The cells undergoing apoptosis, represented by the morphological changes of apoptotic nuclei, were observed and imaged from ten eye views at 20× magnifications under a laser scanning confocal microscope LSM 710 (Zeiss Axio Observer D1).

Flow cytometry method. Apoptosis detection was performed using the FITC annexin V with PI (apoptosis detection kit, BD Biosciences, USA) protocol. A549 cells were treated with two 1 or 2 in high glucose DMEM for 24 h. After treatment, the cells were trypsinized, resuspended in PBS, washed twice, and centrifuged to remove PBS. The cells were suspended in 100 μL of binding buffer containing 5 μL of annexin V and PI and incubated for 15 min under darkness. The stained cells were diluted using 450 μL of binding buffer. The cells were analyzed using a FACSVerse flow cytometer (Becton Dickinson), and the data were analyzed using FACSVerse software.⁵⁰

Results and discussion

Synthesis and general properties

Schiff base ligands L1 and L2 were synthesized by condensing 2-aminomethylpyridine or 2-aminoethylpyridine and 5-methylpyridine-2-carboxaldehyde. The copper(II) complexes of the ligands were prepared by the reaction of copper(II) acetate monohydrate, 1,10-phenanthroline, and L1 or L2 in equimolar quantities using ethanol as a solvent. Both complexes 1 and 2 were isolated as green-colored crystalline compounds in good yields. Based on the elemental analysis, the complexes were formulated as [Cu(L1/L2)(phen)](ClO₄)₂, which was confirmed by a single-crystal X-ray structure determination of 2. The ESI mass spectral data of 1 and 2 display base peaks at m/z 391.25 and 405.27, respectively, which can be attributed to the complex cations [Cu(L1)(phen)]²⁺ and [Cu(L2)(phen)]²⁺, respectively. This is supported by the values of molar conductivity in DMF (Λ_M/Ω⁻¹ cm² mol⁻¹: 1, 203; 2, 205), which falls in the range⁵² for 1 : 2 electrolytes. The formation of the dicationic complexes was evidenced by the intense absorption of the ClO₄⁻ stretches at 1090 and 622 cm⁻¹ in the IR spectrum of the complexes. As expected, the band belonging to ν_imine(C=N) (L1, 1610; L2, 1611 cm⁻¹) was shifted to 1585 cm⁻¹ due to the coordination of imine nitrogen with copper(II). The sharp bands observed at 530–571 cm⁻¹ can be assigned to ν(Cu–N), probably originating from the coordination of L1 (1) or L2 (2) and phen (1 and 2) ligands.

Single crystal X-ray structure of [Cu(L2)(phen)](ClO₄)₂

The ORTEP representation of the complex, [Cu(L2)(phen)](ClO₄)₂2, including the coordinated atom numbering scheme, is shown in Fig. 1a. The crystal structure refinement data, selected bond lengths and bond angles are listed in Table S1 and Table 1, respectively. The asymmetric unit of 2 contains one dicationic complex molecule and two perchlorate anions. The copper atom in the complex is coordinated by all three nitrogen atoms (N1, N2, and N3) of tridentate ligand L2 and both nitrogen atoms (N4 and N5) of phen. The value of the structural index τ of 0.06 [τ = (β − α)/60, where α = N2–Cu1–N5 = 165.67° and β = N1–Cu1–N4 = 169.43°] reveals that the coordination geometry around copper(II) is best described as distorted square pyramidal.⁵³ The corners of the square plane of the geometry are occupied by N1 and N2 nitrogen atoms of the facially coordinated L2 and N4 and N5 nitrogen atoms of phen. The N3 nitrogen atom of L2 occupies the apical position at a distance (2.250(3) Å) longer than the equatorial nitrogen atoms (1.991–2.022 Å), as a consequence of the presence of two electrons in the d_z² orbital of copper(II). It is interesting to note that the τ value of 2 is very much lower than that of its dipica analogue [Cu(dipica)(phen)]²⁺ (τ, 0.77),⁵⁴ indicating that both the apical coordination of 6-methypyridine (mpy) nitrogen atom (N3) of L2 at a longer distance and the six-membered chelate ring formed by L2 decrease the steric congestion at copper(II) and change (i) the meridional fashion to facial and (ii) coordination geometry from trigonal bipyramidal towards square pyramidal. Additionally, the incorporation of the six-membered chelate ring and axial coordination of mpy facilitates stronger coordination of pyridyl N1 nitrogen atom (Cu–N_py: 2, 2.007(2); [Cu(dipica)(phen)]²⁺, 2.058(5) Å) and phen N4 and N5 nitrogen atoms (Cu–N_phen: 2, 2.022(2) and 2.012; [Cu(dipica)(phen)]²⁺, 2.157(5) Å). Notably, the incorporation of phen in the coordination sphere of 2 (Scheme 1) displaces the axially coordinated imine nitrogen of L2 (Scheme 1a) to a more strongly bound equatorial position (Scheme 1b; 1.991(3) Å) and decreases the steric congestion at copper(II) resulting in the distorted square pyramidal geometry, as revealed by the increase in trans bond angles (N1–Cu1–N4 = 169.43° and N2–Cu1–N5 = 165.67°). Another important structural observation is the inter-pair C–H⋯π anti-parallel non-covalent interactions (Fig. 1b) involving C(23)–H of the pyridine ring of phen and the centroid (C_g) of the mpy ring of L2 (C(23)–H⋯C_g(mpy) = 2.942 Å), which is probably responsible for the stabilization of the distorted square pyramidal geometry.⁵⁵

Fig. 1 (a) An ORTEP view of [Cu(L2)(phen)]²⁺2 with atom numbering of the complex and thermal ellipsoids at 40% probability. (b) Interpair C–H⋯π interaction in [Cu(L2)(phen)]²⁺2 involving the π-ring of the methyl pyridyl ring of L2 and the hydrogen atom [C(23)–H] of the pyridyl moiety of phen.

