Newly developed palladium complexes featuring ONS donor ligands: synthetic method, characterization, CT-DNA interaction analysis, BSA protein binding study and in vitro cytotoxicity

Abstract

Two new palladium(II) complexes (C1 and C2) were synthesized using imine- and azo-functional ligands, namely, (E)-4-hydroxy-3-(1-((2-(methylthio)phenyl)imino)ethyl)-2H-chromen-2-one (HL¹) and (E)-6-((2-(ethylthio)phenyl)diazenyl)-7-hydroxy-4-methyl-2H-chromen-2-one (HL²). The ligands and their corresponding complexes were comprehensively characterized by NMR, IR, mass spectrometry, and UV-Vis spectroscopy. The molecular structures of the complexes were confirmed using single-crystal X-ray diffraction study. Both complexes exhibited strong interactions with CT-DNA and BSA, demonstrating high binding affinity as confirmed by UV-Vis and fluorescence spectroscopy. Furthermore, the in vitro cytotoxic activity of the complexes was tested on MCF-7 (human breast epithelial adenocarcinoma) and MCF-10A (human breast epithelial) cell lines using clonogenic and nuclear fragmentation assays. Both complexes showed potent anti-proliferative activities, with C2 displaying slightly higher potency than C1.

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Introduction

It has been more than four decades since the approval of the highly potent anticancer drug cis-platin. In the last four decades, cisplatin has been used extensively as a chemotherapeutic drug for numerous cancers, including sarcomas, carcinomas, lymphomas and others.^1,2 Despite being a highly effective and widely used chemotherapeutic agent, the search for other potent metal-based drugs remains a topic of great interest. This continued search is driven largely by the wide range of adverse side effects associated with cisplatin therapy, such as nephrotoxicity, neurotoxicity, ototoxicity, tumor resistance, nausea, thrombosis, fatigue, and alopecia.³ In recent years, platinum-group metal complexes incorporating multidentate ligands have emerged as promising alternative chemotherapeutic candidates, offering enhanced cancer cell selectivity and reduced tumor resistance.^4–7

If we consider palladium complexes specifically, they exhibit diverse mechanisms of action, making them promising candidates for the development of novel cancer therapies. Notably, these complexes demonstrate strong tumor-targeting capabilities, selectively accumulating in cancerous tissues while sparing healthy cells. This targeted behavior not only enhances the therapeutic efficacy but also reduces the systemic toxicity, a major limitation of many conventional anticancer agents. Consequently, the design and synthesis of new palladium complexes have become areas of significant research interest.^8–12

Moreover, selecting suitable multidentate (ONS donor) ligands is crucial when designing palladium complexes, as the inherently high ligand-exchange rate of Pd(II) can lead to unintended interactions with non-target biomolecules in the human body. Such off-target binding may prevent the complexes from reaching and interacting with DNA—the primary therapeutic target—thereby diminishing their anticancer potency.¹³ In recent years, numerous palladium complexes have been synthesized that demonstrate strong anticancer activity, in some cases exhibiting greater therapeutic potential than cisplatin.^14–18 Among these, palladium complexes incorporating azo and Schiff-base ligands have emerged as particularly promising candidates. These ligands create well-defined coordination environments that effectively encapsulate the palladium center, resulting in highly stable complexes capable of circulating through the bloodstream and reaching target cells.¹⁹ For effective anticancer action, such complexes must ultimately interact with and damage cellular DNA, thereby disrupting uncontrolled cell proliferation. To reach the vicinity of DNA, however, they must first be transported through the bloodstream. Human serum albumin (HSA), the most abundant transport protein in blood plasma, plays a key role in this process by binding and carrying palladium complexes to their target sites.²⁰

Building on the extensive prior knowledge available on the coordination chemistry of palladium complexes, we synthesized two ligands, (E)-4-hydroxy-3-(1-((2-(methylthio)phenyl)imino)ethyl)-2H-chromen-2-one (HL¹) and (E)-6-((2-(ethylthio)phenyl)diazenyl)-7-hydroxy-4-methyl-2H-chromen-2-one (HL²), along with their corresponding Pd(II) complexes, designated as C1 and C2, respectively (Scheme 1).

Scheme 1 Synthetic procedure of the palladium(II) complexes, C1 and C2.

The synthesized ligands and their Pd(II) complexes were characterized using a variety of spectroscopic and analytical techniques, including IR, NMR, mass spectrometry, UV-Vis absorption spectroscopy, and single-crystal X-ray diffraction. Density functional theory (DFT) and time-dependent DFT (TD-DFT) calculations were carried out to gain deeper insight into the electronic structures of the complexes. Furthermore, the binding interactions of the ligands and complexes with calf thymus DNA (CT-DNA) were examined using multiple spectroscopic methods to evaluate their proficiency as DNA-cleaving or DNA-interacting anticancer agents.^21–23 To assess their binding affinity with the transport protein human serum albumin (HSA), absorption and fluorescence spectroscopy were employed. Bovine serum albumin (BSA) was used as a substitute for HSA due to its lower cost, widespread availability, and strong structural homology to HSA. BSA consists of a single polypeptide chain containing 583 amino acid residues, with a molecular mass of approximately 66 000 Da, and possesses two tryptophan (Trp) residues located at positions 134 and 213.²⁴ The in vitro cytotoxicity of the palladium complexes was evaluated using the MTT assay, where MTT refers to 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide against cancerous human breast epithelial cells (MCF-7) and non-tumorigenic human breast epithelial cells (MCF-10A). The anti-proliferative efficacy of the complexes against the MCF-7 cell line was further assessed using a clonogenic assay. In addition, the apoptotic potential of complexes C1 and C2 toward the MCF-7 cells was investigated through a nuclear fragmentation Assay.

Experimental

Materials and methods

The reagents and solvents that were used in these syntheses were purchased from Aldrich. The organic chemicals and inorganic salts were made available from commercial sources and used without further purification. ¹H and ¹³C NMR spectra were recorded on Bruker 300 MHz and 400 MHz instruments (mentioned in each spectrum) in DMSO-d₆ solvent. HRMS mass spectra were recorded on a Waters quadruple time-of-flight mass spectrometer (Xevo G2 Q-TOF). Electronic spectra were taken on a Shimadzu UV-1900i spectrophotometer. IR spectra were recorded on a PerkinElmer Spectrum Two FT-IR Spectrometer.

