Ru(II)-diphosphine/N,S-mercapto complexes and their anti-melanoma properties†

Nádija N. P. da Silva ^a, Marcos V. Palmeira-Mello ^a, Nathália O. Acésio ^b, Carlos A. F. Moraes ^a, João Honorato ^c, Eduardo E. Castellano ^c, Denise C. Tavares ^b, Katia M. Oliveira *^d and Alzir A. Batista *^a
aDepartament of Chemistry, Federal University of São Carlos – UFSCar, CEP 13565-905, São Carlos, SP, Brazil. E-mail: daab@ufscar.br
^bUniversity of Franca – UNIFRAN, CEP 14404-600, Franca, SP, Brazil
^cPhysics Institute of São Carlos, University of São Paulo – USP, CEP 13560-970, São Carlos, SP, Brazil
^dInstitute of Chemistry, University of Brasília – UnB, CEP 70910-900, Brasília, DF, Brazil. E-mail: katia.oliveira@unb.br

First published on 14th November 2024

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Introduction

Cancer, one of the most serious public health problems and a leading cause of death worldwide, is marked by cells that proliferate rapidly and uncontrollably and which infiltrate various tissues and organs. Among the diverse types of cancer, melanoma stands out: it increases at a higher rate than other forms of cancer.¹ Melanoma emerges from mutations in genes that regulate the growth and viability of melanocytes, which are cells that produce melanin and thus offer protection against harmful ultraviolet radiation (UV) and contribute to skin pigmentation. Cutaneous melanoma is the most prevalent form, but melanoma also manifests in tissues like mucous membranes, uveal tract, and leptomeninges.^2,3

In its early stages, melanoma can be effectively treated with surgery, and the survival rate is high. However, diagnosing melanoma late, as in the case of advanced or metastatic melanoma, restricts treatment options.² Therefore, developing methodologies to treat melanoma more effectively while producing fewer or no side effects is crucial, especially when it comes to the advanced stages of the disease.⁴

In clinical practice, chemotherapy based on metal complexes has stood out since cisplatin (cis-[PtCl₂(NH₃)₂], cis-diamminedichloroplatinum(II)) was discovered and approved by the FDA (Food and Drug Administration).⁵ This drug displays broad cytotoxic action and has proven effective for treating lung, head, ovarian, testicular, and esophageal cancer, for instance. Unfortunately, many issues related to side effects and acquired resistance have been reported after continuous use of this chemotherapy.^6–8

To mitigate the undesired effects of cisplatin, researchers have developed new platinum-based compounds with different structures, where mono- or bidentate ligands replace amine ligands to modulate electronic, steric, and basicity effects. Examples of these compounds include carboplatin, oxaliplatin, and nedaplatin, among others.⁹ Although some side effects have been reduced, others have persisted during chemotherapeutic treatment. Consequently, the scientific community has been exploring alternative metal centers in an endeavor to develop complexes that target tumor cells more effectively and selectively.^10,11

Ruthenium has been extensively researched in this context, and ruthenium complexes have been shown to exhibit promising cytotoxic and antitumor activities.¹² Among the ruthenium complexes with potential chemotherapeutic action,^13–15 the BOLD-100 or KP-1339 (trans-[tetrachlorobis(1H-indazole)ruthenate(III)]) complex, developed by Keppler and colleagues, is noteworthy. This compound induces DNA damage, cell cycle arrest, and apoptosis and is now undergoing clinical trials.¹⁰ In this regard, our research group has developed novel ruthenium complexes with different phosphine co-ligands to enhance the cytotoxic activity of the final complexes by taking advantage of the synergism that might occur between the metal center and the ligands in the coordination sphere.^16–18

N,S-Mercapto comprises a group of molecules that can act as ligands and which have attracted our attention. When these ligands coordinate to ruthenium, notably cytotoxic complexes arise, particularly complexes featuring phosphine as co-ligands.^18–20

In this study, we aimed to synthesize novel ruthenium complexes containing 2-mercapto-4(3H)-quinazoline as ligand and different phosphines as co-ligands. Our goal was to establish a possible structure–activity relationship and to explore how changing the substituents in the phosphine groups affects the cytotoxicity of the ruthenium complexes toward melanoma cell lines. Additionally, we have investigated how the ruthenium complexes interact with biomolecules such as CT-DNA and human serum albumin (HSA).

