Reversible Coordination Modes in Piperazine‐Based Ru(II)‐p‐Cymene Acylthiourea Complexes: Modulating DNA Groove Binding for Selective Antimelanoma Activity

Abstract

Acylthiourea ligands, featuring hard (N, O) and soft (S) donor atoms, enable diverse coordination modes in half‐sandwich Ru(II)‐p‐cymene complexes. Herein, we report the synthesis and full characterization of eight half‐sandwich Ru(II)‐p‐cymene complexes, comprising four monodentate (1m–4m) and four bidentate (1b–4b) species prepared under distinct synthetic conditions. The structures of the complexes 1m, 2m, 3m, 3b, and 4b were confirmed by single‐crystal X‐ray diffraction, while FT‐IR, UV–vis, ESI‐MS, and ¹H/¹³C NMR spectroscopy unambiguously distinguish the two binding modes. The reversible coordination mode was monitored by ¹H NMR and UV–vis spectroscopy, with base‐induced deprotonation favoring bidentate binding and HCl addition regenerating the monodentate form. Cytotoxic assays revealed that metal coordination markedly enhanced the antiproliferative activity of the ligands. Most bidentate complexes exhibited higher cytotoxicity than their monodentate analogues. In particular, 2b displayed a sixfold increase in potency (IC₅₀ = 6.05 μM) compared to 2m (36.4 μM) and showed much lower toxicity toward normal ARPE‐19 cells (IC₅₀ = 55.0 μM), indicating greater selectivity toward cancer cells. Molecular docking suggests DNA groove binding as the anticancer mechanism, highlighting the role of bidentate coordination, with 2b identified as the most promising Ru(II)‐p‐cymene complex.

Piperazine‐based monodentate and S,N‐bidentate Ru(II)‐p‐cymene complexes were synthesized through distinct coordination strategies and evaluated for antimelanoma activity. Among the investigated compounds, the S,N‐bidentate complex 2b exhibited significantly enhanced cytotoxicity (IC₅₀ = 6.1 ± 0.8 ) against melanoma cells, particularly the A375 cell line, compared with the monodentate analog 2m.

1. Introduction

Piperazine represents an important heterocyclic scaffold in antitumor drugs such as abemaciclib [1], entrectinib [2], olmutinib [3], and palbociclib [4]. Its incorporation frequently confers favorable pharmacokinetic properties [5], most notably improved aqueous solubility [6], while maintaining antitumor efficacy. Piperazine‐bearing compounds have also demonstrated preclinical anticancer efficacy through diverse mechanisms, including activation of the mitochondrial pathway (caspases‐3/‐8/‐9) [6] and inhibition of topoisomerase IIa [7]. Hybrid molecules combining piperazine with urea or thiourea moieties have shown enhanced antiproliferative effects, particularly against breast cancer cell lines [8], highlighting the synergistic potential of these structural motifs [9].

N‐Acylthiourea ligands have attracted considerable attention due to their ability to modulate the physicochemical and biological properties of metal complexes, including lipophilicity, stability, selectivity, and membrane permeability [10]. These compounds, which incorporate both amide and thioamide functionalities, represent an important class of molecules widely utilized in medicinal chemistry [11]. Their derivatives exhibit diverse biological activities and have found applications across numerous pharmaceutical domains [12, 14]. Recent studies highlight their therapeutic potential, particularly in anticancer research, where interactions of carbonyl (C=O) and thiocarbonyl (C=S) groups with DNA and topoisomerase I contribute to cytotoxic effects against solid tumors [15, 18]. Moreover, the C=S moiety facilitates peptide cross‐linking, expanding their biochemical relevance [19]. The sulfur‐ and nitrogen‐containing core of acylthioureas provides complementary hydrogen‐bond donor and acceptor sites, enabling versatile coordination and biomolecular interactions [20]. Notably, certain acylthiourea derivatives have been identified as CDK2/cyclin E inhibitors, effectively blocking cell‐cycle progression from G1 to S phase [21].

In half‐sandwich Ru(II)‐p‐cymene complexes, acylthiourea ligands display rich coordination chemistry [22]. They can bind as neutral monodentate S‐donor [23] or, following deprotonation of the thioamide N—H, as monoanionic bidentate chelates through either S,N or S,O chelation [24, 25]. The preferred coordination is governed by solvent polarity [23], pH [26], the substitution pattern of the thioamide (primary vs secondary) [27], and intramolecular hydrogen bonding [28], with primary acylthioureas typically favoring S,N chelation [29]. The anticancer activity of Ru–arene complexes has been comprehensively reviewed by the Lee group [30], and in another systematic study by the groups of Sadler and Dyson [30, 32], ruthenium(II)–arene scaffolds, especially those incorporating specific ligands, have emerged as promising candidates with cytotoxicity profiles often complementary to platinum drugs [33, 34]. Furthermore, structural and conformational tuning of these ruthenium(II) architectures has been widely documented to dictate their biomolecular affinity, effectively modulating their DNA‐binding pathways and, in certain strategic designs, successfully directing the complexes from traditional intercalators into selective groove binders [35, 36]. Within this class, the coordination mode of acylthiourea ligands has been repeatedly correlated with biological outcomes [37]. Bidentate S,N‐chelated complexes generally exhibit superior cytotoxic potency compared to their monodentate (S‐coordinated) counterparts, an effect attributed to the increased stability and altered cellular uptake imparted by the anionic chelate [29]. Gabriel H. Ribeiro et al. comprehensively reviewed the anticancer properties of Ru(II)–acylthiourea complexes in their literature survey [10]. The comparative cytotoxic activities of the mono‐ and bidentate S,N complexes are illustrated in Scheme 1.

SCHEME 1.

Selected examples of previously reported anticancer Ru(II)‐p‐cymene acylthiourea complexes (i–v) [24, 29, 40, 42] illustrating neutral monodentate S‐coordination versus monoanionic bidentate S,N‐chelation modes and their associated anticancer activities.

Despite these advances, piperazine‐substituted acylthioureas and their ruthenium(II)–arene complexes remain largely unexplored. Merging the established anticancer pharmacophores of piperazine and acylthiourea within a single ligand framework offers an attractive strategy to generate compounds with enhanced activity and selectivity [38]. In our earlier work [28], we demonstrated that deliberate control over the coordination mode of acylthiourea ligands, specifically the interconversion between monodentate (S‐coordinated) and bidentate (S,N‐chelated) binding, can serve as an effective strategy to modulate the physicochemical and structural properties of ruthenium complexes. Building on this foundation, our subsequent study [39] systematically evaluated how variations in auxiliary ligands influence anticancer activity while maintaining a single acylthiourea framework.

In the present study, we further extend this design strategy by introducing structural variation within the acylthiourea ligand itself, specifically through modification of the N‐aryl substituent, to probe its impact on cytotoxicity. Accordingly, we report the rational design and synthesis of a new series of piperazine‐containing N‐acylthiourea ligands and their ruthenium(II)–p‐cymene complexes, which adopt either monodentate S‐coordination or bidentate S,N‐chelation, thereby offering an expanded platform to investigate structure–activity relationships. Accordingly, the impact of coordination mode and ligand design on cytotoxicity is evaluated to establish structure–activity relationships and identify promising lead candidates.

2. Results and Discussion

2.1. Synthesis and Acid–Base‐Controlled Modulation of Monodentate and Bidentate Coordination in Ru(II) Complexes

The reaction of the dimeric ruthenium precursor [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ with acylthiourea ligands in a 1:2 metal‐to‐ligand ratio afforded two distinct classes of half‐sandwich Ru(II) complexes depending on the reaction conditions: neutral monodentate S‐coordinated complexes 1m–4m and monoanionic bidentate S,N‐chelated complexes 1b–4b, as illustrated in Scheme 2. Complexes 3m and 3b have been previously disclosed by our group [28]. As widely documented for acylthiourea ligands [24, 26, 29], coordination mode is highly sensitive to solvent polarity and medium pH. In nonpolar solvents under neutral conditions, the ligands bind exclusively through the thiocarbonyl sulfur atom in their neutral form [41, 43, 44], whereas polar protic solvents combined with mild base promote deprotonation of the thioamide —NH, favoring formation of a stable six‐membered S,N‐chelate.

SCHEME 2.

Synthesis of monodentate (1m–4m) and bidentate (1b–4b) Ru(II)–p‐cymene acylthiourea complexes and their interconversion through reversible acid–base modulation of coordination mode.

Guided by these principles, monodentate complexes 1m–4m were selectively prepared in toluene at room temperature, while bidentate complexes 1b–4b were obtained in methanol using sodium bicarbonate as base. A systematic study revealed that the coordination mode is fully reversible and can be controlled by simple acid–base manipulation. Treatment of monodentate complexes 1m–4m with organic (Et₃N) or inorganic (NaHCO₃) bases quantitatively yielded the corresponding bidentate derivatives 1b–4b. Conversely, exposure of 1b–4b to HCl in dichloromethane rapidly regenerated the protonated, monodentate forms 1m–4m. These findings establish that basic conditions favor bidentate coordination via amide deprotonation, while acidic environments promote reprotonation, restoring monodentate sulfur‐bound coordination. This reversible switching, monitored by ¹H NMR spectroscopy, highlights the dynamic coordination behavior of acylthiourea ligands and offers a versatile strategy for fine‐tuning the physicochemical and biological properties of Ru(II)–arene metallodrugs. All complexes were fully characterized by Fourier‐transform infrared (FT‐IR), UV–vis, electrospray ionization mass spectrometry (ESI‐MS), ¹H/¹³C NMR spectroscopy, and elemental analysis. Single‐crystal X‐ray diffraction analysis unambiguously confirmed the solid‐state structures of representative monodentate complexes 1m and 3m as well as bidentate complexes 2b, 3b, and 4b. The structural analyses of 3m and 3b were reported previously by our group [28].

2.2. Spectroscopic Differentiation of Monodentate and Bidentate Coordination Modes

The FT‐IR spectra provided initial evidence of the distinct coordination modes. The free acylthiourea ligands displayed characteristic vibrational bands at 3163–3152 cm⁻¹ (ν _N—H, amide), 3025–3019 cm⁻¹ (ν _N—H, thioamide), 1701–1687 cm⁻¹ (ν_C=O), 1241–1235 cm⁻¹ (ν _C=S), and 1331–1320 cm⁻¹ (ν _C—N) [38]. In the monodentate complexes 1m–4m, these bands underwent only minor shifts (Figures S1–S4), consistent with neutral S‐coordination through the thiocarbonyl sulfur atom [45, 46]. In contrast, the bidentate complexes 1b–4b showed disappearance of the amide —NH stretch, a marked redshift of ν _C=O to 1603–1597 cm⁻¹, and a slight shift of ν _C=S to 1228–1217 cm⁻¹, confirming deprotonation and anionic S,N‐chelation (Figures S5–S8) [24]. A medium‐intensity band near 500 cm⁻¹, attributed to ν _Ru—S, was present in all complexes and appeared more intense in the monodentate series [34, 45]. Electronic absorption spectra further distinguished the two coordination modes. Monodentate complexes exhibited two absorption bands at 258–262 nm (π → π*) and 323–328 nm (n → π*) (Figure S9) [42]. The bidentate complexes displayed an additional lower‐energy band at 354 nm attributable to a ligand‐to‐metal charge transfer (LMCT) transition arising from the anionic chelate, along with π → π* and n → π* bands at 252 and 285 nm (Figure S10) [47].