Earle's MEM (#Cat.: 41500034) and DMEM/Ham's F-12 (#Cat.: 11320033) were obtained from Gibco. MTT reagent (#Cat.: 475989) and Crystal Violet (#Cat.: 115940) were obtained from Merck. Hoechst 33528 Stain (#Cat.: 14530) and methanol (#Cat.: 34860) were obtained from Sigma Aldrich. All commercially obtained reagents were utilized as received without further purification.

Syntheses

Synthesis of (E)-4-hydroxy-3-(1-((2-(methylthio)phenyl) imino)ethyl)-2H-chromen-2-one (HL¹). HL¹ was synthesised by the reaction of 0.1 g (0.48 mmol) of 3-acetyl-4-hydroxy-2H-chromen-2-one and 0.068 g (0.48 mmol) 2-(methylthio)aniline in methanol (10 mL) under refluxing condition for about 1 h. A light-yellow precipitate was formed, which was filtered and washed with methanol. The yield was 0.134 g (84%).

Anal. calc. for C₁₈H₁₅NO₃S: C, 66.44; H, 4.65; N, 4.30. Found: C, 66.65; H, 4.74; N, 4.35. ¹H NMR (300 MHz, DMSO-d₆) in ppm: δ 2.07 (s, 3H), 2.30 (s, 3H), 7.31–7.37 (m, 3H), 7.43–7.49 (m, 3H), 7.70 (t, J = 6 Hz, 1H), 8.02 (d, J = 5.8 Hz, 1H), 15.46 (s, 1H). ¹³C NMR (75 MHz, DMSO-d₆) in ppm: δ 14.6, 20.6, 97.7, 116.9, 120.1, 124.4, 126.0, 126.3, 126.6, 127.5, 129.8, 133.5, 135.2, 135.9, 153.7, 177.3. IR (cm⁻¹) in KBr: 3380 ν(O–H); 3012 ν(C–H); 1693 ν(C=O); 1600 ν(C=N); 746 ν(C–S). HRMS: calculated for [M + H]⁺ (m/z): 326.0851; found: 326.0544. UV-Vis (in CH₃CN), λ_max (ε, M⁻¹ cm⁻¹): 326 (23 904), 244 (33 240).

Synthesis of (E)-6-((2-(ethylthio)phenyl)diazenyl)-7-hydroxy-4-methyl-2H-chromen-2-one (HL²). At first, an ice-cold solution of 2.7 g (13.45 mmol) 2-(ethylthio)aniline and 1 : 1 HCl (10 mL) was made using an ice bath. To this solution, an ice cold solution of NaNO₂ (2.0 g in 10 mL water) was added under stirring condition. Then, the whole solution was added to an ice-cold solution of Na₂CO₃ (6 g in 25 mL water) and 2.37 g (13.45 mmol) 7-hydroxy-4-methyl-2H-chromen-2-one with continuous stirring. An orange-red precipitate was obtained, which was filtered and washed with cold water, then dried using a desiccator. Further purification was done using a silica gel (mesh 60–120) column. A bright orange-red band of HL² was eluted by 40% (v/v) ethyl acetate-petroleum ether mixture. Pure HL² was yielded by evaporating the solvent under reduced pressure. The yield was 3.25 g (71%).

Anal. calc. for C₁₈H₁₆N₂O₃S: C, 63.51; H, 4.74; N, 8.23. Found: C, 63.27; H, 4.81; N, 8.95. ¹H NMR (300 MHz, CDCL₃): δ 1.43 (t, J = 7.3Hz, 3H), 2.52 (s, 3H), 2.91 (q, J = 7.3 Hz, 2H), 6.23 (s, 1H), 6.97 (s, 1H), 7.57–7.39 (m, Ar-H), 7.87 (d, J = 7.6 Hz, 1H), 8.22 (s, 1H), 13.31 (s, 1H). ¹³C NMR (75 MHz, DMSO-d₆) in ppm: δ 14.2, 18.5, 25.5, 102.6, 110.7, 112.5, 113.3, 116.8, 124.4, 127.1, 154.0, 155.3, 157.3, 159.9, 160.7, 161.6. IR (KBr, cm⁻¹): 3527 ν(O–H); 2805 ν(C–H); 1628 ν(C=O); 1351 ν(N=N); 817 ν(C–S). HRMS: calculated for [M + H]⁺ (m/z): 341.0960; found: 341.0916. UV-Vis (in CH₃CN), λ_max (ε, M⁻¹ cm⁻¹): 410 (7040), 317 (36435).

Synthesis of [Pd(L¹)Cl] (C1). At first, 0.045 g (0.25 mmol) of palladium chloride (PdCl₂) was added in acetonitrile and solvation was achieved by heating the solution. Then, to this solution, HL¹ was added (0.082 g, 0.25 mmol) and the resulting mixture was refluxed for about 5 h. Then, the mixture was cooled to room temperature and filtered. Orange coloured single crystals of the complex suitable for diffraction were obtained after a few days by the slow evaporation of the solvent. The yield obtained was 0.087 g (74%).

Anal. calc. for C₁₈H₁₄ClNO₃PdS: C, 46.37; H, 3.03; N, 3.00. Found: C, 46.68; H, 3.17; N, 3.07. ¹H NMR (300 MHz, DMSO-d₆): δ 2.08 (s, 3H), 2.40 (s, 3H), 7.31–7.49 (m, 5H), 7.71 (t, J = 6.8 Hz, 1H), 7.95 (d, J = 7.6 Hz, 1H), 8.02 (d, 1H). IR (KBr, cm⁻¹): 3060 ν(C–H); 1697 ν(C=O); 1605 ν(C=N); 758 ν(C–S). HRMS: calculated for C₁₈H₁₄NO₃PdS [M–Cl]⁺ (m/z): 430.7937; found: 430.7211. UV-Vis (in DMSO), λ_max (ε, M⁻¹ cm⁻¹): 450 (10 856), 363 (24 488).

Synthesis of [Pd(L²)Cl] (C2). 0.05 g (0.28 mmol) of palladium chloride (PdCl₂) was added in acetonitrile and solvation was achieved the same way. Then, to this solution, HL² (0.095 g, 0.28 mmol) was added. The resulting mixture was refluxed for about 5 h. Then, the mixture was cooled to room temperature and filtered. Dark orange coloured single crystals of the complex suitable for diffraction were obtained after a few days by the slow evaporation of the solvent. The yield obtained was 0.102 g (76%).