Experimental section

General methods and materials

All the syntheses were conducted under argon atmosphere. RuCl₃·nH₂O, H2mq (2-mercapto-4(3H)-quinazoline), dppe [1,2′-bis(diphenylphosphine)ethane], dppm [1,1′-bis(diphenylphosphine)methane], dppen [1,2′-bis(diphenylphosphine)ethene], bipy (2,2′-bipyridine), calf-thymus DNA (CT-DNA), human serum albumin (HSA), and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were purchased from either Merck or Sigma Aldrich and were used as received. The cis-[RuCl₂(dppm)₂], cis-[RuCl₂(dppe)₂], cis-[RuCl₂(dppen)₂], and cis-[RuCl₂(bipy)₂] precursor complexes were synthesized by using a well-established procedure.^21–24

The IR spectra of the ruthenium complexes were acquired from 4000 to 400 cm⁻¹ on a Bomem–Michelson 102 Fourier transform infrared spectrometer; KBr pellets were used. Molar conductivity was measured on a Meter Lab CDM2300 instrument; 1 × 10^–3 M DMSO or dichloromethane solutions of the ruthenium complexes were employed. Elemental analyses were carried out on a Fisions CHNS analyzer model EA 1108 at the Central Analytical Laboratory of the Department of Chemistry at the Federal University of São Carlos, São Carlos, São Paulo, Brazil. UV-visible absorption spectra were recorded from 250 to 700 nm on a Hewlett-Packard 8452A diode array spectrophotometer; DMSO or dichloromethane solutions of the ruthenium complexes were placed in quartz cuvettes of 1 cm optical path length. Cyclic voltammetry assays were performed on an EGeG Princeton Applied Research Model 273A Potentiostat/Galvanostat by using an electrochemical cell with a three-electrode system, namely Ag/AgCl as the reference electrode and two platinum plates as the auxiliary and working electrodes, immersed in 0.1 mol L⁻¹ tetrabutylammonium perchlorate (TBAP, Fluka Chemica) solution in dichloromethane. ³¹P{¹H}, ¹H, COSY (¹H–¹H), ¹³C{¹H}, HSQC (¹H–¹³C), and HMBC (¹H–¹³C) NMR spectra were recorded on a 9.4 T Bruker Avance III 400 MHz spectrometer.

[RuCl(H2mq)(dppm)₂]PF₆ (Ru1)

To obtain Ru1, 0.018 g (0.10 mmol) of H2mq was added to a Schlenk flask containing a deaerated methanol/dichloromethane mixture (5:1). The system was stirred and refluxed until the ligand was completely dissolved. Then, 0.10 g (0.10 mmol) of cis-[RuCl₂(dppm)₂] and 0.019 g (0.10 mmol) of KPF₆ were added. The system was kept under identical conditions for an additional hour, and then the volume was reduced to around 2 mL. Ethyl ether was introduced to induce precipitation of a yellow powder, which was filtered, rinsed with distilled water and ethyl ether, and dried under vacuum. Yield: (0.080 mmol; 0.098 g) 76%. Anal. calcd for C₅₈H₅₀ClF₆N₂OP₅SRu: C, 56.71; H, 4.10; N, 2.28; S, 2.61. Found: C, 56.86; H, 3.96; N, 2.44; S, 3.03. ESI-MS calcd for [Ru(2mq)(dppm)₂]⁺ ([M]⁺): m/z 1047.1560. Found: m/z 1047.1554. Selected IR (KBr, v, cm⁻¹) 3056 (vC_sp²–H); 1625 (vCN); 1560 (vCC + vCN); 1261 (vC–S); 1906 (vP–C); 836 (vPF₆⁻); 726 (δC–H (ring)); 560 (δPF₆⁻); 482 (vRu–S); 414 (vRu–N). ³¹P{¹H} NMR (162 MHz, D₂O) δ −1.73 (1P, dt, J = 41.7, 28.5 Hz); −3.82 (1P, dt, J = 37.9, 27.7 Hz); −22.17 (1P, ddd, J = 324.0, 41.8, 27.7 Hz); −23.68 to −26.55 (1P, m); −143.04 (1P, hept, PF₆⁻). ¹H NMR (400 MHz, acetone-d₆): δ 13.54 (1H, s, H2mq); 12.23 (1H, s, H2mq) 8.34–6.80 (44H, m, aromatics of dppm and H2mq); 5.7–5.0 (4H, m, dppm). ¹³C NMR (101 MHz, acetone-d₆, 298 K, ppm) δ 173.64 (1C, s, H2mq (CS)); 157.61 (1C, s, H2mq (CO)); 138.95–125.92 (52C, m, aromatics of dppm and H2mq); 117.40 (1C, s, dppm); 116.84 (1C, s, dppm).