Table 1 Bond lengths (Å) and bond angles (°) for copper(II) complexes

	X-ray		DFT^a
	2	2	1
a Calculated from the B3LYP level of theory with mixed basis sets LANL2DZ/6-31G* using methanol as a solvent employing the CPCM method.
Cu(1)–N(1)	2.007(2)	2.067	2.132
Cu(1)–N(2)	1.991(3)	2.045	1.977
Cu(1)–N(3)	2.250(3)	2.358	2.224
Cu(1)–N(4)	2.022(2)	2.073	2.022
Cu(1)–N(5)	2.012(3)	2.075	2.236
N(5)–Cu(1)–N(1)	96.15(11)	97.28	95.34
N(5)–Cu(1)–N(2)	165.67(12)	166.31	171.67
N(5)–Cu(1)–N(3)	116.18(11)	116.95	104.48
N(5)–Cu(1)–N(4)	81.64(11)	80.96	79.44
N(1)–Cu(1)–N(2)	83.26(10)	81.58	78.77
N(1)–Cu(1)–N(3)	101.57(10)	101.93	149.54
N(1)–Cu(1)–N(4)	169.43(11)	163.71	103.76
N(2)–Cu(1)–N(3)	77.82(11)	76.49	78.78
N(2)–Cu(1)–N(4)	96.30(10)	96.28	107.56
N(3)–Cu(1)–N(4)	88.62(10)	93.19	102.47

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DFT-optimized structure and quantum chemical descriptors

The theoretical calculations help validate the experimental results and provide further insight into the complexes. In the present study, DFT calculations were incorporated to provide a comparison between experimental and theoretical values.⁵⁶ Geometry optimization for 1 and 2 in methanol was carried out by employing the B3LYP, 6-31G*/LANL2DZ levels, and the Gaussian 09 computer software.³⁶ The bond lengths, bond angles, optimized energies, frontier molecular orbital (FMO), and highest occupied molecular orbital (HOMO)–lowest unoccupied molecular orbital (LUMO) energy gap are also determined. The optimized structures of 1 and 2 (Fig. 2) in methanol show the donor sites N1, N2, and N3 of L1 and N5 of phen (1) or N1 and N2 of L2 and N4 and N5 of phen (2) coordinated to Cu(II) in the four corners of the square plane. Consequently, the N4 of phen (1) or the N3 of L2 (2) occupies the axial position in the coordination sphere of the Cu(II) ion with square pyramidal geometry. The geometrical parameters, like calculated bond lengths and bond angles, are in good agreement with the single-crystal X-ray data of 2 (Table 1). All strain energy minimized structures reproduced the observed X-ray structure to a maximum tolerance of 0.108 Å for 2 in the Cu–N bond lengths. The maximum deviation in the N–Cu–N bond angle for 2 is 5.72°. The small discrepancies in bond lengths and bond angles are attributable to non-covalent C–H⋯π inter-pair anti-parallel interactions within the lattice in the solid-state, which are not observed in the solution. Generally, in such instances, the normal discrepancy is higher than the mean between the observed and calculated structures,⁵⁷ but the average structure is sufficient to represent the observed structure.

Fig. 2 DFT-optimized structures and energy profile diagram of complexes [Cu(L1)(phen)]²⁺1 (a) and [Cu(L2)(phen)]²⁺2 (b). Doublet spin state: HOMO and LUMO in restricted spin calculations were carried out using the B3LYP level of theory with mixed basis sets LANL2DZ/6-31G* using methanol as a solvent employing the CPCM method.

Interestingly, the tridentate 3N L1 ligand coordinated meridionally, while L2 coordinated facially to copper(II) due to a rigid five-membered (L1) or flexible six-membered (L2) chelate ring. As a result, the geometry of the optimized structure in 1 differed from that in 2. Two longer estimated bond angles (1, N5–Cu–N2 and N1–Cu–N3; 2, N5–Cu–N2 and N1–Cu–N4) were used to determine the trigonality index (τ) for 1 (0.37) and 2 (0.04). The geometry of the copper center in 1 is severely distorted, while 2 is slightly distorted square pyramidal, and the τ for 2 is identical to the experimental result (τ, 0.06). In addition, the difference in the twist angle between the N–Cu–N (L1 or L2) and N–Cu–N (phen) planes estimated for 1 (1.3) and 2 (1.9) is due to the higher aromatic ring delocalization in the phen ring of 2 compared to the phen ring of 1, which enforces the planarity around the Cu(II) center in 2 because of the flexible six-membered chelate ring. This has beneficial effects on the DNA binding ability, mode of binding, and the DNA cleavage efficiency of the complex (cf. below).

The HOMO and LUMO are the main orbitals among FMOs that take part in chemical reactions. The HOMO acts as an electron donor, while the LUMO acts as an electron acceptor, and the gap between HOMO and LUMO characterizes the ability of electron transfer or chemical reactivity of the molecule.⁵⁸ It has been found that HOMO is largely distributed over the phen ligand in 1, on the metal, the methylpyridine (mpy) nitrogen atom of L2, and the phen ligand in 2. LUMO is distributed over the metal center, imine, pyridine (py), and mpy nitrogen atoms of L1 or L2 and phen nitrogen atoms in 1 or 2 (Fig. 2). The E_HOMO of 1 (−7.199) > 2 (−7.260) reveals that L1 releases more electrons; hence, these are involved in the strongest σ-bonding to Cu(II). Thus, the σ-donor capability of the L1 is tuned by the disparity in the planarity of phen because of the rigid 555 chelate rings in 1 rather than the flexible 655 chelate rings in 2, which reflects the presence of synergy between L1 or L2 and phen ligands when bound to the metal. The E_LUMO of 1 (−4.323) < 2 (−4.124) indicates that the π* orbital of phen in 2 is delocalized more than that of phen in 1. Therefore, the more delocalized π* orbital of phen in 2 is involved in a stronger π back-bonding with Cu(II) to stabilize the Cu(I) oxidation state more than that observed with the phen in 1. This is in good agreement with the observed trend in E_1/2 values and illustrates that the delocalization of the π* orbital of phen is important in stabilizing the lower oxidation of Cu(I) (cf. below). There is a significant energy difference between E_LUMO+1 and E_LUMO; LUMO has around 1.4 (1) or 1.5 (2) times the value of LUMO+1, indicating a high-energy transition from LUMO+1 to LUMO. However, E_HOMO−1 and E_HOMO values are comparable, indicating that these orbital transitions differ very little (Table 2).

Table 2 Calculated quantum chemical descriptors for the copper complexes using methanol as a solvent and the CPCM method at the B3LYP/LANL2DZ/6-31G* level of theory

Parameters	1	2	Cisplatin
a In eV. b In eV^−1.
Optimized energy (×10^4)^a	−3.905	−4.012
HOMO^a	−7.199	−7.260	−6.618
HOMO−1^a	−7.205	−7.270
LUMO^a	−4.323	−4.124	−1.970
LUMO+1^a	−3.110	−2.824
Energy gap, ΔE^a	2.877	3.136	4.647
IP^a	7.193	7.260	6.618
EA^a	4.323	4.124	1.970
η	1.438	1.568	2.324
σ	0.695	0.637	0.430
μ	−5.761	−5.692	−4.294
S	0.347	0.319	0.215
ω	11.537	10.330	3.967
χ	5.761	5.692	4.294
N	0.086	0.096	0.252
ΔNmax	4.006	3.630	1.848