Anal. calc. for C₁₈H₁₅ClN₂O₃PdS: C, 42.92; H, 3.14; N, 5.82. Found: C, 42.57; H, 3.23; N, 5.65. ¹H NMR (300 MHz, CDCl₃): δ 1.50 (t, J = 7.9 Hz, 3H), 3.06 (q, J = 7.5 Hz, 2H), 2.55 (s, 3H), 6.27 (s, 1H), 7.08 (s, 1H), 7.74–7.58 (m, Ar–H), 7.87 (d, J = 10.8 Hz, 1H), 8.19 (s, 1H). IR (KBr, cm⁻¹): 2963 ν(C–H); 1635 ν(C=O); 1334 ν(N=N); 760 ν(C–S). HRMS: calculated for C₁₈H₁₅ClN₂O₃PdS [M–Cl]⁺ (m/z): 444.9838; found: 444.9446. UV-Vis (in DMSO), λ_max (ε, M⁻¹ cm⁻¹): 330 (29 964), 396 (sh.), 515 (16 099).

Crystallographic data gathering and refining

A Bruker AXS D8 Quest CMOS diffractometer was used to collect diffraction data with the help of graphite monochromatized Mo-K_α radiation (λ = 0.71073), and reflection data were recorded using the ω scan method. The SAINT program was used for cell refinement, indexing, and scaling of the data, and SADABS was used to make the multi-scan absorption corrections. All data were corrected for Lorentz and polarization effects. The refinement was performed by considering the anisotropic effect of all non-hydrogen atoms, and the hydrogen atoms were placed geometrically in accordance with the riding model. The structures were resolved by the direct method and refined using the SHELXL-2016 program²⁵ by full-matrix least-squares techniques. All crystallographic parameters are given in SI, Tables S1–S3.

Computational method

Geometry optimizations of the palladium complexes were performed using density functional theory (DFT) with the B3LYP hybrid exchange–correlation functional.²⁶ For the Pd atom, the LANL2DZ basis set with its associated effective core potential was employed,²⁷ while all remaining atoms were treated using the 6-31G(d) basis set.²⁸ Vibrational frequency calculations were carried out for each optimized structure to ensure that all geometries correspond to true minima on the potential energy surface. All calculations were performed using the Gaussian 09 software package,²⁹ and molecular structures and orbitals were visualized using GaussView 5. Probable electronic transitions were obtained using time-dependent DFT (TD-DFT)³⁰ in conjunction with the conductor-like polarizable continuum model (CPCM)³¹ in acetonitrile, simulating the solvent environment used in the experimental studies. The fractional contributions of different fragments to the molecular orbitals were analyzed using the GaussSum program.³²

Hirshfeld surface analysis of the complexes

In order to visualize the intermolecular interactions in the crystal, molecular Hirshfeld surfaces (HS) and the associated 2D-fingerprint plots of C1 and C2 were calculated using Crystal Explorer 17.5 software. Bond lengths to hydrogen atoms were automatically set to standard values, while the structural input CIF files of the crystals C1 and C2 were read into the software for calculations. For each point on the Hirshfeld isosurface, two distances, d_e (the distance from the point to the nearest nucleus external to the surface) and d_i (the distance to the nearest nucleus internal to the surface), are defined. The normalized contact distance (d_norm) based on d_e and d_i is given by eqn (1):where r^vdW_i and r^vdW_e are the van der Waals radii of the atoms. The value of d_norm is negative or positive depending on the intermolecular contacts being shorter or longer than the van der Waals separations. Graphical plots of the molecular Hirshfeld surfaces mapped with d_norm use a red-white-blue colour scheme, where bright red spots highlight shorter contacts, white areas represent contacts around the van der Waals separation, and blue regions are devoid of close contacts. To visualize the molecular moiety, transparent mapped surfaces are usually shown. For a given crystal structure and set of spherical atomic electron densities, the Hirshfeld surface is unique. Thus, it suggests the possibility of gaining additional insight into the intermolecular interactions of the molecular crystals. The 2D fingerprint plot resolved into different contacts contributed to the total Hirshfeld surface area of complexes C1 and C2.³³

DNA interaction studies

The binding studies of the complexes with calf thymus (CT) DNA were conducted in Tris–HCl/50 mM NaCl buffer (pH 7.4). A stock solution of CT DNA was prepared in this buffer, and its concentration was determined spectrophotometrically by measuring the absorbance at 260 nm and dividing the value by the molar extinction coefficient (ε₂₆₀ = 6600 M⁻¹ cm⁻¹). Stock solutions of the complexes and ligands (∼10⁻⁴ M) were prepared in DMSO, and were appropriately diluted with Tris buffer as required for the experiments. Absorption titration studies were performed by the gradual addition of CT DNA solution to a fixed concentration of the complex (or ligand) solution, while monitoring the changes in absorbance. A competitive binding study with ethidium bromide (EB) was carried out using fluorescence spectroscopy to evaluate whether the ligands and complexes are capable of displacing EB from the EB-CT DNA adduct. The EB-CT DNA adduct was prepared by mixing 15 µM EB with 30 µM CT DNA in Tris-HCl/NaCl buffer (pH 7.4). Changes in the emission spectra were recorded upon the incremental addition of the ligands or complexes to this adduct solution. The observed fluorescence quenching was attributed to the displacement of EB from its intercalated EB-CT DNA complex.³⁴

BSA protein binding studies

The binding ability of the compounds toward bovine serum albumin (BSA) was examined using both absorption and fluorescence spectroscopy. A BSA stock solution (∼10⁻⁵ M) was prepared in 500 mM phosphate-buffered saline (PBS) at pH 7.4 and stored at 4 °C in the dark. The concentration of BSA was determined spectrophotometrically by measuring its absorbance at 280 nm using a molar extinction coefficient of 66 400 M⁻¹ cm⁻¹. Stock solutions of the ligands and complexes (∼10⁻⁴ M) were prepared in DMSO and were appropriately diluted with PBS, as required for the experiments. Absorption titration was carried out by maintaining a constant concentration of BSA, while gradually adding the ligand or complex solutions. For fluorescence titration, the intrinsic tryptophan fluorescence of BSA was recorded from 290–500 nm (excitation at 280 nm; slit width: 10 nm). Synchronous fluorescence spectra were also collected by fixing the excitation wavelength at 240 nm and scanning the emission from 255–400 nm. Two wavelength intervals, Δλ = 15 nm and Δλ = 60 nm, were used to monitor the microenvironmental changes around the tyrosine (Tyr) and tryptophan (Trp) residues of BSA, respectively.^35,36