¹H NMR spectroscopy proved particularly diagnostic (Figure 1). The monodentate complexes displayed two distinct downfield —NH signals (Figures S11−S14) at 11.17–11.27 ppm (amide) and 12.54–12.89 ppm (thioamide), whereas the bidentate complexes showed only the thioamide —NH resonance at 12.51–12.78 ppm, confirming selective deprotonation of the acyl amide proton upon chelation (Figures S15−S18). The aromatic regions were largely similar, but significant differences appeared in the p‐cymene protons. Monodentate complexes exhibited two doublets (5.40–5.15 ppm) for the arene ring and a single doublet (1.24–1.30 ppm) for the isopropyl methyl groups, reflecting high symmetry. In the bidentate complexes, the reduced molecular symmetry induced by the strained six‐membered chelate ring splits the arene signals into three or four doublets (5.29–4.39 ppm) and the isopropyl methyl resonance into two distinct doublets (∼1.15 and 1.17 ppm), consistent with literature precedents for distorted piano‐stool geometries [29]. Figure 1 highlights the differences in —NH and p‐cymene proton environments between the mono‐ and bidentate complexes, as well as the free ligand and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ precursor.

FIGURE 1.

Stacked ¹H NMR spectra (selected regions) of (a) bidentate complex 1b, (b) monodentate complex 1m, (c) free ligand L ₁ , and (d) the dimeric precursor [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂. Key diagnostic features include the disappearance of the amide —NH signal upon S,N‐chelation, the persistence of both —NH signals in the neutral monodentate form, and the characteristic splitting of p‐cymene arene protons induced by the reduced symmetry of the bidentate chelate.

The ¹³C NMR spectroscopy provided additional confirmation of the coordination mode (Figures 2 and S19–S26). In monodentate complexes, the carbonyl and thiocarbonyl carbons resonated at 169.34–174.21 ppm (C=O) and 179.85–180.08 ppm (C=S), respectively. Upon S,N‐chelation, both signals shifted markedly downfield; the C=O signal shifted to 179.01–181.18 ppm, and the C=S signal shifted to 188.09–188.76 ppm, indicating deshielding due to electron withdrawal by the metal through both nitrogen and sulfur donors. This pattern contrasts with reported S,O‐chelated acylthiourea systems, where only the carbonyl carbon experiences significant deshielding, further validating exclusive S,N‐chelation in the bidentate series [26, 48]. Figure 2 presents a comparative ¹³C NMR analysis of the free ligand and its corresponding mono‐ and bidentate complexes, clearly illustrating these shifts.

FIGURE 2.

Comparative ¹³C NMR spectra (carbonyl and thiocarbonyl region) of the free ligand L ₁  [38], monodentate complex 1m, and bidentate complex 1b, illustrating the pronounced downfield shifts of both C=O (Δδ ≈ 179.01 ppm) and C=S (Δδ ≈ 188.38 ppm) upon formation of the anionic S,N‐chelate, diagnostic of bidentate coordination versus minimal shifts in the neutral S‐monodentate complex.

2.3. Solid‐State Structures of the Complexes

Single crystals suitable for X‐ray diffraction analysis were obtained by slow diffusion of diethyl ether into dichloromethane solutions of the complexes. Crystallographic data, including unit cell parameters, selected bond lengths, and bond angles, are summarized in Tables S1–S3. The molecular structures of the newly synthesized complexes 1m, 2b, and 4b, along with the previously reported 3m and 3b [28], are depicted in Figure 3, with their atom labeling schemes. As expected for half‐sandwich Ru(II)–arene complexes, all compounds adopt a pseudo‐octahedral “piano‐stool” geometry, with the η⁶‐bound p‐cymene ligand occupying the seat and the three remaining donor atoms (Cl, S, and/or N donors) forming the legs [40, 49]. In monodentate complexes (1m and 3m), the ruthenium center is coordinated by one thiocarbonyl sulfur donor and two chloride ligands. In contrast, the bidentate complexes (2b, 3b, and 4b) exhibit κ²‐S,N‐chelation from the thioamide ligand together with a single chloride anion, thereby forming a five‐membered metallacyclic ring. Upon coordination, elongation of the C—S bond is observed, with C=S distances increasing 1.656–1.672 Å in the free ligands to 2.404–2.423 Å in monodentate and 2.392–2.403 Å in bidentate (S,N) complexes, reflecting coordination‐induced elongation and Ru—S bonding interactions [38].

FIGURE 3.

Crystal structure of the complexes 1m, 2b, 3m, 3b, and 4b.

Examination of the solid‐state structures reveals distinct intramolecular hydrogen‐bonding patterns that correlate strongly with the observed coordination mode. In the monodentate complexes, two independent N—H···O and N—H···Cl hydrogen bonds are present: one between the thioamide N—H and the carbonyl oxygen of the same ligand, and another involving the amide N—H of the acyl fragment and a coordinated chloride. In the bidentate complexes, only the N—H(thioamide)···O=C hydrogen bond is observed, as the second potential N—H donor participates in chelation through nitrogen. These hydrogen‐bonding interactions appear to dictate the preferred chelation mode of the thioamide ligands. Primary thioamides bearing an N—H proton capable of forming a strong intramolecular N—H···O=C contact favor S,N‐bidentate coordination, which generates a stable six‐membered pseudo‐chelate ring involving the hydrogen bond [42]. This arrangement effectively blocks the carbonyl oxygen from metal coordination by satisfying its hydrogen‐bond acceptor role internally and introducing steric encumbrance around the oxygen atom. In contrast, secondary thioamides lacking the thioamide N—H proton are unable to form this stabilizing interaction and typically adopt alternative binding modes (e.g., O,S‐chelation) in related systems [33, 34, 50, 53]. To understand the intermolecular interactions in monodentate and bidentate complexes, one representative monodentate complex (1m) and one bidentate complex (2b) were selected for detailed investigation. The packing diagram of complex 1m (Figure S27) reveals weak van der Waals interactions between the chlorido ligand of one molecule and an aromatic hydrogen atom of a neighboring molecule. Additional stabilization of the crystal packing arises from weak van der Waals contacts between the thiocarbonyl sulfur atom and the hydrogen atoms of the p‐cymene moiety. In contrast, the crystal packing of complex 2b (Figure S28) is stabilized by van der Waals interactions involving contacts between the chlorido ligands and piperazine hydrogen atoms, as well as between the aromatic carbon adjacent to the acyl group and piperazine hydrogen atoms. A similar observation was also observed in their complexes 3m and 3b [28]. The present results thus underscore the critical role of intramolecular hydrogen bonding in controlling the coordination chemistry of multifunctional thioamide ligands at the Ru(II)–arene center.

2.4. Stability Test

The stability of representative complexes 2m and 2b was evaluated by ¹H NMR spectroscopy in DMSO‐d ₆ over 72 h at room temperature. The neutral monodentate complex 2m exhibited excellent stability, with no detectable decomposition or isomerization observed throughout the entire period (Figure S29). In contrast, the anionic bidentate complex 2b underwent partial conversion to the monodentate form within the first 10 min of dissolution (Figure S30), consistent with literature reports on related Ru(II)–acylthiourea systems in DMSO‐d ₆ [42]. This rapid initial equilibrium (approximately 7% conversion) is likely due to proton abstraction by trace water or residual acidity in the solvent, followed by slower subsequent kinetics. After 12 h, weak signals corresponding to the free ligand became detectable. However, even after 72 h, the bidentate S,N‐chelate species 2b remained the dominant form (ca. 74%), indicating that, despite the reversible nature of the coordination mode, the S,N‐chelated form retains sufficient kinetic stability under biologically relevant conditions to reach its cellular targets intact. These findings highlight the dynamic yet controllable coordination behavior of this ligand class.

Given the pronounced hydrophobic character of complexes 2m and 2b, ¹H NMR spectra recorded in DMSO‐d ₆ and D₂O at higher concentrations displayed broadened and poorly resolved signals, which hindered reliable peak assignment. Although partial improvement in resolution was achieved at lower concentrations, UV–vis spectrophotometry proved more suitable for examining the solution behavior of these complexes. Accordingly, the stability of complexes 2m and 2b in a DMSO/H₂O (5:5, v/v) mixture was studied using UV–vis spectroscopy in the 200–800 nm range (Figures S31–S32). The spectra displayed characteristic absorption bands that remained unchanged in both position and intensity over time, indicating that the complexes are stable in the mixed solvent system and do not have significant structural degradation or decomposition under the studied conditions [54, 55]. To assess their stability under biologically relevant conditions, complexes 2m and 2b were examined in Dulbecco's modified Eagle medium/Ham's F‐12 (DMEM/F12) medium over a 24‐h period. UV–vis spectroscopic monitoring showed no appreciable changes in the absorption profiles, indicating the preservation of their structural integrity under these conditions [39]. These results demonstrate that the complexes exhibit adequate stability to support subsequent biological investigations. The corresponding spectra are provided in Figures S33–S34.

2.5. Interconversion Between Monodentate and Bidentate Coordination Modes: In Situ ¹H NMR Studies

The switching between monodentate (κ¹‐S) and bidentate (κ²‐S,N) coordination modes was monitored by ¹H NMR spectroscopy in CDCl₃. The conversion of the monodentate complex 1m to its bidentate counterpart 1b was achieved by incremental addition of diisopropylamine (DIPA) to a solution of 1m (Figure 4; full stacked spectra in Figure S35). The reaction proceeded cleanly, as shown by the complete disappearance of the acylamide N—H resonance of 1m and the simultaneous emergence of diagnostic signals for 1b. Notably, the two doublets attributed to the p‐cymene protons in 1m were transformed into four distinct doublets, consistent with the formation of the bidentate 1b species. Additionally, the septet signals corresponding to DIPA underwent a downfield shift upon protonation, merged with piperazine peaks, confirming its role as a base. A parallel transformation was observed for 3m to 3b upon the addition of triethylamine (TEA) as the base [28]. In both cases, deprotonation of the acylamide N—H functionality emerges as the key step that unlocks S,N‐chelation, highlighting the broad applicability of this base‐promoted binding mode transformation across the present ligand series.

FIGURE 4.