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In most stable compounds, the HOMO–LUMO gap, viz. 1 (2.877 eV) < 2 (3.136 eV), defines the properties of the electrons and explains the eventual charge transfer interactions taking place within the molecules. This reflects the ability of electrons to move from occupied orbitals to unoccupied orbitals, thereby providing a fundamental basis for judging the chemical reactivity, selectivity, and stability of the compounds. As a result, the energies of the FMOs were examined to study some quantum chemical descriptors,^59–63 such as ionization potential (IP = −E_HOMO), electron affinity (EA = −E_LUMO), energy gap (ΔE = E_LUMO − E_HOMO), chemical hardness (η = E_LUMO − E_HOMO/2), absolute softness (σ = 1/η), global softness (S = 1/2η), electrochemical potential (μ = −(IP + EA)/2), electrophilicity index (ω = μ²/2η), nucleophilicity index (N = 1/ω), additional electronic charges (ΔN_max = σ/η), and electronegativity (χ = −μ) during excitation of 1 and 2. The quantum chemical descriptors of 1 and 2 are compared with the special parameters of cisplatin⁶⁴ (Table 2) to determine the anticancer properties. A high E_HOMO denotes the donation of an electron by a suitable molecule, while a low E_LUMO implies electron acceptance. As E_HOMO increases and E_LUMO decreases, this indicates an increase in biological activity. Electron freedom, or the energy gap between HOMO and LUMO, is crucial in determining biological reactivity since activity increases as ΔE decreases. The ΔE of soft molecules (σ) is small, while that of hard molecules (η) is large. Consequently, soft molecules have more biological activity than hard molecules. A molecule can readily donate electrons to coordinate the proper structure if the global electronegativity (χ) is small or the chemical potential (μ) is large. This indicates that the electron delocalizes the entire structure. As the nucleophilicity index (N) increases and the electrophilicity index (ω) decreases, biological activity also increases. The greater the ΔN_max value of a compound, the greater its biological activity. The increasing biological activity of the compound is implied by its increasing global softness (S) value. The computed values indicate the possible biological activity of 1 and 2, and the most advantageous parameters are ΔE, σ, η, ΔN_max, and S. For the present copper(II) complexes, their anticancer activities are greater than those of cisplatin. It is observed from the above parameters that they are good candidates for cancer drugs.

Electronic and EPR spectral and redox properties

The electronic spectra of 1 and 2 are very similar to each other (Fig. S1 and S2) and show a broad low energy ligand field (LF) band (λ_max: 1, 684; 2, 675 nm), followed by a shoulder (λ_max: 1, 896; 2, 856) in the visible region revealing a square-derived coordination geometry around Cu(II) with slight distortion.⁶⁵ The observed molar absorptivity of 2 (ε_max, 280 M⁻¹ cm⁻¹) is higher than that of 1 (ε_max, 100 M⁻¹ cm⁻¹) due to the presence of a flexible six-membered chelate ring, which increases the distortion in square-based Cu(II) coordination geometry of 2. The intense absorption bands observed in the range of 268–323 nm are attributed to the intraligand π → π* transitions.

The polycrystalline EPR spectra of 1 and 2 are isotropic (Fig. S3 and S4). The frozen solution EPR spectra of both the complexes exhibit spectral features typical of mononuclear Cu(II) species (Fig. S5 and S6). They are axial with g_∥ > g_⊥ > 2.0 and G = [(g_∥ − 2)/(g_⊥ − 2)] = 5.2–5.5, suggesting the presence of a d_x²−y² ground state in copper(II) located in square-based geometries.⁶⁶ A square-based CuN₄ chromophore is expected^67,68 to show a g_∥ value of 2.200 and A_∥ value in the range of 180–200 × 10⁻⁴ cm⁻¹, and the distortion or incorporation of strong axial interaction is expected to enhance the g_∥ value and decrease the A_∥ value.^67,68 Therefore, the observed values of g_∥ (2.233) and A_∥ (188–189 × 10⁻⁴ cm⁻¹) are consistent with the CuN₅ chromophore of the complexes in the solution. The g_∥/A_∥ values (118–119 cm) fall within the range (105–135 cm) expected for complexes with square planar geometry,⁶⁹ suggesting that there is a slight distortion from planarity, which is evident from the X-ray crystal structure of 2.

The cyclic (CV) and differential pulse voltammetric (DPV) techniques were employed to investigate the redox behavior of 1 and 2 in the DMF solution on stationary glassy carbon electrodes (Fig. S7 and S8). They display well-defined cathodic and anodic waves revealing the reversible Cu^II/Cu^I redox couple owing to the linearity of the plots of i_pcversus ν^1/2 (D: 1, 7.7; 2, 8.2 × 10⁻⁶ cm² s⁻¹) and the value of the peak current ratio (i_pa/i_pc: 1, 0.7; 2, 0.9). However, the value of the difference in peak potentials (ΔE_p: 1, 100; 2, 86 mV) is close to the Fc/Fc⁺ couple (ΔE_p, 86 mV; E_1/2, 0.406 V vs. SCE) under similar experimental conditions. Interestingly, the less negative redox potential, E_1/2 of 2, is higher, and peak potential separation, ΔE_p, is lower than that of 1 (E_1/2 and ΔE_p: 1, −0.077 V and 100 mV; 2, −0.072 V and 86 mV) for the Cu^II/Cu^I couple. This indicates that the smallest structural reorganization between copper(II) and copper(I) species renders the Cu(II) to Cu(I) electron transfer more facile. This is due to the presence of a flexible six-membered chelate ring forming a boat conformation, which is attracted by the steric hindrance between the 6-methyl group in the pyridine moiety of L2 and phen ligands, thereby distorting the Cu(II) coordination geometry.⁷⁰ The redox potentials of the Cu^II/Cu^I couple from the DPV (Fig. S9 and S10) are −0.063 V (1) and −0.059 V (2) vs. SCE.

Stability studies

Both complexes are soluble in DMSO and 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer (pH 7.1), stable in air, and non-hygroscopic. The complexes in DMSO, buffer, and cell growth medium (DMEM) were subjected to UV-visible spectral analyses at various time intervals (0, 24, and 48 h). After 0, 24, and 48 h in solution, no new absorption peaks emerged for 1 and 2, and there was no discernible red or blue shift in the position of the absorption peaks (Fig. S11 and S12). However, after 24 h, the intensity of the absorption peaks for 1 and 2 decreased, indicating a decrease in the concentration. Consequently, the findings show that the structures of 1 and 2 did not change significantly in DMSO, buffer, or DMEM, which establishes the basis to conduct further biological studies.