[Ru(2mq)(dppm)₂]PF₆ (Ru2)

In a Schlenk flask containing 20 mL of deaerated methanol, 0.022 g (0.12 mmol) of H2mq and 0.011 g (0.13 mmol) of NaHCO₃ were added. The system was stirred and refluxed for 1 h, and then 0.10 g (0.10 mmol) of cis-[RuCl₂(dppm)₂] and 0.023 g (0.12 mmol) of KPF₆ were added. The system was stirred and refluxed for other 24 h. After that, the volume was reduced to approximately 2 mL, and distilled water was added to induce precipitation of a yellow solid, which was filtered, washed with distilled water and ethyl ether, and dried under vacuum. Yield: (0.083 mmol; 0.100 g) 79%. Anal. calcd for C₅₈H₄₉F₆N₂OP₅SRu: C, 58.44; H, 4.14; N, 2.35; S, 2.69. Found: C, 58.23; H, 4.23; N, 2.32; S, 2.46. ESI-MS calcd for [Ru(2mq)(dppm)₂]⁺ ([M]⁺): m/z 1047.1554. Found: m/z 1047.1582. Selected IR (KBr, v, cm⁻¹): 3061 (vC_sp²–H); 1613 (vCN); 1436 (vCC + vCN); 1244 (vC–S); 1906 (vP–C); 839 (vPF₆⁻); 732 (δC–H (ring)); 561 (δPF₆⁻); 484 (vRu–S); 418 (vRu–N). ³¹P{¹H} NMR (162 MHz, D₂O): δ 0.92 to −0.51 (1P, m); −8.07 to −9.23 (1P, m); −11.68 (1P, ddd, J = 312.9, 42.2, 26.8 Hz); −22.33 (1P, ddd, J = 67.3, 40.2, 27.8 Hz); −143.64 (1P, hept, PF₆⁻). ¹H NMR (400 MHz, DMSO-d₆): δ 8.19–6.56 (54H, m, aromatics of dppm and H2mq); 5.60–4.60 (4H, m, H2mq). ¹³C NMR (101 MHz, DMSO-d₆, 298 K, ppm) δ 178.12 (1C, s, H2mq (C–S)); 165.86 (1C, s, H2mq (CO)); 159.76 (1C, s, H2mq); 148.48 (1C, s, H2mq); 138.20–116.76 (52C, m, aromatics of dppm and H2mq).

[Ru(2mq)(dppe)₂]PF₆ (Ru3)

Ru3 was synthesized according to the procedure reported in the literature.¹⁹ Yield: (0.077 mmol; 0.094 g) 75%. Anal. calcd for C₆₀H₅₃F₆N₂OP₅SRu: C, 59.07; H, 4.38; N, 2.30; S, 2.63. Found: C, 58.98; H, 4.46; N, 2.49; S, 2.82.

[Ru(2mq)(dppen)₂]PF₆ (Ru4)