¹H NMR spectral changes illustrating the base‐induced transformation of 1m to 1b upon titration with DIPA dissolved in CDCl₃ over the range of 0.2–1.2 equivalents.

The reverse process, bidentate‐to‐monodentate conversion, was affected by the addition of HCl to CDCl₃ solutions of the S,N‐chelated complexes (exemplified by 1b to 1m in Figure 5; full spectra in Figure S36). Protonation promptly restored the amide —NH signal and collapsed the four arene doublets of 1b into the original two‐doublet pattern of 1m, fully regenerating the monodentate coordination mode. An analogous reversal was also observed for the 3b to 3m transformation [28]. These experiments demonstrate that the mono‐ and bidentate forms are interconvertible under mild acid–base control, with the acylamide N—H acting as a reversible “coordination switch” that governs access to the nitrogen donor and, consequently, the overall binding mode of these multifunctional thioamide ligands.

FIGURE 5.

¹H NMR spectral changes illustrating the HCl‐induced transformation of 1b to 1m upon titration with HCl (1.25 M in MeOH, added in CDCl₃) over the range of 0.2–1.6 equivalents.

2.6. Coordination Mode Interconversion Monitored by UV–Vis Spectrophotometry

The switching between monodentate and bidentate coordination modes was further probed by UV–vis absorption spectroscopy in dichloromethane ([Ru] ≈ 1 × 10⁻⁵ M). Titration of the monodentate complex 1m with diisopropylamine (0.2–1.0 equiv) induced well‐defined spectral changes (Figure 6a). A new shoulder emerged at 285 nm, accompanied by a pronounced blueshift of the absorption maximum from 260 to 250 nm (Figure 6a). These observations indicate a ligand coordination mode transition from monodentate to bidentate, facilitated by the base‐induced deprotonation of the amide hydrogen. In contrast, the reverse transformation of 1b to 1m was examined by incremental addition of a 1.25 M HCl solution in dichloromethane. Progressive acidification resulted in the disappearance of the characteristic shoulder peak associated with the bidentate complex, while the absorption maximum at 250 nm exhibited a redshift back to 260 nm, consistent with reprotonation of the amide moiety and re‐establishment of the monodentate coordination (Figure 6b).

FIGURE 6.

UV–vis spectral changes showing (a) the base‐induced conversion of 1m to 1b upon incremental addition of diisopropylamine (0.2–1.0 eq.) and (b) the acid‐induced reverse of 1b to 1m upon titration with 1.2 eq. of HCl solution.

Notably, the base‐mediated conversion (1m → 1b) occurred at lower equivalents of reagent compared to the acid‐mediated reverse process (1b → 1m), underscoring the greater efficiency of the deprotonation pathway. A similar trend was observed for the interconversion between complexes 3m and 3b, exhibiting analogous spectral shifts and coordination mode changes [28].

2.7. Partition Coefficient (log P) and Lipophilicity Assessment

The lipophilicity of monodentate complexes 1m–4m and bidentate complexes 1b–4b, an important determinant of pharmacokinetic and membrane transport behavior, was evaluated by determining their partition coefficients (log P) using the octanol/water model system. Lipophilicity is closely associated with drug absorption, membrane permeability, intracellular distribution, and receptor accessibility in biological environments [56, 57]. Admirable linear correlations were obtained for all complexes in MeOH/water and n‐octanol systems, with regression coefficients (R ² > 0.98) as illustrated in Figures S45–S52, while the corresponding absorption wavelengths are summarized in Table S8.

The experimentally determined log P values for monodentate complexes (1m–4m) were calculated as 3.49 ± 0.02, 4.21 ± 0.03, 4.41 ± 0.01, and 4.47 ± 0.02, and for bidentate complexes (1b–4b) were calculated as 3.87 ± 0.01, 4.34 ± 0.02, 4.31 ± 0.02, and 4.91 ± 0.01, respectively. The bidentate forms show slightly higher log P values, probably due to less hydrogen‐bonding ability compared to the monodentate complexes, which contain two —NH groups in addition to the two chloride ligands. In general, compounds possessing log P values within the range of −0.04 to 5.60 are considered favorable for cellular uptake and bioaccumulation, indicating that these Ru(II) complexes possess physicochemical properties compatible with potential therapeutic applications [58, 59]. Structural variations within the acylthiourea system also influenced lipophilicity [60]. Incorporation of electron‐donating methyl substituents enhanced the lipophilic character. Furthermore, the positive log P values obtained for all complexes indicate preferential partitioning into the lipid phase [61], suggesting their capability to permeate biological membranes effectively. Overall, the lipophilicity trend followed the order 4b > 4m > 3m > 2b > 3b > 2m > 1b > 1m, highlighting the strong influence of ligand architecture on the physicochemical behavior of these Ru(II) systems.

2.8. DNA Binding and Fluorescence Quenching Studies

The interaction of the synthesized Ru(II)‐p‐cymene complexes with calf thymus DNA (CT‐DNA) was investigated using UV–vis absorption titration and ethidium bromide (EB) competitive fluorescence displacement assays. The intrinsic binding constants (K _b) were determined from the UV–vis titration data using the Wolfe–Shimer equation. As summarized in Tables 1 and S9 and Figure S53, the K _b values for the series ranged from 1.58 × 10⁵ to 12.01 × 10⁵ M⁻¹, indicating a strong affinity for the DNA secondary structure. Linear fitting of [DNA]/(ε _a−ε _f) versus [DNA] (Figure 7a) confirmed the reliability of the association. Among the complexes, the bidentate complex 2b exhibited the highest binding affinity (K _b = 12.01 × 10⁵ M⁻¹), suggesting that its specific S,N‐chelation mode and structural configuration were highly favorable for stabilizing the intercalation between DNA base pairs, which correlated with its superior cytotoxic profile.

TABLE 1.

Stern–Volmer quenching constant (K _sv) and apparent binding constant (K _app) values for complexes.

Compound	K b (M −1 )	K sv (M −1 )	K app (M −1 )
1m	3.03 × 105	0.97 × 104	4.85 × 105
2m	7.75 × 105	0.63 × 104	3.15 × 105
3m	1.58 × 105	0.88 × 104	4.40 × 105
4m	7.93 × 105	0.66 × 104	3.30 × 105
1b	2.66 × 105	1.02 × 104	5.10 × 105
2b	12.01 × 105	1.04 × 104	5.20 × 105
3b	2.82 × 105	0.85 × 104	4.25 × 105
4b	2.95 × 105	0.50 × 104	2.50 × 105

FIGURE 7.

Linear fitting plots for DNA binding studies of Ru(II) complexes 1m–4b. (a) [DNA]/(ε _a−ε _f) versus [DNA] for the ct‐DNA titration. (b) I ₀/I versus [Q] for the Stern–Volmer fluorescence quenching assay.

The ability of the complexes to compete with ethidium bromide for DNA binding sites was further evaluated using a fluorescence displacement assay. Incremental addition of each Ru(II) complex resulted in pronounced quenching of the characteristic EB‐DNA fluorescence emission. The Stern–Volmer quenching constants (K _SV), obtained from the linear slopes in Figure 7b, ranged from 0.50 × 10⁴ to 1.04 × 10⁴ M⁻¹. The corresponding apparent binding constants (K _app), calculated at an EB concentration of 5 μM, fell between 2.50 × 10⁵ and 5.20 × 10⁵ M⁻¹. These K _app values show excellent agreement with the K _b values obtained from absorption spectroscopy, validating the consistency of the binding results across both techniques. In general, the DNA binding mode of a compound can be distinguished from UV–vis spectral changes as follows: (i) intercalative binding, characterized by hypochromism along with a bathochromic shift, arising from strong π–π stacking interactions between the planar aromatic chromophore and DNA base pairs [62]; (ii) nonintercalative binding, including groove or electrostatic interactions, which typically results in hyperchromism due to perturbation of the DNA helix [63]; and (iii) groove binding, also indicated by hypochromism without a significant shift in the absorption maximum [64]. Collectively, the data demonstrate that the synthesized complexes exhibit high affinity for CT‐DNA and effectively displace EB from its intercalation sites, supporting a groove DNA‐binding mode that correlates with their observed anticancer potency [65, 66].

Furthermore, characteristic spectral changes during the interaction of the lead complex 2b with DNA are clearly illustrated in Figure 8. Upon the incremental addition of CT‐DNA, the electronic absorption spectra of 2b exhibited a progressive increase in absorbance at approximately 265 nm, accompanied by a slight blueshift, which was diagnostic of noncovalent association and electronic perturbations within the aromatic system upon binding [64, 67]. Correspondingly, Figure 8b shows that the intense fluorescence of the EB–DNA complex was gradually quenched as the concentration of 2b increased from 0 to 30 µM. These visual observations further substantiated that the synthesized complexes effectively competed for the DNA binding sites, providing a solid physicochemical rationale for the potent in vitro anticancer activities observed in this study.

FIGURE 8.

(a) Electronic absorption titration spectrum of 2b. [Compound] = 1.0 × 10⁻⁴ M, [DNA] = 0–70 µM. (b) Fluorescence quenching curves of EB bound to DNA in the presence of 2b. [DNA] = 5 µM, [EB] = 5 µM, and [compound] = 0–30 µM.

2.9. Biological and Anticancer Activity

The cytotoxic properties of the synthesized acylthiourea ligands and their corresponding Ru(II)‐p‐cymene complexes were evaluated against two human melanoma cell lines (A375 and A2058), with the human retinal epithelial cell line (ARPE‐19) serving as the normal control. Overall, metal coordination markedly enhanced the anticancer activity relative to the free ligands, indicating that complexation with Ru(II) centers plays a critical role in modulating cytotoxic efficacy (Table 2).

TABLE 2.

IC₅₀ values (72‐h incubation) of the acylthiourea ligands, their complexes, and cisplatin in A375, A2058 (melanoma), and ARPE‐19 (normal retinal) cell lines.