DNA binding properties

Upon incremental addition of CT DNA to 1 (Fig. 3) and 2 (Fig. S13), the ligand-centered band (268 nm) shows a decrease in molar absorptivity (hypochromism: 1, ∼60; 2, ∼32%) with red shift (1, 3; 2, 2 nm) at R = 25 (R = [DNA]/[Cu]). The observed hypochromic effect, accompanied by a slight red shift, strongly indicates the binding of 1 and 2 to CT DNA through a partial intercalative interaction (Table S2).⁷¹ The intrinsic binding constant (K_b) value (Fig. 3 and Fig. S13) demonstrates the strong DNA binding affinity of the complexes and follows the order 1 (1.36 × 10⁵ M⁻¹) > 2 (1.06 × 10⁵ M⁻¹). Compared to the partially intercalating metal complex {[Ru(bipy)₂(dppz)]²⁺ (K_b, 1.35 × 10⁶ M⁻¹)}, the K_b values of 1 and 2 are 10 times lower.⁷² However, relative to the previously published ternary copper(II) complexes of phen bases, [Cu(pmdt)(phen)](ClO₄)₂ (K_b, 3.1 × 10³ M⁻¹)⁷³ and [Cu(dipica)(phen)](ClO₄)₂ (K_b, 2.7 × 10³ M⁻¹),⁷⁴ the noted DNA binding affinity of these complexes is 100 times greater. In contrast, 1 and 2 exhibit comparable DNA binding affinities when compared to the established intercalator (EthBr, K_b, 4.94 × 10⁵ M⁻¹)⁷² and mixed ligand copper(II) complexes that contain phen, [Cu(cur)(phen)(ClO₄)] (K_b, 7.69 × 10⁵ M⁻¹) and [Cu(acac)(phen)(ClO₄)] (K_b, 3.18 × 10⁵ M⁻¹).⁷⁵ The higher binding strength in 1 is attributed to the presence of a rigid five-membered chelate ring. The rigidity enhances the partial intercalative interaction of the phen ligand between the base pairs of DNA. The hydrophobic interaction of the 6-methyl group with the hydrophobic DNA surface further supports an increase in the DNA binding affinity^56,76 in 1. The lower binding affinity of 2 is due to the presence of a flexible six-membered chelate ring. The flexibility reduces the efficiency of the partial intercalative interaction of the phen ligand, and the decrease in the hydrophobic interaction of the 6-methyl group in 2.

Fig. 3 Absorption spectra of 1 (10 × 10⁻⁶ M) in 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1 in the absence (R = 0) and presence (R = 25) of increasing amounts of CT DNA. Inset: plot of [DNA] vs. [DNA]/(ε_a − ε_f) at R = 25 of 1.

The fluorescence intensity of EthBr bound to DNA at 594 nm (λ_ex, 520 nm) shows a remarkable decrease (Fig. S14 and S15) with the addition of 1 (78%) or 2 (77%), indicating that DNA-bound EthBr molecules are released into solution.⁷⁷ Such fluorescence quenching may be caused by the strong partial intercalative interaction of 1 or 2 (cf. above), leading to the displacement of DNA-bound EthBr. The slope of the plot of I₀/I versus [complex] gives K_SV (Fig. S14 and S15, inset). The fluorescence quenching follows Stern–Volmer behavior (K_SV: 1, 4.26; 2, 4.12 × 10⁵ M⁻¹). The apparent DNA binding constant⁷⁸ values (K_app: 1, 5.20; 2, 5.00 × 10⁶ M⁻¹) suggest that 1 intercalates more strongly than 2. Compared to copper(II) complexes based on phen such as [Cu(L)(phen)](ClO₄)₂ (K_app, ∼10⁵ M⁻¹),⁷⁹ [Cu(L1–L5)(phen)-(H₂O)](ClO₄)₂ (K_app, 3.60–4.18 × 10⁵ M⁻¹),⁸⁰ [Cu(phdpa)(phen)-(H₂O)](ClO₄)₂ (K_app, 9.59 × 10⁴ M⁻¹)⁸¹ and [Cu(ETDPA)-(phen)](ClO₄)₂ (K_app, 3.40 × 10³ M⁻¹),⁸² the K_app values of 1 and 2 are 10–1000 times higher.

The observed circular dichroic (CD) spectrum of CT DNA (2 × 10⁻⁵ M) consists of a positive band (270 nm) due to base stacking and a negative band (240 nm) due to helicity (Fig. S16 and S17), which is characteristic of DNA in the right-handed B form.^56,83 When 1 or 2 is incubated with DNA at the 1/R (= [Cu complex]/[DNA]) value of 3, the CD spectrum of DNA (Fig. S16 and S17) changes both positive (vast increase in intensity; red-shift: 1 and 2, 3 nm) and negative bands (slight decrease in intensity; red-shift: 1 and 2, 4 nm), which is consistent with partial intercalative interaction⁸⁴ of phen ring.

Both complexes display a well-defined cathodic and anodic wave within the time scale of the cyclic voltammetry (CV) experiments (2% DMF/5 mM Tris-HCl/50 mM NaCl buffer, pH 7.1). Redox-active complexes 1 and 2 show the reversible Cu(II)/Cu(I) couple (E_1/2: 1, −0.079; 2, −0.073 V/ΔE_p: 1, 99; 2, 89 mV/i_pa/i_pc: 1, 0.8; 2, 0.6). Upon the addition of CT DNA (R = [DNA]/[Cu] = 5), a significant reduction in cathodic and a slight decrease in anodic peak currents (Fig. S18 and S19) are observed due to the slow diffusion of the DNA-bound complex to the electrode surface.⁸⁵ The observed redox potential (E_1/2) shift for 1 is less positive (+2 mV) and for 2 is less negative (−4 mV), which implies that the binding of 1 and 2 to DNA is through partial intercalative interaction. Additionally, the E_1/2 values from differential pulse voltammetry (DPV: 1, −0.063; 2, −0.058 V) for the Cu(II)–Cu(I) redox couple of the complexes (Fig. S20 and S21) are shifted (1, +1; 2, −6 mV), revealing a stronger DNA binding affinity. The shift in the value of the formal potential (ΔE°′) can be used to estimate the ratio of the equilibrium binding constant (K⁺/K²⁺) values of the Cu(I) and Cu(II) forms to DNA, which are calculated to be 1.0 (1) and 1.3 (2), respectively, suggesting their stabilization in the Cu(I) oxidation state upon binding to CT DNA.⁸⁶

DNA cleavage properties

Upon incubation of the pUC19 plasmid DNA (20 μM in base pairs) and 1 or 2 (10–100 μM) in the absence of external additives for 1 h at 37 °C in 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer (pH = 7.1), followed by electrophoresis, the fastest migration is observed for the SC DNA (form I). A single strand of SC DNA relaxes upon cleavage, resulting in slower-moving nicked circular (NC) DNA (form II; 43%), and no linear DNA (LC, form III) is discernible, indicating that neither of the DNA strands is cleaved by 1 or 2 (Fig. S22). The amount of form I decreases whereas that of form II increases when the concentrations of 1 or 2 are increased (100–160 μM) while keeping the DNA concentration constant (20 μM), causing the complete conversion to form II [97%; conc. 140 (1) or 130 μM (2)], which demonstrates that the cleavage is concentration-dependent. Due to the absence of form III in either 1 or 2, single-strand DNA scission was interpreted (Fig. 4).⁸⁷

Fig. 4 Agarose gel showing cleavage of 20 μM SC pUC19 DNA incubated with 1 (top) or 2 (bottom) in a 2% DMF 50 mM Tris-HCl/NaCl buffer (pH 7.1) at 37 °C for 1 h. Lane 1, DNA and lane 2–8, DNA + 1 (top) or 2 (bottom) (100, 110, 120, 130, 140, 150, and 160 μM). Forms I and II are supercoiled and nicked circular forms of DNA, respectively.