Biological study

Cell lines and culture condition

The MCF-7 (i.e., human breast epithelial adenocarcinoma cell line) and MCF-10A (i.e., human breast epithelial cell line) cell lines were obtained from NCCS-Pune, India. The MCF-7 and MCF-10A cells were cultured in Earle's MEM and DMEM/Ham's F-12 medium, respectively. Both media were supplemented with 10% FBS and 1% Anti-Anti. Cells were harvested in an incubator, where the humidified condition was maintained properly, and the temperature and the CO₂ concentration were kept at 37 °C and 5%, respectively.

In vitro cytotoxicity

Cytotoxic effects of both C1 and C2 complexes against MCF-7 and MCF-10A cells were tested by MTT assay with our established protocol.³⁷ In brief, 1 × 10⁴ cells per well were plated in 96 well plates, incubated for 24 h, and then treated with various doses of C1 and C2 complexes ranging from 1.56 µM to 100 µM for 24 h. On the very next day, MTT solution was added for 3 h, followed by the addition of DMSO–methanol (1 : 1). Finally, the absorbance values were obtained at 570 nm, and the cell viability and cytotoxicity values were calculated.

Clonogenic assay

The colony-forming inhibition abilities of the C1 and C2 complexes against MCF-7 cells were tested by clonogenic assay with our established protocol.³⁸ In brief, 500 cells per well were plated in 6 well plates, incubated for 24 h, and then treated with C1 (8.33 and 16.66 µM) and C2 (4.73 and 9.46 µM) complexes for 24 h. On the very next day, methanol and 0.05% crystal violet were utilized for cell fixing and staining, followed by washing with PBS. Finally, images of all wells were taken with a digital camera, and the relative colony numbers were calculated.

Nuclear fragmentation assay for apoptosis detection

The nuclear fragmentation abilities of the C1 and C2 complexes against MCF-7 cells were tested by nuclear fragmentation assay with our established protocol.³⁹ In brief, 1 × 10⁵ cells per well were plated on sterile coverslips, incubated for 24 h, and then treated with C1 (8.33 and 16.66 µM) and C2 (4.73 and 9.46 µM) complexes for 24 h. On the very next day, 4% formaldehyde and Hoechst 33 258 were utilized for cell fixing and staining, followed by washing with PBS. Finally, the coverslips were mounted in glycerol and the nuclear morphologies were observed under a fluorescent microscope (Olympus).

Statistical analysis

GraphPad Prism 8.0.2 Software was used to perform the statistical analysis. All experiments were repeated independently three times, and the observed data were recorded as (mean ± SD). One-way ANOVA was performed to evaluate the significant difference between the control and treated groups, where the P < 0.05 was considered statistically significant. CompuSyn software was used for the inhibitory concentration (IC₅₀) value calculations, which was specifically performed based on Chou-Talalay's method.⁴⁰

Results and discussion

Synthesis and spectral characterization

The coumarin-based Schiff base ligand HL¹ was synthesized by reacting 3-acetyl-4-hydroxy-2H-chromen-2-one with 2-(methylthio)aniline in methanol. The corresponding Pd(II) complex C1 was then obtained by reacting HL¹ with PdCl₂ in a 1 : 1 molar ratio in acetonitrile under reflux conditions (Scheme 1). Ligand HL² was prepared via the conventional diazo-coupling reaction between 2-(ethylthio)aniline and 7-hydroxy-4-methylcoumarin. The Pd(II) complex C2 was synthesized by reacting HL² with PdCl₂ in a 1 : 1 molar ratio in acetonitrile under reflux. The structures of the ligands and complexes were confirmed using a combination of spectroscopic and analytical techniques, including NMR, IR, UV-Vis spectroscopy, and mass spectrometry. Structures of the complexes were confirmed by single-crystal X-ray diffraction method.

NMR spectroscopy

¹H NMR spectra of HL¹ and C1 were recorded in DMSO-d₆. In the spectrum of HL¹, the phenolic O–H proton appeared as a singlet at 15.46 ppm, while the three protons of the –SCH₃ group appeared as a singlet at 2.30 ppm. In the spectrum of C1, the O–H signal disappeared, and the –SCH₃ peak shifted downfield to 2.40 ppm, indicating coordination of the palladium center through the phenolic oxygen and thioether sulfur atoms. For HL² and C2, ¹H NMR spectra were recorded in CDCl₃. The phenolic O–H proton of HL², observed at 13.31 ppm, disappeared in the spectrum of C2, while the –SCH₂ protons experienced a downfield shift from 2.91 ppm in HL² to 3.06 ppm in C2. These observations similarly suggest that complexation occurs via phenolic oxygen and thioether sulfur atoms (SI, Fig. S1–S4 and S11–S12).

IR spectroscopy

In the IR spectrum of HL¹, a broad band at 3380 cm⁻¹ was assigned to O–H stretching, and a band at 1600 cm⁻¹ corresponded to C=N (imine) stretching. Upon complexation to form C1, the O–H stretching band disappeared, and the C=N stretching band shifted to 1557 cm⁻¹, indicating coordination of the ligand to the Pd(II) center. Similarly, the IR spectrum of HL² exhibited an O–H stretching band at 3527 cm⁻¹ and an N=N (azo) stretching band at 1351 cm⁻¹. In the corresponding complex C2, the O–H band disappeared, and the N=N band experienced a red shift to 1334 cm⁻¹, consistent with coordination to palladium (SI, Fig. S5–S6 and S13–S14).

Mass spectrometry

High-resolution mass spectra (HRMS) of the ligands and complexes were recorded in methanolic solution (SI, Fig. S7–S8 and S15–S16). The molecular ion peak for HL¹ appeared at m/z 326.0544, corresponding to [M + H]⁺, while HL² showed a peak at m/z 341.0960, also corresponding to [M + H]⁺. For C1, the molecular ion peak was observed at m/z 430.7211, assignable to [M–Cl]⁺. For C2, a peak at m/z 444.9446 was observed, corresponding to [M–Cl]⁺.