Compound	IC50 value ± SD (μM)
	A375	A2058	ARPE‐19
L 1	48.2 ± 2.5	59.0 ± 2.8	58.9 ± 3.1
L 2	65.7 ± 1.2	>100	>100
L 3	>100	88.5 ± 3.2	>100
L 4	49.0  ± 3.4	88.2 ± 3.4	62.6 ± 4.8
1m	48.3 ± 3.3	46.3 ± 4.2	>100
2m	36.4 ± 4.2	>100	>100
3m	55.0 ± 3.5	>100	>100
4m	33.4 ± 1.5	49.0 ± 2.2	47.7 ± 3.7
1b	30.7 ± 3.4	42.5 ± 3.5	>100
2b	6.1 ± 0.8	22.3 ± 2.1	55.0 ± 2.8
3b	33.7 ± 3.2	63.8 ± 3.5	67.9 ± 2.5
4b	22.7 ± 1.8	32.6 ± 2.2	25.2 ± 3.1
Cisplatin	20.5 ± 2.8	>100	>100

Structure–activity relationship (SAR) analysis revealed that bidentate complexes generally display improved cytotoxicity compared to their monodentate counterparts across the majority of ligand pairs, with the advantage being particularly pronounced for several derivatives (Figures 9 and S54). This trend supports the beneficial role of the rigid S,N‐chelated architecture in enhancing biological performance. In particular, complex 2b emerged as the lead compound of the series, exhibiting a pronounced sixfold increase in potency (IC₅₀ = 6.1 ± 0.8 μM in A375 cells) relative to its monodentate analog 2m (IC₅₀ = 36.4 ± 4.2 μM), while maintaining good selectivity toward melanoma cells over normal ARPE‐19 cells (IC₅₀ = 55.0 ± 2.8 μM). This derivative best exemplifies the combined advantages of the bidentate coordination mode and appropriate N‐aryl substitution. Complex 4b, while active against melanoma cells (IC₅₀ = 22.7 ± 1.8 μM in A375), also exhibited notable cytotoxicity toward normal ARPE‐19 cells (IC₅₀ = 25.2 ± 3.1 μM). This reduced selectivity is attributed to the strong electronic influence of its N‐aryl substituent, which may promote nonspecific interactions. Such behavior provides important insight for future ligand optimization aimed at improving the therapeutic window within this series.

FIGURE 9.

The cell viability of (a) melanoma A375 cells and (b) melanoma A2058 cells treated with complexes (0, 5, 10, 15, 25, 35, 50 μM) for 72 h.

These results highlight the significance of the bidentate coordination mode in enhancing the anticancer efficacy of Ru(II)‐p‐cymene complexes, particularly when paired with optimized ligand substitution. The notable activity of 2b likely reflects the synergistic effects of the Ru(II)–arene platform and the piperazine substituent, a motif associated with enhanced bioavailability [5]. Moreover, recent analyses emphasize that ligand optimization within Ru(II)–acylthiourea systems is critical for maximizing biological performance [10].

To evaluate the antiproliferative effects of complex 2b, morphological changes in A375 melanoma cells were examined after treatment with various concentrations (2.5, 5, and 10 μM) for 72 h (Figure 10). While untreated control cells displayed a normal polygonal morphology and high confluency, exposure to 2b induced a dose‐dependent reduction in cell density accompanied by cellular shrinkage and rounding. Specifically, at 10 μM, the majority of cells became sparse and lost adherence, indicating severe growth inhibition. To further assess the kinetics of this response, a time‐course study was performed at 10 μM (Figure S55), which revealed a progressive deterioration in cell viability from 24 to 72 h. These morphological observations established that 2b impaired melanoma cell viability in both a dose‐ and time‐dependent manner.

FIGURE 10.

(a) The cell morphology and (b) Western blot analysis of p21 and Cyclin B1 protein expression in A375 cells treated with 2b (0–10 μM) for 72 h.

To further elucidate the molecular mechanism underlying this growth inhibition, the expression levels of the cell cycle‐regulated proteins p21 and Cyclin B1 were investigated by Western blotting. As illustrated in Figure 10b, treatment with 2b induced a distinct, dose‐dependent modulation of these molecular markers. At a lower concentration of 5 μM, a noticeable upregulation of p21 (1.6‐fold relative to control), accompanied by a prominent accumulation of Cyclin B1 (1.8‐fold), was observed, reflecting an initial perturbation of the G2/M checkpoint regulatory pathway. Upon increasing the concentration to 10 μM, Cyclin B1 levels further accumulated to 3.2‐fold, whereas p21 expression returned near the baseline level. This biphasic response of p21 at the higher concentration likely correlates with the severe cellular stress and the active onset of apoptosis observed in the morphological analysis, where advanced cell death often leads to the decline of early stress‐responsive regulators. This sustained impairment of cell viability likely stems from the interference with essential proliferative pathways through the robust DNA groove binding established in the previous sections, providing a cellular context for the enhanced potency of the bidentate species. This pattern was consistent with the disruption of replication and transcription through DNA groove binding, a recognized mechanism for Ru(II)–arene complexes [68].

To further explore how the lead complex 2b exerted its anticancer effects, a wound‐healing assay was performed on A375 melanoma cells to evaluate the inhibition of cell migratory capacity. Treatment with 5 μM 2b significantly delayed wound healing compared to the untreated control group, with a clear scratch gap remaining after 24 h (Figure 11a). Quantitative analysis confirmed a dose‐dependent inhibitory effect, which was supported by the migration rates being approximately 84% in controls versus approximately 54% in the 5 μM treatment group after 24 h (p < 0.05) (Figure 11b). To further elucidate the underlying molecular mechanism, the expression level of Matrix Metalloproteinase‐9 (MMP‐9), a key enzyme involved in cancer cell migration and invasion, was examined by Western blotting. As shown in Figure 11c, treatment with 2b resulted in a profound, dose‐dependent decrease in MMP‐9 protein levels, with the relative expression sharply dropping to 0.2 at the effective concentration of 5 μM.

FIGURE 11.

(a) The cell migration effects, (b) quantitative analysis, and (c) Western blot analysis of MMP‐9 protein expression in A375 melanoma skin cancer cells treated with compound 2b.

This potent antimigratory capability correlates closely with the high DNA‐binding affinity of 2b (K _b = 12.01 × 10⁵ M⁻¹). These results suggest that 2b may suppress melanoma cell migration and proliferation primarily by interfering with critical signaling pathways or gene expressions through a robust DNA groove‐binding mode, thereby suppressing the spread and proliferation of melanoma cells [69]. This finding not only explained the superior cellular activity of 2b but also reaffirmed the core role of the bidentate coordination mode in enhancing the biological efficacy of ruthenium(II)–arene complexes.

2.10. In Silico Studies

2.10.1. Structural Optimization and Energy‐Level Calculations

Based on the data in Table S5, the optimized structures of compounds 1m–4m and 1b–4b were obtained using the DMol³ module in Materials Studio, with all energies reported in Hartree (Ha). The total energy reflects the overall thermodynamic stability of each optimized structure. Among the complexes’ series, 4m (−3141.617 Ha) exhibited the lowest total energy, indicating the highest stability, while 1b with −2564.062 Ha was comparatively less stable.

The HOMO–LUMO energy gaps (ΔE) for compounds 1m–4m and 1b–4b were calculated as 2.544, 2.578, 2.662, 2.530, 2.460, 1.990, 2.663, and 2.315 eV, respectively. Among these, 3m exhibited the largest gap (2.662 eV), consistent with higher kinetic stability and reduced reactivity, whereas 2b showed the smallest gap (1.990 eV), indicating comparatively enhanced chemical reactivity (Table S6). For complexes 1m–3m and all bidentate analogs, the HOMO electron density is largely delocalized over the Ru center, chloride ligands, the thiourea sulfur atom, and the p‐cymene framework. In contrast, complex 4m shows a distinct LUMO distribution that is predominantly located on the mesitylene moiety. For the monodentate series, the LUMO is mainly distributed over the acylthiourea fragment with additional contribution from the Ru center. In the bidentate series, the LUMO is localized on different ligand regions depending on structure: in 1b and 2b, it is concentrated on the carbonyl group and the carbonyl‐linked phenyl unit; in 3b, it is mainly localized on the piperazine fragment; and in 4b, it is primarily distributed over the acylthiourea moiety (Figure S56).

Chemical hardness (η) and chemical softness (s) are important descriptors of relative stability, reflecting a compound's resistance to changes in electron density and electron‐transfer processes. As shown in Table S6, complex 2b exhibits the lowest hardness and highest softness, indicating comparatively lower stability relative to the other complexes. A lower electrophilicity index (ω) corresponds to greater nucleophilicity, whereas higher ω values indicate stronger electrophilic character. Accordingly, compounds can be classified as strong (ω > 1.5 eV), moderate (0.8–1.5 eV), or weak (ω < 0.8 eV) electrophiles. On this basis, all synthesized compounds, particularly 2b, fall within the strong electrophile category, in line with their elevated ω values and implying a greater tendency to interact with biomolecular targets. Moreover, a reduced HOMO–LUMO energy gap is associated with increased metal–ligand orbital mixing and enhanced metal‐to‐ligand electron donation, which can further accentuate the electrophilic character of the metal center [70]. In conclusion, from an electronic structure perspective, molecules with lower HOMO–LUMO energy gaps exhibit greater chemical softness and electrophilicity, facilitating stronger interactions with biological targets such as DNA and proteins. This enhanced biomolecular interaction may account for the superior cytotoxic activity observed for complex 2b compared to its monodentate counterpart.

2.11. Molecular Docking Investigation With dsDNA

To further discuss the binding model of the interaction between the complexes and DNA, a molecular docking study was performed for the representative complex 2b with duplex B‐DNA (PDB ID:1AIO). The docking results showed that complex 2b preferentially occupied the major groove of B‐DNA. As illustrated in Figure 12, 2b adopted a crescent‐shaped conformation that matched the helical curve of the DNA, and this geometric complementarity allowed it to stabilize effectively within the DNA structure. Detailed interaction analysis revealed that the binding stability of 2b was provided by a key hydrogen bond with the DA17 residue and extensive van der Waals contacts with neighboring nucleotides, including DT5, DG16, DC18, DC19, and DA20. To experimentally validate this major‐groove binding mode, a fluorescence competitive displacement assay using methyl green (MG), a well‐known major‐groove binder, was performed. As shown in Figure 12d, the fluorescence intensity of the MG–dsDNA complex gradually decreased upon the addition of 2b at ratios of 1:1 and 1:2, indicating that 2b successfully displaced MG from the major groove of DNA.

FIGURE 12.

Molecular docking model and fluorescence displacement assay of 2b binding to dsDNA (PDB: 1AIO). (a) Overall and (b) local docking structures of the 2b‐dsDNA complex. (c) LigPlot+ analysis showing interactive networks, where green dashed lines and red spoked arcs represent hydrogen bonds and hydrophobic interactions, respectively. (d) Fluorescence emission spectra of the MG–dsDNA complex in the presence of 2b (0–10 μM; [MG] = 5 μM) at ratios of 1:0, 1:1, and 1:2, demonstrating competitive major‐groove binding.

These computational findings were highly consistent with the high binding constants (K _b and K _app) measured in the titration and displacement assays, confirming that 2b interacted strongly with DNA through noncovalent groove binding. Meanwhile, this binding mode was consistent with reported Ru(II)–DNA interactions, in which hydrogen‐bonding and hydrophobic contacts facilitate major‐groove association [33]. Moreover, noncovalent groove binding, distinct from cisplatin‐like covalent cross‐linking, represents a recognized mechanism for Ru(II)–arene anticancer complexes [71], including acylthiourea derivatives demonstrating pronounced DNA affinity [72]. In this regard, the integration of the piperazine scaffold represents a pivotal architectural design element that dictates both biomolecular recognition and pharmacokinetic performance. While its basic nitrogen centers actively support localized B‐DNA major‐groove pocket association via robust electrostatic interactions, they simultaneously govern key systemic properties, including aqueous solubility and cellular accumulation, effectively translating structural rigidity into enhanced antimelanoma potency.