The time-dependent cleavage of DNA by 1 or 2 was investigated {[complex] = 140 (1) or 130 (2) μM and [DNA] = 20 μM in bp} as a function of time (0–60 minutes). The amount of NC DNA increases as the reaction time is extended, while the amount of SC DNA steadily decreases (Fig. 5 and S23), and the rate of DNA cleavage reaches its maximum within 55 minutes. The appearance of NC DNA and the disappearance of SC DNA follow pseudo-first-order kinetic profiles and fit well into a single-exponential decay curve (Fig. 5 and S23). The copper(II) center may be crucial in fostering Lewis acid–base activity, which would increase the rate at which DNA cleaves.

Fig. 5 (a) Agarose gel showing cleavage of 140 μM SC pUC19 DNA incubated with 1 (140 μM) in a 2% DMF 50 mM Tris-HCl/NaCl buffer (pH 7.1) at 37 °C for 1 h. Lane C, DNA and lane 1–13, DNA + 1 (0, 5, 10, 15, 20, 25, 30, 35, 40, 45, 50, 55, and 60 min). Forms I and II are supercoiled and nicked circular forms of DNA, respectively. (b) Disappearance of the supercoiled form (SC DNA) and formation of the nicked circular form. Inset: (% SC DNA) versus time for 1 (140 μM).

When supercoiled pUC19 DNA was treated with methyl green, the cleavage reaction mediated by the complexes was not quenched (Fig. S24, lane 7), while it was completely cleaved to form II. This suggests that the complexes prefer to bind through partial intercalation rather than groove binding (cf. above).⁸⁸ The mechanism by which 1 or 2 cleaves pUC19 DNA was examined (Fig. S24) and explained in the presence of singlet oxygen quencher NaN₃ (100 μM), hydroxyl radical scavenger DMSO (20 μM), superoxide anion generator SOD (0.5 units), and H₂O₂ producer catalase (0.5 units) in an aerobic environment.⁸⁹ As illustrated in Fig. S24, the mechanism of DNA cleavage by 1 or 2 is as follows: DNA cleavage activity is not affected by NaN₃, DMSO, SOD, or catalase (lanes 3–6), ruling out the potential of cleavage by singlet oxygen, hydroxyl radical, superoxide, and H₂O₂, respectively.

This finding suggests that a hydrolytic mechanism may be the cause of the DNA cleavage response by 1 or 2. In general, two main processes are involved in DNA cleavage: oxidative cleavage and hydrolytic cleavage. In the presence of external reducing agents, the mixed-ligand Cu(II) complexes, including phen as a coligand with the CuN₅ chromophore, like [Cu(L)(phen)](ClO₄)₂ ⁸¹ and [Cu(dipica)(phen)](ClO₄)₂,⁵⁵ cleave plasmid DNA oxidatively. In most cases, this involves a photoactive or redox-active metal center and produces reactive singlet oxygen or hydroxyl radical species. This damages the sugar and/or base and produces fragmented species that are not religated. There are no disadvantages to hydrolytic cleavage without external additives because it is possible to religate the cleaved products enzymatically. Therefore, to ascertain the hydrolytic nature of the cleavage reaction, the NC form obtained from the cleavage of SC DNA was reacted with a T4 ligase enzyme.^90,91 We observed 90% conversion of the NC DNA to its original SC form (Fig. 6, lanes 3 and 5), providing direct evidence in favor of the hydrolytic mechanism. Consequently, the hydrophobic interaction of the 6-methyl moiety supports the partial intercalation of planar phen with rigid 555 (1) or flexible 556 chelate rings (2). This leads to the distortion of the DNA double helix due to the increased DNA binding of 2 compared to 1, resulting in significant hydrolytic DNA cleavage activity⁹² at lower concentrations of 2 (130 μM) rather than 1 (140 μM).

Fig. 6 Analysis of the capacity of T4 DNA ligase to religate DNA cleaved by complexes. Lane 1, DNA control. Lane 2: products (NC pUC19) obtained from the reaction with 1; lane 3, lane 2 + T4 ligase; lane 4, products (NC pUC19) obtained from the reaction with 2; and lane 5, lane 4 + T4 ligase.

BSA interaction

Tryptophan emission⁹³ quenching experiments were carried out to measure the protein binding affinity of complexes by adding increasing concentrations (0–4 mM) of complexes to BSA (1 mM). At 300 K, the fluorescence intensity of BSA decreases to 63 (1), 62% (2), and at 310 K, it decreases to 69% (1, 2), with a red shift of 4–10 nm (Fig. 7 and S25). The observed Stern–Volmer plots (Fig. S26–S28) show a good linear relationship⁹⁴ in which K_SV (Table S3) increases as temperature rises (300 K: 1, 4.31; 2, 4.08 × 10⁵ M⁻¹ and 310 K: 1, 5.43; 2, 5.62 × 10⁵ M⁻¹), indicating that the most likely source of fluorescence quenching is a dynamic quenching mechanism. The obtained bimolecular quenching rate constant (k_q) is in the order of 10¹³ M⁻¹ s⁻¹, which is 1000 times greater than the diffusion-controlled quenching maximum limit (2.0 × 10¹⁰ M⁻¹ s⁻¹).⁹⁵ Moreover, when the temperature is increased (300 K, 1, 1.50; 2, 3.23 × 10⁵ M⁻¹ and 310 K, 1, 2.26; 2, 3.88 × 10⁵ M⁻¹), the effective quenching constant, K_a (Table S3), derived using the modified Stern–Volmer equation,⁹⁶ increases.

Fig. 7 Changes in the fluorescence spectra of BSA through titration with 1 at 300 K (left, A) and 310 K (right, B). The concentration of BSA was 1 × 10⁻⁶ mol L⁻¹, and the concentration of 1 was varied from (a) 0.0 to (k) 4.0 × 10⁻⁶ mol L⁻¹, with pH 7.4 and λ_ex 280 nm.