UV-Vis study

The absorption spectra of the ligands and complexes were recorded in acetonitrile and DMSO, respectively (SI, Fig. S9, S10 and S17, S18). HL¹ exhibited a sharp absorption peak at 326 nm (ε = 23 904 M⁻¹ cm⁻¹), along with a higher energy peak at 244 nm (ε = 33 240 M⁻¹ cm⁻¹). HL² showed a sharp peak at 317 nm (ε = 36 435 M⁻¹ cm⁻¹) and an additional band at 410 nm (ε = 7040 M⁻¹ cm⁻¹). Upon complexation, C1 displayed a broad band at 450 nm (ε = 10 856 M⁻¹ cm⁻¹), along with a higher energy peak at 363 nm (ε = 24 488 M⁻¹ cm⁻¹). Complex C2 showed a high-energy peak at 330 nm (ε = 29 964 M⁻¹ cm⁻¹), a lower energy peak at 515 nm (ε = 16 099 M⁻¹ cm⁻¹), and a shoulder at 396 nm. These spectral shifts and new low-energy bands indicate ligand-to-metal charge transfer (LMCT) transitions and confirm complex formation.

Crystallographic structure analysis

The structures of C1 and C2 were determined by single-crystal X-ray diffraction. ORTEP diagrams with labeled atoms are shown in Fig. 1. Detailed crystallographic data, including selected bond lengths and angles, are provided in the SI (Tables S2 and S6). C1 crystallizes in the monoclinic system with space group C2/c. In C1, ligand HL¹ coordinates to the palladium center via one nitrogen atom (N1), one oxygen atom (O1), and one sulfur atom (S1), forming a five-membered chelate ring [N1–C12–C17–S1–Pd1] and a six-membered chelate ring [N1–C10–C9–C1–O1–Pd1]. The trans angles around the metal center are nearly equal (<O1–Pd1–S1 = 175.03(8)°; <N1–Pd1–Cl1 = 176.64(8)°), while the remaining angles are close to 90° (<O1–Pd1–N1 = 94.02(10)°; <N1–Pd1–S1 = 88.38(8)°; <O1–Pd1–Cl1 = 89.28(7)°; <S1–Pd1–Cl1 = 88.29(3)°). The bond lengths are Pd1–N1 = 2.019(3) Å, Pd1–O1 = 1.982(2) Å, Pd1–S1 = 2.2174(9) Å, and Pd1–Cl1 = 2.3036(8) Å. These parameters indicate that the palladium center adopts an almost planar geometry (τ₄ = 0.06). On the other hand, C2 was crystallized in a triclinic crystal system having space group P1̄. Ligand HL² coordinates through O1, N1, and S1, forming a six-membered chelate ring [N1–N2–C7–C16–O1–Pd1] and a five-membered chelate ring [N1–C6–C1–S1–Pd1]. The trans angles around the metal center are nearly equal (<O1–Pd1–S1 = 179.68(9)°; <N1–Pd1–Cl1 = 177.35(9)°), and the remaining angles are close to 90° (<N1–Pd1–O1 = 92.59(12)°; <N1–Pd1–S1 = 87.57(10)°; <O1–Pd1–Cl1 = 89.68(9)°; <S1–Pd1–Cl1 = 90.16(4)°). Bond lengths are Pd1–N1 = 1.983(3) Å, Pd1–O1 = 1.992(3) Å, Pd1–S1 = 2.2404(11) Å, and Pd1–Cl1 = 2.3118(12) Å. These data confirm that the palladium center in C2 also adopts an essentially planar, square-planar geometry (τ₄ = 0.03), consistent with previously reported Pd(II) complexes.⁴¹

Fig. 1 ORTEP plot of C1 (A) and C2 (B) with 35% ellipsoid probability.

Computational analysis

The geometries of C1 and C2 were optimized using the DFT/B3LYP method. The calculated bonding parameters are in good agreement with the experimental X-ray crystallographic data. Energies and compositions of selected molecular orbitals (MOs) are provided in the SI (Tables S4 and S6). Contour plots of representative MOs of C1 and C2 are also shown in the SI (Fig. S23 and S24). For C1, the HOMO is composed of 64% ligand (HL¹), 17% Pd(dπ), and 19% Cl(pπ), while the LUMO consists of 70% HL¹, 22% Pd(dπ), and 8% Cl(pπ), indicating significant ligand–metal interactions. In C2, the HOMO contains 71% ligand (HL²), 18% Pd(dπ), and 11% Cl(pπ), whereas the LUMO is predominantly ligand-centered (97% HL²) with minor Pd(dπ) contribution (3%), highlighting the major role of the ligand in electronic transitions.

Time-dependent DFT (TD-DFT) calculations were performed in DMSO to elucidate the electronic transitions (SI, Tables S5 and S7). For C1, the HOMO → LUMO transition at 518 nm (f = 0.0386) and HOMO−1/HOMO−2 → LUMO transitions at 468 nm (f = 0.0129) correspond to the experimental absorption band at 450 nm (ε = 10 856 M⁻¹ cm⁻¹), involving mixed Pd(dπ)/Cl(pπ)/ligand (π) → ligand (π*) transitions. Transitions at 393 nm (HOMO−1 → LUMO+1, f = 0.1414) and 365 nm (HOMO−3 → LUMO+1, f = 0.1472) correspond well with the experimental band at 363 nm (ε = 24 448 M⁻¹ cm⁻¹), characterized by Cl(pπ)/ligand (π) → ligand (π*) transitions. For C2, experimental absorption bands at 330 nm (ε = 29 964 M⁻¹ cm⁻¹) and 515 nm (ε = 16 099 M⁻¹ cm⁻¹) are in good agreement with TD-DFT transitions at 326 and 312 nm (HOMO → LUMO+2/HOMO−6 → LUMO) and 502 and 478 nm (HOMO/HOMO−1 → LUMO), respectively. The low-energy transitions involve mixed Pd(dπ)/Cl(pπ)/ligand (π) → ligand (π*) character, confirming the nature of the experimentally observed bands.