2.12. ADMET Studies

To theoretically validate the aforementioned physicochemical benefits imparted by the piperazine ring, SwissADME was utilized to evaluate the ADMET profiles of all complexes, including parameters such as hydrogen‐bond donor/acceptor capacity (Table S7), BBB permeability, absorption, TPSA, hepatotoxicity, and aqueous solubility. The Ru(II) complexes investigated in this study did not fully satisfy Lipinski's rule due to their higher molecular weights and showed a low predicted bioavailability score (0.17). Nevertheless, they complied with most Veber parameters, including log P ≤ 5, TPSA ≤ 140 Ų, hydrogen‐bond acceptors ≤ 10, and hydrogen‐bond donors ≤ 5, despite exceeding the molecular weight criterion [73]. For intestinal absorption, Log P values should be below 5.6, a criterion satisfied by all complexes, indicating favorable lipophilicity. In general, the bidentate complexes exhibited slightly higher lipophilicity compared with the corresponding monodentate complexes. The %ABS values were calculated using the equation 109 − (0.3345 × TPSA). All complexes exhibited predicted %ABS values higher than 81%, indicating favorable intestinal absorption and good oral absorption potential. Furthermore, the bidentate complexes showed slightly higher absorption values than the corresponding monodentate complexes [74]. The predicted log K _p values (−3.18 to −4.17 cm/s) indicate moderate skin permeability for all compounds, with compound 4m showing the highest permeation potential and compound 2b the lowest [75].

3. Conclusions

This study presents a detailed investigation into the coordination behavior of acylthiourea ligands in Ru(II) half‐sandwich complexes, highlighting the reversible modulation between monodentate and bidentate binding modes under acid–base control. FT‐IR spectroscopy confirmed neutral monodentate coordination via the thiocarbonyl sulfur atom, while redshifts in the C=O and C=S stretching frequencies in bidentate complexes indicated S,N‐chelation following amide deprotonation. ¹H and ¹³C NMR spectroscopy served as powerful tools for distinguishing coordination environments, with pronounced chemical shift changes in —NH, carbonyl, and p‐cymene proton resonances reflecting ligand deprotonation, chelation, and increased structural asymmetry. In situ NMR tube experiments demonstrated base‐promoted conversion from monodentate to bidentate coordination, while subsequent acid treatment reversed this process, confirming the dynamic, pH‐responsive nature of the system. X‐ray crystallographic analysis of representative complexes further corroborated the proposed coordination modes, revealing characteristic bond lengths and angular distortions associated with chelate formation. Notably, this is the first systematic study to establish reversible acid–base switching between mono‐ and bidentate coordination in acylthiourea–Ru(II) complexes. Our results demonstrate that bidentate coordination is promoted under basic conditions via amide deprotonation, whereas acidic conditions restore monodentate sulfur coordination through reprotonation. This reversible behavior highlights the dynamic and tunable coordination properties of acylthiourea ligands in transition metal complexes. These findings not only deepen the understanding of acylthiourea coordination chemistry but also introduce a versatile, stimulus‐responsive platform with potential applications in catalysis, molecular recognition, and bioinorganic system design. The switchable coordination modes of the Ru(II)‐p‐cymene complexes are central to their biological performance. The bidentate coordination motif, driven by the chelate effect, generates a stable, crescent‐shaped geometry that enables precise geometric complementarity with the B‐DNA major groove. This structural rigidity is essential for maximizing noncovalent binding affinity and selective cytotoxic activity. In contrast, the monodentate coordination mode offers less optimal geometric alignment, which correlates with the reduced anticancer potency observed across the series. Thus, the acid–base‐mediated coordination switch acts as a molecular control element, allowing for the modulation of target affinity and therapeutic index. It has been shown that metal coordination significantly improved the cytotoxicity of the ligands, with bidentate complexes outperforming their monodentate counterparts. Complex 2b showed a sixfold increase in potency over 2m and demonstrated good selectivity toward cancer cells. Docking results suggest that its enhanced activity arises from DNA groove binding, highlighting the crucial role of the bidentate coordination mode in boosting the anticancer efficacy of Ru(II)‐p‐cymene complexes. Computational calculations, including quantum‐chemical descriptors, indicate that complex 2b exhibits the highest electrophilicity and the strongest propensity for interaction with biomolecular targets.

4. Experimental Section

4.1. Material and Methods

The ligands, along with their respective mono‐ and bidentate Ru(II) complexes, were synthesized under ambient conditions. All solvents utilized in the synthesis were subjected to purification protocols in accordance with standard procedures prior to their application. The reagents employed were of analytical or reagent grade and were utilized without any further purification. Ruthenium(III) chloride trihydrate (RuCl₃·3H₂O), α‐phellandrene, and other organic precursors required for ligand synthesis, as well as bases, were obtained from Sigma‐Aldrich. The compound 4‐(4‐phenylpiperazin‐1‐yl)aniline was synthesized utilizing a method documented in the existing literature [76]. Fargo MP‐10 melting point apparatus was used to determine the melting point of the synthesized complexes. FT‐IR spectra were recorded on a Bruker Alpha OPUS spectrophotometer using KBr pellets. The absorption spectra of the complexes were measured on an Agilent 8453 UV‐vis spectrophotometer. ¹H/¹³C NMR spectra were obtained on a JEOL 400 MHz spectrometer, with tetramethylsilane (TMS) as the internal reference. Elemental (CHNS) analyses were achieved using a UNICUBE elemental analyzer. ESI‐MS was carried out on a Waters ZQ 4000 mass spectrometer to determine the molecular ion mass of the complexes.

4.2. Synthesis and Characterization

4.2.1. Synthesis of Monodentate Complexes

In order to prepare 1m–4m, the acylthiourea ligands and the ruthenium precursor, bis[dichlorido(η⁶‐p‐cymene)ruthenium(II)], were synthesized following previously reported literature [38, 77]. A toluene solution of the ligand (2 mmol in 50 mL) was prepared and added dropwise to a suspension of [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (1 mmol in 50 mL toluene) under constant stirring. The reaction mixture was stirred at 50–60 °C for 6 h and then allowed to cool to room temperature without disturbance. The resulting mixture was decanted, and the precipitated orange solid was washed with diethyl ether (2 × 30 mL), followed by drying under vacuum at 80 °C for 3 h. The obtained complexes were characterized by various analytical techniques, including ¹H/¹³C NMR, ESI‐MS, FT‐IR, and UV–vis spectroscopy. Single crystals suitable for X‐ray diffraction were obtained for complexes 1m and 3m by slow diffusion of diethyl ether into a dichloromethane solution at room temperature. The general procedure is presented in Scheme 2.

4.2.2. Synthesis of Bidentate Complexes

In order to prepare 1b–4b, to a methanolic suspension of the ligand (2 mmol in 50 mL), the corresponding base (2 mmol), 1,1,3,3‐tetramethylguanidine (TMG), N,N,N′,N′‐tetramethylnaphthalene‐1,8‐diamine (PS), 2,2,6,6‐tetramethylpiperidine (TMP), diisopropylamine (DIPA), triethylamine (TEA), sodium bicarbonate (NaHCO₃), 2,4‐dimethylpyridine (DMP), or pyridine (Py), was added and stirred for 5 min at room temperature. Subsequently, a methanolic solution of [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (1 mmol in 50 mL) was added dropwise under continuous stirring. The reaction mixture was stirred at room temperature for 6 h. Consequently, the reaction mixture was allowed to stand, the solvent was decanted, and the resulting yellow‐orange precipitate was washed with diethyl ether (2 × 30 mL). The crude product was dried under vacuum at 70 °C for 3 h. The obtained complexes were characterized by various analytical techniques, including ¹H/¹³C NMR, ESI‐MS, FT‐IR, and UV–vis spectroscopy. Single crystals suitable for X‐ray diffraction were obtained for complexes 2b, 3b, and 4b by slow diffusion of diethyl ether into a dichloromethane solution at room temperature. The general procedure is presented in Scheme 2.

4.2.3. Interconversion of Mono‐/Bidentate Complexes

Apart from independent synthesis of mono‐/bidentate complexes, we are also able to interconvert mono‐/bidentate complexes. The preparations of bidentate complexes were obtained by reacting equimolar amounts of the monodentate complexes with the corresponding organic bases (Table S4). The reactions were carried out in methanol at room temperature and stirred for 4 h. The resulting products were isolated and purified using the same procedure as described for route 1. The conversion of 1m to 1b was evaluated using eight different organic and inorganic bases. No consistent correlation was observed between the base pK _a values and the isolated yields of the product. The variation in yield is likely influenced by differences in the work‐up process, particularly during the washing step. Among the bases tested, NaHCO₃ afforded the highest yield, which can be attributed to the facile removal of byproducts from the reaction mixture. The complete list of bases, along with their respective pK _a values and product yields, is summarized in Table S4. The transformation from monodentate to bidentate coordination was further confirmed by in situ ¹H NMR studies. In a typical experiment, a deuterated chloroform solution of monodentate complex 1m was mixed with a deuterated chloroform solution of the selected organic base in an NMR tube. The progress of the reaction was monitored at room temperature over a period of 2 h. The general synthetic route to mono/bidentate complexes (route 2) is illustrated in Scheme 2. The conversion of bidentate complexes to their monodentate counterparts was successfully achieved by treating a methanolic suspension of the bidentate complexes with a 1.25 M methanolic solution under ambient conditions for 4 h. Attempts to induce this transformation using various hydrochloride salts, including pyridinium hydrochloride, 4‐bromopyridinium hydrochloride, triethylammonium hydrochloride, and 2‐(chloromethyl) pyridine hydrochloride, were unsuccessful under similar conditions.