When the complexes are added, the 210 nm absorbance peak of BSA⁹⁷ is dramatically reduced, with a red-shift (1, from 210 to 220 nm; 2, from 210 to 222 nm). Meanwhile, the absorption intensity at 280 nm has slightly increased (Fig. 8), indicating that additional aromatic acid residues have extended into the aqueous environment and destroyed the tertiary structure of BSA.⁹⁸ These findings indicate that the interaction between copper(II) complexes and BSA is primarily a static quenching process caused by the formation of a ground-state complex (BSA-1 or BSA-2). The binding constant, K_b, increases with temperature (300 K: 1, 1.10; 2, 2.19 × 10⁶ M⁻¹ and 310 K: 1, 1.34; 2, 2.62 × 10⁶ M⁻¹), indicating that a stable BSA-1 or BSA-2 ground-state complex is formed⁹⁹ (Table S3). Additionally, the value of n, which is close to 1 (Table S3), shows that the binding site in BSA is unique and that the complexes have easy access to the hydrophobic environment of the tryptophan residue.¹⁰⁰ Since the values of ΔG° are negative, it is clear that the binding process is spontaneous, as shown in Table S3. The presence of significant hydrophobic forces of interaction between the ligands of copper(II) complexes and BSA is also supported by the positive values reported for both ΔH° and ΔS°.¹⁰¹ Covalent bonding could be considered in addition to the hydrophobic interaction. However, the obtained value for ΔH° (1, 70; 2, 74 kJ mol⁻¹) is significantly lower than what would be predicted for the formation of covalent bonds, which should be ≥120 kJ mol⁻¹.¹⁰²

Fig. 8 UV-Vis absorption spectra of BSA in the absence and presence of 1 (left, A) and 2 (right, B). (a) Absorption spectrum of BSA. (b) Absorption spectrum of BSA in the presence of 1 and 2 at the same concentration, [BSA] = [Cu complex] = 3.5 × 10⁻⁶ mol L⁻¹.

Cytotoxicity of copper(II) complexes

The antiproliferative activity in vitro against human cervical carcinoma (HeLa) and human lung epithelial adenocarcinoma (A549) cell lines, mouse embryonic fibroblasts (NIH 3T3) and human lung epithelial (L132) normal cells was carried out for 1, 2, cisplatin, free ligands, and copper-acetate using the MTT assay method. The activities of 1 and 2 are much higher than those of the corresponding free ligands in HeLa or A549 cells. They have exhibited a significant inhibitory potency against the proliferation of the HeLa (IC₅₀: 24 h, 1.164 ± 0.085 (1); 1.372 ± 0.104 μM (2) and 48 h, 0.853 ± 0.046 (1); 1.023 ± 0.078 μM (2)) or A549 (IC₅₀: 24 h, 0.496 ± 0.043 (1); 0.494 ± 0.041 μM (2) and 48 h, 0.199 ± 0.015 (1); 0.201 ± 0.016 μM (2)) cell line at a very low concentration in dose- and time-dependent manners. Since the phen co-ligand possesses higher lipophilicity¹⁰³ to penetrate the cell membrane and act as a recognition element for non-covalent interaction with DNA, the IC₅₀ values of 1 and 2 against the A549 cell line are marginally lower (∼2 to 3 times) than those of copper(II)-phen complexes [(phdpa)Cu(phen)(H₂O)](ClO₄)₂ (IC₅₀, 3.10 μM),⁸⁴ [Cu(L)(phen)], and [Cu(sal)(phen)] (IC₅₀, 1.42 μM).⁸³ They are lower (∼6 to 16 times) than those of [(phdpa)Cu(phen)(H₂O)](ClO₄)₂ (IC₅₀, 3.10 μM),⁸⁴ [Cu(L)(phen)](ClO₄)₂ (IC₅₀, 3.55 μM),¹⁰⁴ [Cu(phen)₂(CH₃CN)](ClO₄)₂ (IC₅₀, 4.00 μM),¹⁰⁵ [Cu(Aldpa)(phen)](ClO₄)₂ (IC₅₀, 7.60 μM),¹⁰⁶ [Cu(pabt)(phen)]ClO₄ (IC₅₀, 7.60 μM)¹⁰⁷ and [Cu(ETDPA)(phen)](ClO₄)₂ (IC₅₀, 8.00 μM).¹⁰⁸ Interestingly, compared to [Cu(L1–L5)(phen)(H₂O)](ClO₄)₂ (IC₅₀, 3.85–46.75 μM),⁸² these complexes are likely to be significantly lower (∼8 to 95 times).⁸² They are remarkably much lower (∼170 to 200 times) than those of [Cu(cur)(phen)(ClO₄)] (IC₅₀, 85.30 μM) and [Cu(acac)(phen)(ClO₄)] (IC₅₀, >100 μM).⁷⁷ Further, the cytotoxicity of 1 and 2 against the HeLa and A549 cancer cell lines was comparable to that of the prevalent benchmark metallodrug, cisplatin, under the same experimental conditions. The IC₅₀ values of 1 and 2 were ∼90 (HeLa) or ∼200 (A549) times lower than that of cisplatin (HeLa: 48 h, IC₅₀ = 16.4 μM; A549: 48 h, IC₅₀ = 40 μM),^117,118 which is certainly a beneficial factor for considering these complexes as promising candidates for new antitumor agents. In addition, 1 and 2 do not cause any damage to the normal mouse embryonic fibroblast cells (NIH 3T3: 48 h, IC₅₀ > 500 μM) or normal human lung epithelial cells (L132: 48 h, IC₅₀ > 250 μM), indicating that they are non-toxic to healthy cells, which is expected for a better drug.

Reactive oxygen species (ROS) detection

The ROS levels in the A549 cancer cell line are assessed in this study by employing an oxidant-sensitive fluorescent dye. Cellular esterase can alter the non-polar fluorescent dye dichlorodihydrofluorescein diacetate (DCFH-DA) into the polar DCFH. However, this DCFH is not visible under a microscope and requires intracellular ROS to change into a fluorescent DCF. Our findings propose that complexes treated with A549 exhibit fluorescence rather than the absence of fluorescence observed in the control cells (Fig. 9). This suggests that both complexes accelerate the apoptosis caused by ROS in A549 cells.¹⁰⁹

Fig. 9 Reactive oxygen species (ROS) generation in the absence (C) and presence of Cu(II) complexes (1 and 2) and visualized using a fluorescence microscope.