To obtain Ru4, 0.014 g (0.14 mmol) of H2mq and 32 μL of triethylamine were added to a Schlenk flask containing 30 mL of a deaerated methanol/dichloromethane mixture (1:1). After 10 min, 0.10 g (0.10 mmol) of cis-[RuCl₂(dppen)₂] and 0.028 g (0.15 mmol) of KPF₆ were added. The system was stirred and refluxed for 12 h, and then the volume was reduced to approximately 2 mL. Distilled water was incorporated to induce precipitation of a yellow powder, which was filtered, washed with distilled water and ethyl ether, and dried under vacuum. Yield: (0.078 mmol; 0.095 g) 76%. Anal. calcd for C₆₀H₄₉F₆N₂OP₅SRu: C, 59.26; H, 4.06; N, 2.30; S, 2.64. Found: C, 59.38; H, 4.15; N, 2.72; S, 2.69. ESI-MS calcd for [Ru(2mq)(dppen)₂]⁺ ([M]⁺): m/z 1071.1554. Found: m/z 1071.158. Selected IR (KBr, v, cm⁻¹): 3056 (vC_sp²–H); 1625 (vCN); 1560 (vCC + vCN); 1261 (vC–S); 1906 (vP–C); 836 (vPF₆⁻); 726 (δC–H (ring)); 560 (δPF₆⁻); 482 (vRu–S); 414 (vRu–N). ³¹P{¹H} NMR (162 MHz, D₂O) δ 69.08–66.87 (1P, m); 59.57 (1P, m); 56.73 (1P, m); 53.16–51.73 (1P, m); −143.64 (1P, hept, PF₆⁻). ¹H NMR (400 MHz, DMSO-d₆): δ 8.95–6.54 (44H, m, aromatics of dppen and H2mq); 6.22–5.71 (4H, m, dppen). ¹³C NMR (101 MHz, DMSO-d₆, 298 K, ppm) δ 171.51 (1C, s, H2mq (C–S)); 159.02 (1C, s, H2mq); 144.88 (1C, s, H2mq (CO)); 138.40 (1C, s, H2mq); 134.60–127.11 (52C, m, aromatics of dppen and H2mq); 122.88 (4C, s, dppen).

[Ru(2mq)(bipy)₂]PF₆ (Ru5)

To synthesize Ru5, 0.044 g (0.24 mmol) of H2mq was added to a Schlenk flask containing 32 mL of an ethanol/water mixture (1:1). The system was stirred and refluxed until the ligand was completely dissolved. Subsequently, 0.10 g (0.20 mmol) of cis-[RuCl₂(bipy)₂] and 0.114 g (0.61 mmol) of KPF₆ were added. The system was maintained under the same conditions until precipitation of a red solid, which was separated by filtration, washed with 3 mL of distilled water and 3 mL of ethyl ether, and dried under vacuum. Yield: (0.184 mmol; 0.135 g) 89%. Anal. calcd for C₂₈H₂₁F₆N₆OPSRu: C, 45.72; H, 2.88; N, 11.42; S, 4.36. Found: C, 45.83; H, 3.03; N, 11.34; S, 4.52. ESI-MS calcd for [Ru(2mq)(bipy)₂]⁺ ([M]⁺): m/z 591.0536. Found: m/z 591.0553. Selected IR (KBr, v, cm⁻¹): 3072 (vC_sp²–H); 1603 (vCN); 1523 (vCC + vCN); 1245 (vC–S); 844 (vPF₆⁻); 729 (δC–H (ring)); 557 (δPF₆⁻); 479 (vRu–S); 423 (vRu–N). ¹H NMR (400 MHz, DMSO-d₆): δ 12.76 (1NH, s, H2mq); 9.73–7.16 (19H, m, aromatics of bipy and H2mq); 5.34 (1H, d, H2mq). ¹³C NMR (101 MHz, DMSO-d₆, 298 K, ppm) δ 174.74 (1C, s, H2mq (C–S)); 160.63 (1C, s, H2mq); (1C, s, H2mq (CO)); 159.22–118.16 (26C, m, aromatics of bipy and H2mq).

X-ray crystallography

Ru1, Ru2, Ru3, and Ru5 monocrystals were obtained by slow evaporation in acetone, dichloromethane/methanol, acetone/methanol, and methanol, respectively. The X-ray single-crystal diffraction measurements were carried out at the Institute of Physics, University of São Paulo in São Carlos, São Paulo, Brazil. Data were collected on a Rigaku XtaLAB Mini (ROW) or XtaLAB Synergy Dualflex HyPix diffractometer, with MoKα (λ = 0.71073). The crystal structures were determined through direct methods by using the SHELXL²⁵ program and refined by the least-squares method with the aid of the SHELXL program. The crystallographic data and structures of Ru1, Ru2, Ru3, and Ru5 were generated with the OLEX²⁶ program.

Studies on the interaction with DNA

Interaction with human serum albumin (HSA)

Interactions between Ru2–Ru5 and HSA (from Sigma Aldrich) were studied by measuring HSA fluorescence suppression in the presence of different concentrations of a ruthenium complex. To investigate the type of interaction, the experiment was conducted at 298, 303, or 310 K. The HSA solution was prepared by solubilization in Tris-HCl buffer (pH 7.4), and the concentration was determined by UV-vis spectroscopy on the basis of the molar absorptivity at 280 nm (ε = 36500 cm⁻¹ mol⁻¹ L).