4.2.3.1. L₁RuCl₂‐m (1m)

Complex 1m was synthesized following the general procedure for the synthesis of monodentate complexes by using L ₁ (0.833 g, 2 mmol) and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Orange color solid. Yield: 1.155 g, 80%. M.p.: 214 °C. Anal. Calcd. for C₃₄H₃₈Cl₂N₄ORuS (%): C, 56.50; H, 5.30; N, 7.75. Found: C, 56.68; H, 5.55; N, 7.72. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 258 (22698), 328 (5454). FT‐IR (KBr, cm⁻¹): 3229 (m; ν(amide N—H)), 3041 (m; ν(thiourea N—H)), 1666 (s; ν(C=O)), 1338 (s; ν(C—N), 1226 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.29 (d, J = 8.0 Hz, 6H, H‐19/H‐19´), 2.24 (s, 3H, H‐18), 2.91–3.02 (Sep, J = 8 Hz, 1H, H‐17), 3.38–3.45 (m, 8H, H‐13–H‐16), 5.23 (d, J = 6.0 Hz, 2H, H‐12/H‐12´), 5.39 (d, J = 6 Hz, 2H, H‐11/H‐11´), 6.93 (t, J = 7.2 Hz, 1H, H‐10), 7.00 (d, J = 7.2 Hz, 2H, H‐9/H‐9´), 7.03 (d, J = 7.2 Hz, 2H, H‐8/H‐8´), 7.32 (t, J = 7.2 Hz, 2H, H‐7/H‐7´), 7.39 (d, J = 8.0, 2H, H‐6/H‐6´), 7.47 (t, J = 8.0 Hz, 2H, H‐5/H‐5´), 7.55 (t, J = 8.0 Hz, 1H, H‐4), 8.28 (d, J = 8.2 Hz, 2H, H‐3/H‐3´), 11.27 ppm (s, 1H, NH‐H2), 12.82 ppm (s, 1H, NH‐H1). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 180.02 ( C =S), 169.46 ( C =O), 150.65, 150.36, 130.09, 129.41, 128.32, 128.17, 127.42, 127.00, 124.65, 116.74, 116.07, 103.51, 99.77 (aromatic rings), 84.26, 82.96 (p‐cymene), 49.68, 48.86 (CH₂ carbons of piperazine), 30.48, 29.05 (‐ C H‐(CH₃)₂), 23.16, 22.33 (‐CH‐( C H₃)₂), 18.45 (cymene‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M –HCl, Cl^‐]⁺/found: 651.18/ 651.26 (Figure S37).

4.2.3.2. L₂RuCl₂‐m (2m)

Complex 2m was synthesized following the general procedure for the synthesis of monodentate complexes by using L ₂ (0.861 g, 2 mmol) and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Orange color solid. Yield: 1.148 g, 78%. M.p.: 232 °C. Anal. Calcd. for C₃₅H₄₀Cl₂N₄ORuS (%): C, 57.06; H, 5.47; N, 7.60. Found: C, 56.69; H, 5.53; N, 7.68. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 262 (37700), 328 (9330). FT‐IR (KBr, cm⁻¹): 3202 (m; ν(amide N—H)), 3037 (m; ν(thiourea N—H)), 1712 (s; ν(C=O)), 1335 (s; ν(C—N), 1223 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.29 (d, J = 8 Hz, 6H, H‐19/H‐19´), 2.23 (s, 3H, H‐18), 2.38 (s, 3H, H‐17), 2.91–3.01 (Sep, J = 8 Hz, 1H, H‐16), 3.37–3.45 (m, 8H, H‐12–H‐15), 5.22 (d, J = 8 Hz, 2H, H‐11/H‐11´), 5.38 (d, J = 8 Hz, 2H, H‐10/H‐10´), 6.93 (t, J = 8.0 Hz, 1H, H‐9), 7.00–7.03 (d, J = 8 Hz, 4H, H‐7,7´/H‐8,8´), 7.27 (d, J = 8.0 Hz, 2H, H6‐/H‐6´), 7.32 (t, J = 8.0 Hz, 2H, H‐5/H‐5´), 8.16 (d, J = 8.0 Hz, 2H, H‐4/H‐4´), 7.47 (d, J = 8.0 Hz, 2H, H‐3/H‐3´), 11.19 ppm (s, 1H, NH‐2H), 12.85 ppm (s, 1H, ‐NH‐1H). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 180.08 ( C =S), 169.54 ( C =O), 144.62, 130.02, 129.61, 129.39, 128.61, 128.46, 127.05, 124.71, 116.76, 116.13, 103.58, 99.91, 99.79 (aromatic rings), 84.30, 82.99 (p‐cymene), 48.91 (CH₂ carbons of piperazine), 30.52 (‐ C H‐(CH₃)₂), 22.36, 21.86 (‐CH‐(CH₃)₂), 18.84 (benzene ring‐CH₃), 18.44 (cymene–CH₃). ESI ⁺ ‐MS (MeOH): cal [M –HCl, Cl^‐]⁺/found: 665.19/665.34 (Figure S38).

4.2.3.3. L₃RuCl₂‐m (3m)

Complex 3m was synthesized following the general procedure for the synthesis of monodentate complexes by using L ₃ (0.889 g, 2 mmol) and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Orange color solid. Yield: 1.185 g, 79%. M.p.: 230 °C. Anal. Calcd. for C₃₆H₄₂Cl₂N₄ORuS (%):C, 57.59; H, 5.64; N, 7.46. Found: C, 57.54; H, 5.46; N, 7.41. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 262 (42592), 328 (10686). FT‐IR (KBr, cm⁻¹): 3202 (m; ν(amide N—H)), 3040 (m; ν(thiourea N—H)), 1717 (s; ν(C=O)), 1322 (s; ν(C—N), 1234 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.23 (t, J = 8.0 Hz, 3H, H‐20), 1.29 (d, J = 8.0 Hz, 6H, H‐19/H‐19´), 2.23 (s, 3H, H‐18), 2.65 (q, J = 8.0 Hz, 2H, H‐17), 2.91–3.02 (Sep, J = 8.0 Hz, 1H, H‐16), 3.35–3.47 (m, 8H, H‐12–H‐15), 5.23 (d, J = 6.0 2H, H‐11/H‐11´), 5.38 (d, J = 6.0 2H, H‐10/H‐10´), 6.93 (t, J = 8.0 Hz, 1H, H‐9), 7.00–7.03 (m, 4H, H‐7,7´/H‐8,8´), 7.29–7.32 (m, 4H, H‐5,5´/H‐6,6´), 7.38 (d, J = 8.0 Hz, 2H, H4‐/H‐4´), 8.20 (d, J = 8.0 Hz, 2H, H‐3/H‐3´), 11.19 ppm (s, 1H, NH‐H2), 12.86 ppm (s, 1H, ‐NH‐H1). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 180.03 ( C =S), 169.36 ( C =O), 157.43, 137.97, 129.85, 129.43, 128.32, 128.19, 127.04, 125.71, 125.40, 124.86, 124.64, 116.77, 116.09, 103.51, 99.79 (aromatic rings), 84.25, 83.00 (p‐cymene), 48.88 (CH₂ carbons of piperazine), 35.28 (‐ C H‐(CH₃)₂), 30.49 (benzene ring‐ C H₂‐CH₃), 22.33 (‐CH‐( C H₃)₂), 21.55 (cymene‐ C H₃), 18.42 (benzene ring‐ CH₂‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M –HCl, Cl^‐]⁺/found: 679.21/679.40 (Figure S39).

4.2.3.4. L₄RuCl₂‐m (4m)

Complex 4m was synthesized following the general procedure for the synthesis of monodentate complexes by using L ₄ (0.917 g, 2 mmol) and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Orange color solid. Yield: 1.222 g, 80%. M.p.: 200 °C. Anal. Calcd. for C₃₇H₄₄Cl₂N₄ORuS (0.5 water) (%): C, 57.43; H, 5.86; N, 7.24. Found: C, 57.36; H, 5.73; N, 7.16. UV–vis (DCM), 5 × 10⁻⁵ M, λ _max nm, ε (dm³·mol⁻¹·cm³): 258 (24516), 328 (7942). FT‐IR (KBr, cm⁻¹): 3220 (m; ν(amide N—H)), 3045 (m; ν(thiourea N—H)), 1676 (s; ν(C=O)), 1336 (s; ν(C—N), 1234 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.24 (d, J = 8.0 Hz, 6H, H‐19/19´), 2.16 (s, 3H, H‐18), 2.23 (s, 3H, H‐17), 2.33 (s, 6H, H‐16/H‐16´), 2.86–2.92 (Sep, J = 8 Hz, 1H, H‐15), 3.39–3.47 (m, 8H, H‐11–H‐14), 5.15 (d, J = 5.8 Hz, 2H, H‐10/H‐10´), 5.31 (d, J = 5.8 Hz, 2H, H‐9/H‐9´), 6.80 (s, 2H, H‐8/H‐8´), 6.92 (t, J = 8 Hz, 2H, H‐8/H‐8´), 7.01–7.05 (m, 4H, H‐6,6´/H‐5,5´), 7.31 (t, J = 8.0 Hz, 2H, H4‐/H‐4´), 7.43 (d, J = 8.0 Hz, 2H, H‐3/H‐3´), 11.17 ppm (s, 1H, NH‐2H), 12.54 ppm (s, 1H, ‐NH‐1H). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 179.87 ( C =S), 174.21 ( C =O), 139.78, 136.14, 134.04, 132.26, 129.49, 129.17, 128.44, 128.36, 127.55, 126.86, 125.43, 124.60, 116.10, 103.20, 100.16 (aromatic rings), 84.06, 82.73 (p‐cymene), 48.84 (CH₂ carbons of piperazine), 30.48 (‐ C H‐(CH₃)₂), 22.38 (benzene ring‐CH‐( C H₃)₃), 21.38 (benzene ring‐ C H₃ (ortho)), 19.53 (benzene ring‐ C H₃ (meta)), 18.51 (cymene‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M –HCl, Cl^‐]⁺/found: 693.22/693.45 (Figure S40).

4.2.3.5. L₁RuCl‐b (1b)

Complex 1b was synthesized following the general procedure for the synthesis of bidentate complexes by using L ₁ (0.833 g, 2 mmol), base (2 mmol), and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Yellow‐orange color solid. Yield: 1.112 g, 81%. M.p.: 217 °C. Anal. Calcd. for C₃₄H₃₇ClN₄ORuS (%): C, 59.51; H, 5.43; N, 8.16. Found: C, 59.68; H, 5.58; N, 8.00. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 252 (36570), 285 (21436), 354 (13422). FT‐IR (KBr, cm⁻¹): 3053 (m; ν(thiourea N—H)), 1603 (s; ν(C=O)), 1315 (s; ν(C—N), 1217 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.16 (d, J = 8.0 Hz, 3H, H‐18), 1.19 (d, J = 8.0 Hz, 3H, H‐18´), 2.13 (s, 3H, H‐17), 2.91–3.02 (Sep, J = 8.0 Hz, 1H, H‐16), 3.31–3.38 (m, 8H, H‐11–H‐15), 4.48 (d, J = 8 Hz, 1H, H‐10), 4.57 (d, J = 8 Hz, 1H, H‐10´), 4.69 (d, J = 8 Hz, 1H, H‐9), 5.26 (d, J = 8 Hz, 1H, H‐9´), 6.90 (t, J = 7.3 Hz, 1H, H‐8), 6.88 (d, J = 8 Hz, 2H, H‐7/H‐7´), 6.97 (d, J = 8 Hz, 2H, H‐6/H‐6´), 7.30 (t, J = 7.3 Hz, 2H, H‐5/H‐5´), 7.39 (d, J = 4.0 Hz, 2H, H‐4/H‐4´), 7.49 (t, J = 4.0 Hz, 2H, H‐3/H‐3´), 7.53 (t, J = 4.0 Hz, 1H, H‐2), 8.22 (d, J = 7.7 Hz, 2H, H‐1/H‐1´), 12.53 ppm (s, 1H, NH). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 188.38 ( C =S), 179.01 ( C =O), 151.27, 149.89, 138.13, 130.94, 129.33, 128.03, 127.44, 124.73, 120.28, 116.51, 116.30, 100.33, 100.09 (aromatic rings), 84.48, 82.59, 80.03, 79.32 (p‐cymene), 49.46, 49.18 ( C H₂ carbons of piperazine), 31.43 (‐ C H‐(CH₃)₂), 23.18, 21.67 (‐CH‐( C H₃)₂), 18.83 (cymene‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M ‐Cl]⁺/found: 651.18/651.23 (Figure S41).