Further, we assessed DCF fluorescence intensity using flow cytometry to investigate the influence of 1 or 2 on ROS levels. The DCF fluorescence intensity is 100% in the control. Hydrogen peroxide (H₂O₂) is next to the superoxide anion and hydroxyl radical, a key member of the class of ROS. It is an important ROS produced by the mitochondrial respiratory chain in many chemically reactive molecules derived from oxygen;¹¹⁰ hence, it is used as a positive control. The A549 cells are treated at 250 μM concentrations of H₂O₂, indicating a higher DCF fluorescence intensity of 149.12 ± 0.49% at the cellular level of ROS. The DCF fluorescence intensity increases to 140.34 ± 0.93% (1) or 148.42 ± 0.62% (2) when A549 cells are treated with 1 or 2. A greater shift denotes higher fluorescence intensity due to the formation of more DCF, resulting in the production of more ROS for 2 than for 1, which is comparable to the DCF fluorescence intensity of H₂O₂ ROS (Fig. 10). This demonstrates that 1 and 2 activate the ROS-generating machinery and lead to increased levels of ROS when exposed to cancer cells, causing apoptosis. Therefore, it is clear that oxidative stress contributes to the effective anticancer properties of Cu(II) complexes.

Fig. 10 (a) DCFDA assay in A549 cells for the generation of ROS using the copper(II) complexes in a time-dependent manner (control, black; complex, red; and H₂O₂, blue) and (b) bar diagram showing % ROS generation obtained from the FACSverse analysis.

Cell cycle arrest studies

By inducing apoptosis and/or cell cycle arrest, which are important mechanisms of action for anticancer metal complexes, cytotoxic drugs may prevent the growth of cancer cells.¹¹¹ This was clarified using flow cytometry to examine the profiles of PI-stained A549 cells treated with IC₅₀ doses of 1 or 2 for 24 h.¹¹² It is possible to quantify the populations in the G0/G1, S, and G2/M phases, which can be connected to the quantity of DNA copies in the single-cell suspension. The percentage of control cells at S phase is 18.48 ± 0.34% (Fig. 11). When exposed to synchronized cells in the S phase, 1 (28.76% ± 0.29%) or 2 (30.91% ± 0.40%) started a startling rise of 10.28% (1) or 12.43% (2). Both the G0/G1 and G2/M phases decreased in tandem with the increase in the S phase. The S phase, which occurs before cell division, is when histone and DNA synthesis occur.¹¹³

Fig. 11 (a) Flow cytometric analysis showing the S phase cell cycle arrest by copper(II) complexes in A549 cells and (b) bar diagram showing % S phase arrest obtained from the FACSverse analysis.

However, DNA replication is halted if DNA breaks occur during the process. According to the flow cytometry data, 1 and 2 cause DNA damage that leads to A549 cells undergoing apoptosis, which arrests the cells in the S-phase. It is clear that, under the same conditions, 2 produces a greater apoptotic effect on cell cycle arrest than 1 (Fig. 11). The impact on cell cycle arrest does not match the IC₅₀ values of the complexes. Given that A549 cancer cells have shorter doubling times than healthy cells, S-phase cell cycle arrest in cancer cells may help deliver differential toxicity. Numerous anticancer drugs, such as cisplatin,¹¹⁴ mitomycin C,¹¹⁵ hydroxyurea,¹¹⁶ and the polyphenolic anticancer agent resveratrol,¹¹⁷ have been shown to cause S-phase arrest in various human cancer cell types. Numerous anticancer drugs are known to suppress the activity of topoisomerase and other essential enzymes involved in DNA duplication.¹¹⁸ Further study into the processes underlying the S-phase arrest brought on by 1 or 2 will be intriguing.

Apoptosis studies

To determine whether the inhibition of cell proliferation and cytotoxicity is caused by apoptosis induction or nonspecific necrosis, the most active cytotoxic complexes 1 and 2 were examined. Apoptosis, also known as programmed cell death, is a type of cell death that is considered when developing anticancer drugs. In order to evaluate the anticancer drug, an apoptosis study that uses acridine orange (AO) and ethidium bromide (EB) staining is introduced. Although EB only stains cells that have lost their membrane integrity, AO is a vital dye that can stain both live and dead cells.¹¹⁹ Viable cells have green fluorescing nuclei with a highly organized structure when viewed under a fluorescence microscope. Early apoptotic cells also have green fluorescing nuclei, but perinuclear chromatin condensation is visible as bright green patches or fragments.

Late apoptotic cells have orange to red fluorescing nuclei with condensed or fragmented chromatin. Necrotic cells have consistently orange to red fluorescing nuclei and have expanded to large sizes; there is no sign of chromatin fragmentation. The complexes promoted the apoptosis of A549 cells, as evidenced by the microphotographs of AO- and EB-stained A549 cells pretreated for 24 hours with the copper(II) complexes. Apoptosis, which is characterized by extremely condensed chromatin that marinates into a horseshoe-shaped structure, can be induced by both complexes (Fig. 12). As a result, the observed morphological changes demonstrate that while both complexes cause cell death, they induce only apoptosis and not necrosis.¹²⁰

Fig. 12 A549 cells stained with acridine orange (AO)/ethidium bromide (EB) in the absence (C) and presence of copper(II) complexes (1 or 2, 0.50 μM) and visualized under a fluorescence microscope.

4′,6′-Diamidino-2-phenylindole (DAPI) staining was also performed to help establish the apoptotic phase. The copper(II) complexes treated with A549 cells and the control were stained with DAPI and observed under a confocal microscope (Fig. 13). In contrast to the treated cells, which exhibit typical indicators of cells undergoing apoptosis, the nuclei of the control cells are permeabilized with detergent (0.1% Triton X-100) and are light and uniformly marked. The treated cells are found to have fragmented or highly condensed nuclei; bright field images indicate cell shrinkage and membrane blebbing, which are characteristics of apoptotic cells. Therefore, the DAPI staining with both complexes¹²¹ shows an apoptotic mode of cell death.

Fig. 13 A549 cells stained with DAPI in the absence (C) and presence of copper(II) complexes (1 or 2, 0.50 μM) and visualized under a fluorescence microscope.

Annexin V and propidium iodide (PI) assays, along with flow cytometry, were used to examine the induction of apoptosis in A549 cells. The IC₅₀ concentrations determined from the cytotoxicity assessment were used to measure the induction of apoptosis after 24 h of treatment with 1 and 2. The assay of annexin V and PI selectively labels cells, allowing for the differentiation of live, apoptotic, and necrotic cells. Annexin V binds to phosphatidylserines externalized to the outer leaflet of the plasma membrane during apoptosis, while PI, a membrane-impermeable dye, binds to the DNA of necrotic cells with damaged plasma membranes.¹²² The analysis of the flow cytometry data indicates (Fig. 14) that the viable cells (annexin V−/PI− cells) in the lower left quadrant comprise 99.67% of the total; the viable cells in the treated ones decrease to 66.82 ± 0.52% (1) or 61.61 ± 0.48% (2). The early apoptotic cells (annexin V+/PI− cells) in the lower right quadrant increased from 0.07% in the untreated cells to 28.67% ± 0.25% (1) or 35.99% ± 0.21% (2), while the late apoptotic cells (annexin V+/PI+) in the upper right quadrant abruptly increased from 0.09% in the untreated cells to 4.29% ± 0.07% (1) or 2.37% ± 0.05% (2) in the treated cells.