In 1000 μL microtubes, increasing aliquots (5–40 μL) of one of the ruthenium complexes in DMSO were added, and the volume was adjusted to 50 μL (5%) with DMSO. To the microtubes, 950 μL of the stock HSA solution was added to reach a final volume of 1000 μL. HSA and DMSO solutions in the absence of a ruthenium complex were used as controls. Aliquots of 200 μL were taken from each microtube and transferred to a 96-well opaque plate; measurements were performed in triplicate. Fluorescence emission was measured from 260 to 500 nm under excitation at 270 nm on a Synergy/H1-Biotek fluorometer equipped with a monochromator.

The fluorescence quenching process was quantitatively analyzed by using the Stern–Volmer equation:²⁷

F0/F = 1 + kqτ0[Q] = 1 + KSV[Q]	(1)

where F₀ and F represent the fluorescence intensity in the absence and presence of the quencher (Ru2–Ru5), respectively; [Q] denotes the quencher concentration; K_SV corresponds to the Stern–Volmer quenching constant; k_q stands for the biomolecular quenching constant; and τ₀ represents the average lifetime of HSA without the quencher (∼10^–8 s).²⁷

To determine the binding constant (K_b) and the number of binding sites (n), eqn (2) was employed:

log[(F0 − F)/F] = logKb + nlog[Q]	(2)

The thermodynamic parameters (ΔH°, ΔS°, and ΔG°) were derived from eqn (3) and (4):²⁸

ln(Kb1/Kb2) = (1/T1 − 1/T2) × ΔH/R	(3)

where K₁ and K₂ are the binding constants at temperatures T₁ and T₂, respectively; and R is the constant of the ideal gases.

ΔG° = −RTlnKb = ΔH° − TΔS°	(4)

General cell culture

The cell lines employed in this study included the human melanoma (A-375) and murine melanoma (B16-F10) cell lines and a non-tumoral human keratinocyte cell line (HaCat). A-375 and B16-F10 were provided by S. S. Maria-Engler, University of São Paulo, São Paulo, Brazil and HaCat cells was provided by P. F. Oliveira, Federal University of Alfenas, Alfenas, Minas Gerais, Brazil. All the cell lines were cultivated in Dulbecco's Modified Eagle Medium Nutrient (DMEM – Sigma Aldrich) supplemented with 10% fetal bovine serum, antibiotics (0.001 mg mL⁻¹ streptomycin and 0.005 mg mL⁻¹ penicillin – Sigma Aldrich), and 2.38 mg mL⁻¹ Hepes (Sigma Aldrich). The cells were cultured as monolayers in 25-cm² disposable flasks at 37 °C under 5% CO₂.

Results and discussion

Synthesis and characterization of ruthenium/diphosphine/N,S-mercapto complexes

The reaction of cis-[RuCl₂(dppm)₂], cis-[RuCl₂(dppe)₂], cis-[RuCl₂(dppen)₂], or cis-[RuCl₂(bpy)₂] with H2mq (2-mercapto-4(3H)-quinazoline) resulted in four novel ruthenium complexes (Ru1, Ru2, Ru4, and Ru5), which we properly isolated and characterized (Fig. 1). Ru3 had previously been synthesized and documented.¹⁹ In this study, we synthesized it specifically to facilitate comparisons between the obtained structures and biological results.

We obtained Ru1–Ru5 as hexafluorophosphate salts of the 1:1 electrolyte type (Fig. 2), confirmed by molar conductance measurements performed in DMSO (55.7–60.50 μS cm⁻¹) and dichloromethane (40 μS cm⁻¹, for Ru1). The elemental analysis data also agreed with the structures proposed for Ru1–Ru5.

The reaction between cis-[RuCl₂(dppm)₂] and H2mq in a dichloromethane/methanol mixture (5:1) gave Ru1, which bears H2mq coordinated to ruthenium through the sulfur atom in a monodentate manner. However, when we introduced sodium bicarbonate into the reaction medium, another coordination mode characterized by chelation and negative charge emerged for the ligand, generating Ru2. As in the case of Ru2, in Ru3–Ru5, H2mq coordinated to ruthenium in a bidentate and anionic fashion, through the S and N atoms.