Hirshfeld surface calculations of complexes

All the available close contacts (intermolecular interactions) present in C1 and C2 were analysed using Hirshfeld surface analysis and fingerprint plots. This provides detailed insight into the crystal packing behaviour and a 3D pictorial description of the supramolecular assembly via hydrogen bonding.⁴² The Hirshfeld surfaces of C1 and C2 were mapped over d_norm (with a range of −0.1 to 1.1 Å), shape index (with a range of −1.0 to 1.0 Å) and curvedness (with a range of −4.0 to 4.0 Å), and are shown in SI, Fig. S19 and Fig. S21. Circular red spots visible on the d_norm surfaces signify the hydrogen bonding contacts and other weak contacts (C–H⋯π/π⋯π) present in the molecule. These red areas are mainly responsible for hydrogen bonding with other molecules. The adjacent pit-bump red and blue areas present in the shape-index surface indicate the presence of π⋯π interactions in the molecule. The 2D fingerprint plot shows various intermolecular interactions, along with their reciprocal appearing as specific spikes (SI, Fig. S20 and S22). The breakdown of the 2D fingerprint plots allows us to unfold all the available individual interactions (H⋯H, C⋯C, H⋯C/C⋯H, H⋯N/N⋯H, H⋯S/S⋯H, H⋯O/O⋯H) in the complexes and their respective contributions to the total surface.⁴³ H⋯H interactions contribute significantly (41.5% for C1 and 32.5% for C2) to the overall surfaces of both structures, which are shown in Fig. 3 and 4.

The Hirshfeld surface study suggests the significant role of H-atom contacts and H⋯C/C⋯H, C⋯C, and H⋯(O/N/S)/(O/N/S)⋯H interactions in giving shape to the crystal packing. Also, the complexes can interact with other molecules via the van der Waals and hydrogen bonding interactions present in the crystals. These kinds of interactions are usually observed between many protein-small molecules. This fact motivates us to perform further studies (based on interaction) to explore the effects of the complexes on BSA and CT DNA (Fig. 2).⁴⁴

Fig. 2 2D-Fingerprint plot (full) and relative inputs of numerous intermolecular contacts to the total HS area for C1.

Fig. 3 2D-Fingerprint plot (full) and relative inputs of numerous intermolecular contacts to the total HS area for C2.

Fig. 4 Change in the absorption spectra of complexes C1 (A) and C2 (B) with the gradual addition of CT DNA. Inset: Plot of [DNA]/(ε_a − ε_f) vs. [DNA].

CT-DNA interaction analysis

Absorption spectral study. UV-Vis absorption titration is a very effective and widely used technique to study the binding mode and efficiency of the binding of metal complexes with CT DNA. When a metal complex binds with DNA, there is a characteristic shift of the wavelength of the electronic spectra of the complex that occurs, either a bathochromic or hypsochromic shift, along with a decrease in the absorption maxima (hypochromicity) or increase in the absorption maxima (hyperchromicity). Hypochromicity is mainly observed when stabilization of the DNA secondary structure occurs after interaction with a metal complex, while hyperchromicity indicates destabilization.⁴⁵

The gradual addition of CT-DNA to HL¹ resulted in a pronounced hyperchromicity of approximately 73% at 327 nm. For HL², a moderate hyperchromicity of about 13% was observed at both the ∼320 nm and 415 nm bands (SI, Fig. S25). In contrast, complex C1 exhibited significant hypochromicity of around 87% at 450 nm, accompanied by an isosbestic point at approximately 350 nm. For complex C2, hypochromicity of about 17% was observed for both the 515 nm and 330 nm bands (Fig. 4). Such spectral changes typically indicate effective association between the chromophoric units of the complexes and the DNA base pairs.

The binding constant (K_b) was determined by plotting [DNA]/(ε_a − ε_f) vs. [DNA] from the spectral absorption titration data using the Wolfe–Shimmer equation [eqn (2)].⁴⁶

Here, [DNA] represents the concentration of DNA, and ε_a, ε_f and ε_b represent the extinction coefficient, the extinction coefficient of the compounds in the absence of DNA and the extinction coefficient of the compounds when fully bound to CT DNA, respectively. The binding constants (K_b) were determined to be 7.16(±0.3) × 10³ M⁻¹ and 4.18(±0.08) × 10³ M⁻¹ for HL¹ and HL², respectively, and 2.62(±0.06) × 10⁵ M⁻¹ and 5.43(±0.2) × 10⁵ M⁻¹ for C1 and C2, respectively. These values are in good agreement with those reported for similar types of complexes.^19,47 The relatively high K_b values indicate the strong nature of the binding interactions of both the ligands and the complexes with CT DNA.

Fluorescence quenching study. In order to provide further evidence of the higher binding affinity of the compounds towards CT DNA, fluorescence quenching experiments were performed. Ethidium bromide (EB) is a widely known fluorescence probe that binds with CT DNA via intercalation to make the EB-CT DNA adduct, which shows high fluorescence intensity at around 610 nm when excited at 540 nm. If the compounds have more binding affinity towards CT DNA than EB, it can displace EB from the EB-CT DNA adduct, thereby significantly decreasing the fluorescence intensity.⁴⁸ In our study, both the ligands and the complexes were sufficiently competitive to displace EB from the EB–CT-DNA adduct. HL¹ and HL² caused decreases in fluorescence intensity of approximately 24% and 20%, respectively (SI, Fig. S26), while complexes C1 and C2 induced reductions of about 28% and 35%, respectively (Fig. 5).

Fig. 5 Emission spectra of EB-CT DNA with increasing concentrations of complexes C1 (A) and C2 (B). Inset: Plots of the emission intensity F₀/F vs. [Complex].

Fluorescence quenching was further analyzed using the Stern–Volmer equation [eqn (3)],⁴⁹ which also enabled the determination of the binding constants for both compounds.

Here, F₀ and F are the fluorescence intensities of EB-CT DNA in the absence and presence of the compounds, respectively, [Q] is the concentration of the added compounds, and K_SV is the Stern–Volmer quenching constant. The K_SV values of HL¹, HL², C1 and C2 were obtained from the slopes of the plots, and were calculated to be 6.26(±0.15) × 10³ M⁻¹, 5.44(±0.2) × 10³ M⁻¹, 2.45(±0.05) × 10⁵ M⁻¹ and 1.51(±0.06) × 10⁵ M⁻¹ for HL¹, HL², C1 and C2, respectively.