4.2.3.6. L₂RuCl‐b (2b)

Complex 2b was synthesized following the general procedure for the synthesis of bidentate complexes by using L ₂ (0.861 g, 2 mmol), base (2 mmol), and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Yellow‐orange color solid. Yield: 1.120 g, 80%. M.p.: 199 °C. Anal. Calcd. for C₃₅H₃₉ClN₄ORuS (%): C, 60.03; H, 5.61; N, 8.00. Found: C, 59.83; H, 5.58; N, 8.00. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 252 (41040), 285 (25420), 354 (14644). FT‐IR (KBr, cm⁻¹): 3031 (m; ν(thiourea N—H)), 1662 (s; ν(C=O)), 1321 (s; ν(C—N), 1228 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.15 (d, J = 7.0 Hz, 3H, H‐17), 1.19 (d, J = 7.0 Hz, 3H, H‐17´), 2.14 (s, 3H, H‐16), 2.46 (s, 3H, H‐15), 2.63–2.73 (Sep, J = 7.0 Hz, 1H, H‐14), 3.31–3.36 (m, 8H, H‐10–H‐13), 4.57 (dd, J = 7.0 Hz, 2H, H‐9/H‐9´), 4.70 (d, J = 7.0 Hz, 1H, H‐8), 5.27 (d, J = 7.0 Hz, 1H, H‐8´), 6.87 (t, J = 8.0 Hz, 1H, H‐7), 6.93 (t, J = 8 Hz, 2H, H‐6/H‐6´), 6.97 (d, J = 8 2H, H‐5/H‐5´), 7.28–7.31 (m, 4H, H‐3,3´/H‐4,4´), 7.38 (t, J = 8.0 Hz, 2H, H‐2/H‐2´), 8.15 (d, J = 8.0 Hz, 2H, H‐1/H‐1´), 12.53 ppm (s, 1H, NH). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 188.27 ( C =S), 179.02 ( C =O), 151.29, 149.85, 141.39, 135.81, 129.62, 128.62, 127.52, 124.73, 120.28, 116.51, 116.31, 100.28 (aromatic rings), 84.57, 82.69, 80.29, 79.38 (p‐cymene), 49.47, 49.21 (CH₂ carbons of piperazine), 31.42 (‐ C H‐(CH₃)₂), 23.13, 21.75 (‐CH‐( C H₃)₂), 18.85 (cymene‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M ‐Cl]⁺/found: 665.19/665.63 (Figure S42).

4.2.3.7. L₃RuCl‐b (3b)

Complex 3b was synthesized following the general procedure for the synthesis of bidentate complexes by using L ₃ (0.889 g, 2 mmol), base (2 mmol), and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Yellow‐orange color solid. Yield: 1.100 g, 77%. M.p.: 205 °C. Anal. Calcd. for C₃₆H₃₉ClN₄ORuS (0.5 water) (%): C, 59.68; H, 5.85; N, 7.75. Found: C, 59.30; H, 5.20; N, 7.76. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 252 (43586), 285 (25402), 354 (15306). FT‐IR (KBr, cm⁻¹): 3152 (m; ν(thiourea N—H)), 1671 (s; ν(C=O)), 1335 (s; ν(C—N), 1226 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.16 (d, J = 8.0 Hz, 3H, H‐18), 1.19 (d, J = 8.0 Hz, 3H, H‐18´), 1.28 (t, J = 8.0 Hz, 3H, H‐17), 2.14 (s, 3H, H‐16), 2.65 (q, J = 8.0 Hz, 2H, H‐14), 2.63–2.73 (Sep, J = 8.0 Hz, 1H, H‐16), 3.32–3.38 (m, 8H, H‐10–H‐13), 4.51 (d, J = 4.0 Hz, 1H, H‐9), 4.60 (d, J = 4.0 Hz, 1H, H‐9´), 4.70 (d, J = 4.0 Hz, 1H, H‐8), 5.26 (d, J = 4.0 Hz, 1H, H‐8´), 6.88 (t, J = 8.0 Hz, 1H, H‐7), 6.93 (d, J = 8.0 Hz, 2H, H‐6/H‐6´), 6.98 (d, J = 8.0 Hz, 2H, H‐5/H‐5´), 7.28–7.32 (m, 4H, H‐3,3´/H‐4,H‐4´), 7.38 (d, J = 8.0 Hz, 2H, H2‐/H‐2´), 8.15 (d, J = 8.0 Hz, 2H, H‐1/H‐1´), 12.53 ppm (s, 1H, ‐NH). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 188.40 ( C =S), 179.33 ( C =O), 151.46, 150.02, 147.86, 135.66, 129.79, 127.63, 124.89, 120.46, 116.69, 116.49, 100.54, 100.13 (aromatic rings), 84.72, 82.78, 80.25, 79.61 (p‐cymene), 49.65, 49.38 (CH₂ carbons of piperazine), 31.60 (‐ C H‐(CH₃)₂), 29.30 (benzene ring‐ C H₂‐CH₃), 23.38, 21.87 (‐CH‐( C H₃)₂), 19.00 (cymene‐ C H₃), 15.93 (benzene ring‐ CH₂‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M ‐Cl]⁺/found: 679.21/679.63 (Figure S43).

4.2.3.8. L₄RuCl‐b (4b)

Complex 4b was synthesized following the general procedure for the synthesis of bidentate complexes by using L ₄ (0.917 g, 2 mmol), base (2 mmol), and [Ru(η⁶‐p‐cymene)Cl(μ‐Cl)]₂ (0.612 g, 1 mmol). Yellow‐orange color solid. Yield: 1.165 g, 80%. M.p.: 237 °C. Anal. Calcd. for C₃₇H₄₃ClN₄ORuS (%): C, 61.01; H, 5.95; N, 7.69. Found: C, 60.61; H, 5.39; N, 7.74. UV–vis (DCM), 5 × 10⁻⁵ M, λ_max nm, ε (dm³·mol⁻¹·cm³): 252 (36570), 285 (21436), 354 (13422). FT‐IR (KBr, cm⁻¹): 2974 (m; ν(thiourea N—H)), 1629 (s; ν(C=O)), 1324 (s; ν(C—N), 1223 (s; ν(C=S)). ¹ H NMR (400 MHz, CDCl₃, 298 K, ppm): 1.18 (d, J = 8.0 Hz, 3H, H‐17), 2.24 (d, J = 8.0 Hz, 3H, H‐17´), 2.08 (s, 3H, H‐16), 2.38 (s, 3H, H‐16), 2.65 (d, 6H, H‐14/H‐14´), 2.73–2.82 (Sep, J = 8 Hz, 1H, H‐13), 3.34–3.36 (m, 8H, H‐9–H‐12), 3.76 (d, J = 6.0 Hz, 1H, H‐8´), 4.71 (d, J = 6.0 Hz, 1H, H‐8), 4.77 (d, J = 6.0 Hz, 1H, H‐7´), 5.03 (d, J = 6.0 Hz, 1H, H‐7), 6.88 (t, J = 8.0 Hz, 1H, H‐6), 6.94 (d, J = 8.0 Hz, 2H, H‐5,5´), 6.98–7.02 (m, 4H, H‐3,3´/H‐4,4´), 7.28 (t, J = 8.0 Hz, 2H, H2‐/H‐2´), 7.43 (d, J = 8.0 Hz, 2H, H1‐/H‐1´), 12.78 (s, 1H, ‐NH). ¹³ C NMR (150 MHz, CDCl₃, 298 K, ppm): δ, 188.76 ( C =S), 181.18 ( C =O), 151.30, 149.79, 138.45, 136.15, 133.13, 127.99, 127.55, 124.61, 120.27, 116.28, 102.21, 98.85 (aromatic rings), 83.91, 82.40, 79.65, 75.48 (p‐cymene), 49 26 (CH₂ carbons of piperazine), 32.03 (‐ C H‐(CH₃)₂), 23.81 (benzene ring‐CH‐( C H₃)₃), 21.33, 20.25 (benzene ring‐ C H₃ (ortho)), 19.94 (benzene ring‐ C H₃ (meta)), 18.54 (cymene‐ C H₃). ESI ⁺ ‐MS (MeOH): cal [M ‐Cl]⁺/found: 693.22/693.32 (Figure S44).

4.3. Lipophilicity Study

The lipophilicity of complexes 1m–4m and 1b–4b was determined by measuring their partition coefficients (Log P) using the shake‐flask method in a biphasic n‐octanol/water system according to a previously reported procedure [78]. Initially, stock solutions of complexes 1m–4m and 1b–4b were prepared by dissolving 5 mg of each complex in 5 mL of methanol‐saturated water (20:80, v/v). These stock solutions were subsequently diluted with water to obtain final concentrations of 5, 10, 15, 20, and 25 μg/mL. Similarly, separate stock solutions were prepared by dissolving 5 mg of each complex in 5 mL of n‐octanol, followed by dilution to the same concentration range (5–25 μg/mL). For the partitioning experiment, equal volumes (2 mL each) of the aqueous and n‐octanol phases were combined and mechanically shaken for 24 h at room temperature to ensure equilibrium distribution of the complexes between the two immiscible phases. After equilibration, the concentrations of complexes 1m–4m and 1b–4b in both phases were quantified by UV–vis spectroscopy at their respective absorption maxima (λ _max = 249–252 nm). The concentrations were calculated according to the Beer–Lambert law using previously established calibration curves. Finally, the partition coefficient (Log P) values were calculated using the following Equation (1) [79]:

$$\log P=\log \left(\frac{c_{\mathrm{O}}}{c_{\mathrm{W}}}\right)$$ (1)

where C _O and C _W represent the equilibrium concentrations of the complexes in the n‐octanol and aqueous phases, respectively.