The IR spectra of Ru1–Ru5 exhibit the same behavior (Fig. S1†). They display bands between 3050 and 3080 cm⁻¹, attributed to the stretching of the C–H bond of aromatic rings. The bands between 1630 and 1430 cm⁻¹ corresponded to stretching of the v(CC) and v(CN) bonds of the aromatic rings of the phosphine, bipyridine, and mercapto ligands. Characteristic vibrations assigned to the v(P–F) and δ(P–F) bonds of PF₆⁻ appeared between 850 and 550 cm⁻¹.³⁰

The electrochemical behavior of Ru1–Ru5 by cyclic voltammetry was investigated. Ru1–Ru5 behaved similarly, with the Ru^II/Ru^III oxidation potential ranging from 0.95 to 1.53 V and the Ru^III/Ru^II reduction potential ranging from 0.48 to 1.06 V (Fig. S2†). Ru1–Ru5 had higher half-wave potential (E_1/2) than the respective precursor complexes (Table 1). As expected, when the chloride ligands, which are σ–π donors, with a σ donor and π acceptor ligand, was replaced, the electron density on the metal center decreased due to electron density back-donation from the metal to the ligand. This electrochemically stabilized ruthenium because its oxidation required a higher potential. Ru1–Ru5 behaved like other phosphine complexes bearing mercapto ligands.^19,31

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The electronic spectra of Ru1–Ru5 showed an intense absorption in the region of 260 nm for Ru1–Ru4, the diphosphine complexes, and 240 nm for Ru5, the bipyridine complex. This absorption referred to intraligand transitions (π–π*) of the aromatic rings of the phosphine, bipyridine, and mercapto ligands. The absorptions from 250 to 340 nm in the spectra of Ru1–Ru4 and from 350 to 520 nm in the spectrum of Ru5 corresponded to metal-to-ligand charge transfer (MLCT) of the dπ_Ru → 3pσ*dπ(phosphine) and dπ_Ru → π*(bipy, mercapto) type, respectively.

We deconvoluted the electronic spectra of Ru1–Ru5. We confirmed the MLCT bands due to dπ_Ru → 3pσ*dπ(phosphine) and dπ_Ru → π*(bipy, mercapto), which were overlapped and resulted in the single MLCT band observed in the electronic spectra. Deconvolution also revealed the d–d transition band, which has a low extinction coefficient and was hidden by the MLCT band (Fig. S3 and S4†).

We characterized Ru1–Ru5 by ¹D and ²D NMR spectroscopy at different nuclei, including ³¹P{¹H}, ¹³C{¹H}, and ¹H. The ³¹P{¹H} NMR spectra of Ru1–Ru5 exhibited four signals with a double double–double (ddd) pattern, which indicated ABMX spin systems. The phosphorus atom trans to nitrogen (N), the most electronegative element among the atoms coordinated to ruthenium, was more deshielded and hence had a greater chemical shift, followed by the phosphorus atom trans to sulfur (S). The chemical shifts referring to phosphorus atoms trans to phosphorus appeared at lower chemical shifts, and the signals corresponding to these atoms were split into two sets of four lines (Fig. S5–S8†) and had higher coupling constant (Table S1†). The ¹³C{¹H} and ¹H NMR spectra and contour maps are shown in Fig. S9–S26.†

The X-ray crystal structures of Ru1, Ru2, Ru3, and Ru5 were determined, and Fig. 3 illustrates their ORTEP diagrams. Ru1, Ru2, Ru3, and Ru5 were hexacoordinated with distorted octahedral geometry: the Ru–S (2.42–2.46 Å) and Ru–P (2.30–2.40 Å) bonds were longer due to the larger atomic radii of phosphorus and sulfur. In comparison, the Ru–N bond (2.13 Å) was shorter. The P–Ru–P and N–Ru–N angles of phosphine and bipyridine, respectively, were larger than the S–Ru–N angles of mercapto ligand (Table 2), as expected.