Both the absorption and fluorescence studies revealed that the ligands and their palladium complexes interact strongly with CT-DNA, with the corresponding binding constants showing good agreement. A detailed comparison of the K_SV values with those reported for similar palladium complexes is provided in SI, Table S8.^50–55 These results clearly indicate that coordination of the ligands to the palladium center enhances their DNA-binding capability.

BSA protein binding experiment

UV-Vis study. Albumins are the most common proteins found in the blood stream that act as a drug carrier. So, the binding of the complexes with bovine serum albumin (BSA) protein was of great interest. The primary investigation was done by recording absorption spectra in the range of 230–450 nm, while keeping the concentration of the BSA solution constant and gradually adding the compounds. BSA shows a characteristic peak at 280 nm (due to the presence of aromatic amino acid segments of the BSA backbone). If any conformational changes happen around these segments, the intensity of the peak at 280 nm shows some degree of hyperchromicity.⁵⁶ The ligands and complexes showed a steady increase of intensity at the band around 280 nm (Fig. 6) (SI, Fig. S27). This suggests the presence of a ground state interaction between the ligand or complexes and the protein. The apparent association constants (K) were calculated from the plot of 1/(A_obs − A₀) vs. 1/[compound] using the following eqn (4):where A_obs is the observed absorbance (at 280 nm) of the solution, and A₀ and A_c are the absorbance of BSA alone and of BSA with the compounds, respectively. The apparent association constants for HL¹, HL², C1 and C2 were calculated to be 2.59(±0.13) × 10⁴ M⁻¹, 6.83(±0.34) × 10³ M⁻¹, 4.08(±0.3) × 10⁵ M⁻¹ and 3.51(±0.25) × 10⁵ M⁻¹, respectively.

Fig. 6 Change in the absorption spectra of BSA with the increasing addition of complexes C1 (A) and C2 (B). Inset: Plot of 1/(A_obs − A₀) vs. 1/[complex].

Fluorometric study. The inherent fluorescence property of BSA, which is at 338 nm upon excitation at 280 nm, comes from the aromatic amino acid (AAA) segments present in BSA. Trp, Tyr and Phe AAA residues⁵⁷ have the relative fluorescence intensities of 100 : 09 : 0.5. Phe has a very low quantum yield, while the fluorescence of Tyr is nearly quenched when ionized or attached to a carboxyl or amino acid group or Trp. Therefore, the fluorescence of BSA essentially arises from the Trp unit, which also depends strongly on the surrounding environments. Any alteration of these surrounding environments causes quenching of the fluorescence intensity of BSA. Changes can happen if there is an interaction between BSA and the compounds.⁵⁸

Both ligands and complexes were non-fluorescent in DMSO solution. When titrated against the BSA solution, HL¹ (65%), HL² (62%), C1 (67%) and C2 (73%) showed significant quenching of fluorescence with the gradual addition of the ligands and complexes, as shown in Fig. 7 (SI, Fig. S28). The hypochromicity nature gives a clear idea that the BSA-complex formed in the ground state gave rise to a conformational change in the surrounding environment of Trp. In order to analyze the whole quenching process, the Stern–Volmer eqn (5) was used. The quenching rate constant (K_q) and Stern–Volmer quenching constant (K_sv) were calculated using the plot of F₀/F vs. [Q]:

where, F₀ and F are the fluorescence intensities of BSA in the absence and presence of the compounds, respectively, [Q] is the concentration of the added compounds, K_SV is the Stern–Volmer quenching constant, and τ₀ (5.42 × 10⁻⁹ s)⁵⁹ is the average fluorescence lifetime of BSA without compounds. The K_SV values were calculated to be 1.23(±0.25) × 10⁴ M⁻¹, 4.89(±0.15) × 10³ M⁻¹, 2.68(±0.14) × 10⁵ M⁻¹ and 4.52(±0.2) × 10⁵ M⁻¹ for HL¹, HL², C1 and C2, respectively. The calculated K_SV values for the complexes are comparable to the reported palladium complexes. The K_q values were also calculated and found to be in the range of 10¹¹–10¹² M⁻¹ S⁻¹, suggesting that the nature of quenching is dynamic rather than static (as the K_q values are higher than that of the average dynamic quenching constant (2.0 × 10¹⁰ M⁻¹ S⁻¹).⁶⁰

Fig. 7 Emission spectra of BSA in the presence of gradual increase in concentrations of the C1 (A) and C2 (B). Inset: Plots of emission intensity F₀/F vs. [complex].

Estimation of the binding constant and counting binding sites⁻

With the help of the Scatchard equation⁶¹ (6), from log [(F₀ − F)/F] versus log[compound] (SI, Fig. S29 and S30), the binding equilibrium constants (K_b) and number of binding sites (n) accessible for interaction with the complexes were estimated.The calculated binding constants (K) and binding site numbers (n) were 9.73(±0.27) × 10³ M⁻¹, 0.983 (±0.01) for HL¹, 5.17(±0.13) × 10⁴ M⁻¹, 0.969 (±0.02) for HL², 3.16(±0.2) × 10⁵ M⁻¹, 0.98(±0.03) for C1 and 4.65(±0.18) × 10⁵ M⁻¹, 0.956 (±0.02) for C2, indicating a high affinity of both the ligands and complexes toward serum albumin. Notably, the complexes exhibited stronger binding than the free ligands. All data related to the BSA interaction study are summarized in Table 1 for clarity.