Fig. 14 (a) Cellular apoptosis induced by copper(II) complexes as determined from the annexin V-FITC/PI staining assay of the A549 cells with four distinct phenotypes: viable cells (lower left quadrant, LL); cells at an early stage of apoptosis (lower right quadrant, LR); cells at a late stage of apoptosis (upper right quadrant, UR); and necrosis (upper left quadrant, UL) and (b) bar diagram showing % early apoptosis obtained from the FACSverse analysis.

The absence of necrotic cells is shown by the dearth of such a large fluctuation [untreated, 0.17%; treated, 0.22 ± 0.015% (1) or 0.03 ± 0.001% (2)] in the upper left quadrant (annexin V−/PI+ cells). The overall percentages of apoptotic cells in 1 and 2 treated cells at IC₅₀ concentration were 33 and 38%, respectively. This indicates that 2 is more potent than 1 at inducing apoptosis in A549 cells. The fact that the lower right quadrant comprised the highest percentage of cells made it clear that both complexes caused early apoptosis in A549 cells. One of the primary mechanisms of action for copper complexes is generally acknowledged to be their capacity to induce apoptosis, which has been extensively reported in the literature.^27,123 These findings demonstrated that 1 or 2 produced apoptosis alone in treated A549 cells in contrast to the control, suggesting that 1 or 2 induced toxicity involving apoptotic cell death.^124,125

Conclusions

Novel mixed-ligand copper(II) complexes of the type [Cu(L1)/(L2)(phen)](ClO₄)₂ are synthesized and characterized. The geometry of [Cu(L2)(phen)](ClO₄)₂ was analyzed through single-crystal X-ray diffraction and was shown to have a CuN₅ chromophore. The coordination environment of the Cu(II) atom can be described as a distorted square pyramidal. A three-dimensional infinite network is formed and dominated by the inter-pair C–H⋯π anti-parallel non-covalent interactions. The chemical reactivity, stability, and selectivity of both complexes in the solution were understood using electronic parameter calculations and structural optimization using DFT. The optimized structures show that the coordination geometry of the copper center is a distorted square pyramidal. They are promising candidates for cancer drugs corresponding to the computed quantum chemical descriptors. Absorption, EPR, and ESI-MS spectral investigations revealed that both complexes preserve their geometry in solution. The binding properties of the complexes with CT DNA and BSA are investigated. The results of electronic absorption and circular dichroic spectroscopy, as well as electrochemistry, indicate that both complexes bind to DNA in a partial intercalative mode with a high affinity for DNA. The binding studies with EthBr have revealed a significant decrease in the fluorescence intensity of the EthBr–DNA system in the presence of both complexes, implying that the complexes can compete for DNA-binding sites with EthBr and displace EthBr from the EthBr–DNA system. Interestingly, both complexes show reversible redox behavior. They cause a considerable decrease in current when they bind to CT DNA, shift to a less negative or less positive Cu(II)/Cu(I) redox potential, and stabilize the Cu(I) oxidation state, resulting in more evident chemical nuclease activity. The mechanism of quenching in BSA is believed to occur through the static mode, implying that complexes bind to BSA via hydrophobic interaction based on protein binding studies. Both complexes show hydrolytic DNA cleavage activity in the absence of an activator (97%, 130–140 μM complex concentration). Their affinity for BSA (K_b, 10⁶ M⁻¹) shows that the complexes are delivered in the blood by serum albumin. The two copper(II) complexes are non-toxic to NIH 3T3 and L132 healthy cells, yet they are more cytotoxic in vitro than cisplatin against HeLa and A549 cancer cells. These results showed that the complexes were particularly detrimental to cancer cells probably due to their higher hydrophobicity. Both complexes were shown to accumulate preferentially in the A549 cells. Live cell confocal microscopy demonstrated ROS generation, crumpling and fragmentation of cell nuclei, elevated ROS levels comparable to H₂O₂, flow cytometry demonstrating early apoptosis (LR quadrant) in A549 cells, and confirmation of apoptotic cell death by AO/EB and DAPI staining and annexin V-FITC/PI assays. Additionally, complexes in the cell cycle analysis prevent the proliferation of A549 cancer cells by causing cell cycle arrest at the S-phase. As a result, early apoptosis, ROS production, and S-phase arrest show that the complexes prevent cancer cell proliferation and trigger apoptosis. The biological properties of the newly developed mixed-ligand copper(II) complexes outlined here provided a substantial anticancer activity and are suitable for in vivo research and further investigation, which can facilitate the discovery of new anticancer agents with enhanced efficacy.

Author contributions

Mariappan Murali: conceptualization, funding acquisition, investigation, methodology, project administration, resources, supervision, writing – original draft, writing – review and editing; Anbarasu Kanchana Mala: formal analysis, investigation, methodology; Somasundaram Sangeetha: data curation, formal analysis, investigation, methodology; Balasubramaniam Selvakumaran: data curation, formal analysis, investigation, methodology; Pitchan Arul Prakash: formal analysis, investigation, methodology; Selvaraj Shanmugavadivel: data curation, investigation; Tamilarasan Ajay Kamal: methodology, software; Mohamed Sultan Mohamed Jaabir: conceptualization, funding acquisition, investigation, project administration, resources, supervision.

Conflicts of interest

There are no conflicts to declare

Data availability

The data supporting this article have been included as part of the supplementary information (SI). Supplementary information is available. See DOI: https://doi.org/10.1039/d5dt02476e.

CCDC 2143125 contains the supplementary crystallographic data for this paper.¹²⁶

Acknowledgements

We thank STIC, Cochin University of Science and Technology, for X-ray crystal structure data. We thank the SAIF, Indian Institute of Technology Madras, for structure solution and refinement and for recording EPR spectra. Author B. S. thanks the Council of Scientific and Industrial Research (CSIR), New Delhi, India, for giving the CSIR SRF (Direct) Fellowship [file no. 08/1365(25505)/2025-EMR-I]. Author TA acknowledges support from SRM TRP Engineering College, India (SRM/TRP/RI/005).

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Footnote

† This paper is in honor of Professor S. Natarajan, Solid State and Structural Chemistry Unit, Indian Institute of Science, Bangalore, India, for his lifetime of dedicated service and profound contributions to the field of Inorganic Solid State Chemistry.

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