4.4. DNA Interaction Studies

Spectroscopic titration is widely employed to evaluate the binding of small molecules to DNA, particularly through UV–vis absorption methods [38]. In this study, UV–vis absorption titrations were conducted to examine the interaction of the synthesized Ru(II)‐p‐cymene complexes (1m to 4b) with CT‐DNA under pH 7.4 conditions. Absorption spectroscopy provides a sensitive approach for probing the binding behavior between metal complexes and biomolecular targets. The spectra obtained during the titration of 2b (10 μM) with CT‐DNA showed a gradual increase in absorbance at approximately 265 nm (0–70 μM), accompanied by a slight blueshift, indicative of noncovalent association between 2b and the DNA helix.

The extent of binding was quantified using the Stern–Volmer equation, [DNA]/(ε _a − ε _f) = [DNA]/(ε _b − ε _f) + 1/K _b(ε _b − ε _f), where [DNA] represents the DNA concentration in base pairs; ε _a is the apparent extinction coefficient, calculated from A(observed)/[complex]; and ε _f and ε _b correspond to the extinction coefficients of the free and fully bound complex, respectively.

4.5. Fluorescence Competitive Displacement Assay

The competitive binding ability of the complexes was investigated through an ethidium bromide (EB) displacement assay. Fluorescence spectra were recorded by maintaining a fixed concentration of DNA (5 μM) and EB (5 μM) while incrementally adding the Ru(II) complexes (0–30 μM). The quenching of the EB‐DNA fluorescence was monitored, and the Stern–Volmer quenching constants (K _sv) were derived from the slopes of the linear plots according to Equation (2):

$$\frac{I_0}{I}=1+\hspace{0.17em}{K}_{\mathrm{sv}}\left[Q\right]$$ (2)

where I ₀ and I are the fluorescence intensities in the absence and presence of the complex (quencher), respectively. The apparent binding constants (K _app) were further calculated from the K _SV values at an EB concentration of 5 μM to evaluate the consistency of the binding results.

4.6. MG Competitive Displacement Assay

To evaluate the specific major‐groove binding affinity of the ruthenium(II) complexes, a fluorescence competitive displacement assay was performed using MG as a major groove‐specific probe. The MG–dsDNA complex was prepared by incubating Ct‐DNA (5.0 μM) with MG (5.0 μM) in Tris–HCl buffer (5 mM Tris–HCl, 50 mM NaCl, pH 7.4) for 30 min at room temperature in the dark. The fluorescence titration was carried out by sequentially adding incremental amounts of the representative complex 2b (with structural ratios of complex to dsDNA at 1:1 and 1:2) into the MG–dsDNA solution. After each addition, the mixture was allowed to equilibrate for 5 min. The fluorescence emission spectra were recorded in the wavelength range of 620–750 nm upon excitation at 600 nm.

4.7. Anticancer Studies

4.7.1. Cell Culture

Human melanoma A375 cells and A2058 cells were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA). The human normal retinal pigment epithelial ARPE‐19 cells were kindly provided by Dr. Chien‐Chih Chiu from the Department of Biotechnology, Kaohsiung Medical University (Kaohsiung, Taiwan). The A375 and A2058 cells were maintained in DMEM supplemented with 10% FBS (fetal bovine serum) and antibiotics (100 units/mL penicillin and 100 μg/mL streptomycin) at 37 °C in a humidified atmosphere of 5% CO₂. In addition, the completed cultured medium for ARPE‐19 cells was DMEM/F12/FBS (10%)/antibiotics (1%) and DMEM/FBS (10%)/antibiotics (1%), respectively.

4.7.2. Cytotoxicity Assay

Cytotoxicity of monodentate (1m–4m) and bidentate species (1b–4b) was studied by using the MTT [3‐(4,5‐dimethylthiazol‐2‐yl)‐5‐(3‐carboxymethoxyphenyl)‐2‐(4‐sulfophenyl)‐2H‐tetrazolium] assay. All the cell lines were cultured and maintained in the complete culture medium. The 5 × 10 ³ cells/well were seeded in a 96‐well plate and incubated overnight in a humidified condition at 37 °C with 5% CO₂. Each complex was dissolved in DMSO to prepare a 1 mM stock solution, which was subsequently diluted with culture medium to obtain the required working concentrations (0–50 µM). The cells were treated with the respective concentrations for 72 h. After treatment with compounds, the cells were washed twice with PBS and incubated with diluted fresh MTT media for 4 h. The data were taken at 490 nm. By using the formula A−B /A × 100 (A = control group and B = treated group), the cell viability (%) was calculated. Each experimental point represents the mean ± SD of three replicates.

4.7.3. Migration Assay

To evaluate cell migration capacity, a wound healing assay was performed using ibidi Culture‐Inserts 2 Well (ibidi GmbH, Martinsried, Germany). Cells were seeded into each well of the insert at a density of 3 × 10⁴ cells per well and incubated at 37 °C in a 5% CO₂ atmosphere to allow for the formation of a confluent monolayer. Following cell attachment, the inserts were carefully removed using sterile forceps to create a cell‐free gap of a defined width. The wells were gently washed with phosphate‐buffered saline (PBS) to remove nonadherent cells, and the medium was replaced with fresh culture medium. Images of the wound area were captured at 0, 12, and 24 h postremoval using an inverted phase‐contrast microscope. The extent of cell migration was quantified by measuring the wound area at each time point using ImageJ software (National Institutes of Health, Bethesda, MD, USA). The wound healing rate (percentage of wound closure) was calculated using the following formula (3):

$$\text{Migration}\hspace{0.17em}\mathrm{rate}\hspace{0.17em}\left(\mathrm{\%}\right)=\hspace{0.17em}\frac{A_0-A_t}{A_0}\hspace{0.17em}\times 100$$ (3)

4.7.4. Western Blotting

To evaluate the expression levels of cell cycle‐ and migration‐regulated proteins, human melanoma A375 cells were treated with various concentrations of complex 2b (0, 2.5, 5, and 10 μM) for 72 h. Following treatment, both control and treated cells were harvested, washed twice with ice‐cold PBS, and lysed in RIPA lysis buffer supplemented with a protease and phosphatase inhibitor cocktail for 30 min on ice. The lysates were centrifuged at 12,000 × g for 15 min at 4 °C to remove cell debris, and the supernatant containing the total protein was collected. Protein concentrations were quantified using a BCA protein assay kit according to the manufacturer's instructions. Equal amounts of protein samples (30 μg) were mixed with 5× SDS sample loading buffer, boiled at 95 °C for 5 min, and subsequently separated by 10% or 12% SDS‐PAGE gel electrophoresis. The resolved proteins were electrotransferred onto polyvinylidene difluoride (PVDF) membranes. To prevent nonspecific binding, the membranes were blocked with 5% nonfat milk in Tris‐buffered saline containing 0.1% Tween‐20 (TBST) for 1 h at room temperature. The blocked membranes were then incubated overnight at 4 °C with specific primary antibodies against p21 (1:1000, Cell Signaling Cat# 2947T), Cyclin B1 (1:1000, Cell Signaling Cat# 12231T), MMP‐9 (1:2000, ABclonal Cat# A0289), α‐tubulin (1:10000, Abclonal Cat# AC012), and GAPDH (1:20000, ABclonal Cat# A19056). After washing three times with TBST buffer, the membranes were incubated with the corresponding horseradish peroxidase (HRP)‐conjugated secondary antibodies for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence (ECL) detection system, and the immunoreactive band intensities were quantified via densitometric analysis using ImageJ software. α‐Tubulin and GAPDH were employed as internal loading controls to normalize protein expression levels.

4.7.5. In Silico Studies

Using the DMOL³ program in the Materials Studio package, the lowest energies, HOMO/LUMO, and the chemical reactivity descriptors were calculated. All complexes were optimized by DFT‐D3 based on the DNP basis set (basis file 3.5) and exchange‐correlation function, which was chosen from GGA and PBE. Geometry optimizations were carried out using the DNP basis set (basis file 3.5) with spin‐unrestricted treatment (up to 1000 iterations; smearing 0.005 Ha; 4 Å cutoff). Grimme's semiempirical dispersion correction was included to improve the description of noncovalent interactions. All compounds were saved as Mol2 for molecular docking studies [80].

4.7.6. Molecular Docking

Molecular docking was performed to investigate the interaction of the synthesized Ru(II)–p‐cymene complexes with duplex B‐DNA. The crystal structure of B‐DNA [d(CGCGAATTCGCG)₂] (PDB ID: 1AIO) was obtained from the RCSB Protein Data Bank and prepared by removing water molecules, adding hydrogens, and optimizing with the CHARMm force field. Docking simulations were conducted using the LibDock module of Discovery Studio 2022 (BIOVIA, Dassault Systems, San Diego, USA), which aligns ligands to the active groove region based on polar and apolar interaction hotspots. The docking pose of 2b complexed with dsDNA was shown by PyMOL, and the interaction was analyzed using LigPlot⁺ to identify hydrogen bonding, π–π stacking, and hydrophobic contacts with DNA residues [38].

4.7.7. Statistical Analysis

Statistical analysis was performed using Student's t‐test for cell assays. * represents p < 0.05 and ** represents p < 0.01; both were considered significant.

Author Contributions

Amir Karim: conceptualization, data curation, formal analysis, investigation, methodology, writing – original draft, writing – review and editing. Xin Wang: data curation, formal analysis, methodology. Yu‐Ting Chu: data curation, formal analysis, software. Rahime Eshaghi Malekshah: formal analysis, methodology, computational study, writing – original draft. En‐De Shu: data curation, formal analysis, methodology. Najeeb Ullah: formal analysis, writing – original draft. Chien‐Chih Chiu: supervision, data curation, writing – review and editing. Ching‐Ming Chien: supervision, funding acquisition, data curation, writing – review and editing. Sodio C. N. Hsu: supervision, funding acquisition, project administration, resources, data curation, writing – review and editing.

Funding

Amir Karim, Rahimeh Eshaghi Malekshah, Najeeb Ullah, Yu‐Ting Chu, and Sodio Hsu were supported by the National Science and Technology Council, Taiwan (NSTC 113‐2113‐M‐037‐006, NSTC 113‐2113‐M‐037‐017, and NSTC 114‐2113‐M‐415‐002‐MY2), the Kaohsiung Medical University (KMU‐TB115002), and the NSYSU‐KMU Joint Research Project (NSYSUKMU 114‐P20).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Crystallographic information files (.cif) for the complexes have been deposited in the CCDC: 2496494 (1m), 2496503 (2b), 2496499 (4b). Additional supporting information and experimental data in support of this article are presented in the Supporting Information.

Data Availability Statement

Crystallographic information files (.cif) for the complexes have been deposited in the CCDC: 2496494 (1m), 2496503 (2b), 2496499 (4b). Other experimental data in support of this article are presented in the Supporting Information.