Table 1 Apparent association constant (K), binding equilibrium constant (K_b), Stern–Volmer quenching constant (K_sv) and number of binding sites (n) of the BSA protein with HL¹, HL², C1 and C2

Complex	UV-vis study	Fluorometric study
	K (M^−1)	K b (M^−1)	K sv (M^−1)	n
BSA + HL^1	2.59(±0.13) × 104	9.13(±0.27) × 103	1.23(±0.25) × 104	0.983(±0.01)
BSA + HL^1	6.83(±0.34) × 103	5.17(±0.13) × 103	4.89(±0.15) × 103	0.969(±0.02)
BSA + C1	4.08(±0.3) × 105	3.16(±0.2) × 105	2.68(±0.14) × 105	0.98 (±0.03)
BSA + C2	3.51(±0.25) × 105	4.65(±0.18) × 105	4.52(±0.2) × 105	0.956 (±0.02)

---

Synchronous fluorometric spectral study

Fluorophore moieties, namely, tryptophan and tyrosine, are of particular interest as these are the primary sources of fluorescence in BSA. Synchronous fluorescence spectroscopic analysis is a good technique to understand the change in the microenvironment near these fluorophores.⁶² Both of these segments show emission individually at different wavelength bands when specific differences between the excitation and emission wavelengths (Δλ = λ_em − λ_ex) were taken. The Δλ value of 15 nm is characteristic of the tyrosine moiety of BSA, and the Δλ value of 60 nm is characteristic of the tryptophan moiety of BSA.⁶³ Any shift of these emission spectra with the gradual addition of any foreign substances is actually related to changes in polarity near the fluorophores.⁶⁴ Now, the synchronous spectra were recorded with the incremental addition of both complexes in BSA solution with a specific Δλ (15 nm or 60 nm) value. C1 and C2 both showed small hypochromic effects of around 33% and 30% in the 298 nm region when Δλ was set to be 15 nm (Fig. 8). However, for Δλ = 60 nm, the band at 337 nm showed a significant decrease in intensity, nearly 65% for C1 and 75% for C2. Additionally, a red shift of 2 nm was observed for C2 in the 337 nm region (Fig. 9).

Fig. 8 The synchronous spectra of BSA (with wavelength difference, Δλ = 15 nm) in the presence of increasing concentration of complexes C1 (A) and C2 (B).

Fig. 9 The synchronous spectra of BSA (with wavelength difference, Δλ = 60 nm) in the presence of increasing concentration of complexes C1 (A) and C2 (B).

All these results conclude that C1 and C2 both interact with BSA very strongly, and the interaction is also suitable for further study regarding the anti-proliferative activity of the complexes.

Biological study

In vitro cytotoxicity. The in vitro cytotoxic activities of the C1 and C2 complexes against MCF-7 cells were evaluated using the MTT assay. Cell viabilities were calculated from absorbance values using the standard formula (SI, eqn (S1)), and the results are presented in Fig. 10. The IC₅₀ values were determined using the CompuSyn software and are summarized in SI, Table S9. These data clearly indicate that complex C2 exhibits higher cytotoxic potency (IC₅₀ = 9.46 µM) than complex C1 (IC₅₀ = 16.66 µM) toward MCF-7 human breast adenocarcinoma cells, consistent with values reported for similar palladium complexes^65,66 (SI, Table S9). In contrast, both complexes display negligible cytotoxicity toward normal MCF-10A human breast epithelial cells, with IC₅₀ values of 125.93 µM for C1 and 140.29 µM for C2.

Fig. 10 Relative cell viability (%) of complexes C1 (A) and C2 (C) against the MCF-7 cell line. Relative cell viability (%) against the MCF-10A cell line for complexes C1 (B) and C2 (D).

These findings confirm that both compounds exhibit significant in vitro cytotoxicity toward human breast epithelial adenocarcinoma (MCF-7) cells, while remaining essentially non-toxic to normal human breast epithelial (MCF-10A) cells.

Clonogenic assay. A clonogenic assay was performed to evaluate the colony-forming ability of MCF-7 cells in the presence of the C1 and C2 complexes. This assay also allowed assessment of their anti-proliferative properties, with the results presented in Fig. 11. The images clearly show a dose-dependent reduction in the colony number for both complexes, indicating that increasing concentrations of C1 and C2 progressively suppress colony formation. These observations confirm that both complexes possess significant in vitro anti-proliferative potential.

Fig. 11 Clonogenic assay of both complexes C1 and C2 (A) against the MCF-7 cell line. Relative colony formation (%) of complexes C2 (B) and C1 (C).

Nuclear fragmentation assay for apoptosis detection. A nuclear fragmentation assay was conducted to evaluate the ability of the C1 and C2 complexes to induce nuclear damage in MCF-7 cells. Changes in nuclear morphology serve as a distinct hallmark of apoptosis. As shown in Fig. 12, the number of nuclear fragments increased progressively with higher concentrations of both complexes. This dose-dependent rise in nuclear fragmentation confirms that C1 and C2 possess significant in vitro apoptotic potential.

Fig. 12 Nuclear fragmentation assay of both complexes C1 and C2 (A) against the MCF-7 cell line. Number of fragmented nuclei in the observed field for complexes C2 (B) and C1 (C).

Conclusions

Two new Pd(II) complexes (C1 and C2) featuring ONS donor pincer ligands (HL¹ and HL²) were synthesized and thoroughly characterized using spectroscopic methods and single-crystal X-ray diffraction. Structural analyses confirmed that both ligands act as tridentate ONS chelators in their respective complexes. DNA-binding studies employing UV-Vis absorption and fluorescence spectroscopy established that the ligands and complexes possess significant affinity for CT-DNA, while protein-binding experiments demonstrated strong interactions with BSA. Biological evaluations using MTT, clonogenic, and nuclear fragmentation assays revealed clear differences in cytotoxic profiles, with complex C2 displaying superior antiproliferative and apoptosis-inducing effects compared to complex C1 in MCF-7 breast cancer cells. These findings collectively suggest that ONS-based Pd(II) complexes, particularly C2, hold substantial promise as potential anticancer agents.

Conflicts of interest

There are no conflicts to declare.

Data availability

The data that support the findings of this study are available from the corresponding author on request.

The data supporting this article have been included as part of the supplementary information (SI). Supplementary information: NMR, IR, HRMS and UV-Vis spectra of all new compounds, XRD analysis, DFT calculation etc. See DOI: https://doi.org/10.1039/d5nj03343h.

CCDC 2359962 and 2359963 contain the supplementary crystallographic data for this paper.^67a,b

Acknowledgements

The authors thank SERB (No. EEQ/2018/000226), New Delhi, India for financial support. A. Halder (09/0096(13956)/2022-EMR-I) thanks CSIR, New Delhi, India for providing a fellowship. S. Guha and Dr G. Das thank DST-INSPIRE Faculty Project (Project Ref No. DST/INSPIRE/04/2020/001299) for their fellowships and the research grant.

Notes and references

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