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Interactions with biomolecules

DNA is a widely exploited target when it comes to developing candidate antitumoral metallodrugs. DNA damage can induce cell death, as seen with cisplatin. Metallodrugs can interact with nucleic acids through non-covalent modes, such as intercalation, insertion, and groove binding, or via a covalent mode, including direct binding of the complex to the DNA structure.³³

In laboratories, several molecular methods are currently used to explore and to classify metallodrug–DNA interactions. Here, we assessed the potential interactions of complexes with CT-DNA by viscosity measurements, competitive assay by fluorescence, circular dichroism, and electrophoresis-based techniques. Variations in DNA viscosity in the presence of a complex provide important information about how a complex interacts with DNA. An intercalating agent, such as thiazole orange, increases the distance between the nitrogenous base pairs, to accommodate the agent, causing the double helix to elongate and DNA viscosity to rise. Covalent interaction elicits the opposite effect—the DNA viscosity decreases because the axial length of the double helix is shortened.^34,35

Before this study, we investigated whether Ru1–Ru5 were stable in the solution. First, we recorded the ³¹P{¹H} NMR spectra of Ru1, Ru2, and Ru4 and the ¹H NMR spectrum of Ru5, dissolved in DMSO, at 0, 24, and 48 h. Based on Fig. S29,† only the spectrum of Ru1, which contains H2mq as a ligand coordinated in a monodentate manner, changed, and new signals appeared. This indicated that DMSO, a coordinating solvent, replaced H2mq and chloride in the coordination sphere of ruthenium. The spectra of Ru2–Ru5 remained unaltered during the time of the experiment (Fig. S29–36†).

Similarly, we investigated the stability of Ru1–Ru5 in a DMSO/culture medium mixture. Except for Ru1, all ruthenium complexes remained stable during the evaluated period (Fig. S31–34†). Given that Ru1 was not stable under the conditions of the biological assays, we did not evaluate its ability to interact with biomolecules or cytotoxic activity.

To characterize how Ru2–Ru5 interact with DNA, we measured viscosity at constant CT-DNA concentration and varying concentrations of the ruthenium complex, to obtain different [ruthenium complex]/[CT-DNA] molar ratios. Unlike thiazole (an intercalating agent) and cisplatin (which covalently interacts with DNA), adding aliquots of a ruthenium complex to the CT-DNA solution did not modify DNA viscosity significantly (Fig. 4A), indicating that Ru2–Ru5 established electrostatic or groove interactions. The confidence limits of the viscosity tests can be found in Table S4.†

We also used circular dichroism (CD) to characterize how Ru2–Ru5 interact with DNA. In the CD spectrum of CT-DNA (Fig. 4A), two bands emerged: the band at 245 nm in the negative region, due to DNA helicity (right-handed twist), and the band at 275 nm in the positive region, due to base stacking. These bands are highly sensitive to interaction between small molecules and DNA. While the CD spectrum of CT-DNA remains unchanged during electrostatic and minor groove binding, intercalation significantly alters both the positive and negative bands.^36–38

Fig. 4B shows the CD spectrum of CT-DNA in the presence of Ru2. There were no notable alterations in the intensity or position of the DNA bands, which suggested that Ru2 and DNA interacted weakly, possibly via electrostatic or groove interactions. Ru3, Ru4, and Ru5 addition to CT-DNA elicited a similar behavior (Fig. S37†).

To confirm whether Ru2–Ru5 interact with CT-DNA through the grooves, we conducted competition assays by using Hoechst 33258, a fluorescent dye that interacts with DNA through the minor groove, furnishing a DNA-Hoechst complex that emits fluorescence at 460 nm when excited at 340 nm. Increasing Ru2 concentration decreased the fluorescence intensity of the CT-DNA-Hoechst complex (Fig. 4C), indicating that Ru2 interacted with DNA via the minor groove, displacing Hoechst from this region and suppressing the fluorescence. Ru3, Ru4, and Ru5 behaved similarly (Fig. S38†). However, their binding constant (K_b) values are higher for the complexes with a similar structure (Ru2–Ru4) and lower for the Ru5 complex (Table S5†).

Furthermore, we employed electrophoretic mobility shift assays in gel to analyze how Ru2–Ru5 interact with DNA. This assay involves analyzing how DNA moves through a solid phase, such as agarose, under an electrical potential difference. Given that DNA carries a negative charge, it tends to migrate toward the anode. The rate at which DNA moves depends on various factors, including size and conformation. Longer DNA fragments migrate more slowly than shorter ones.³³

For this analysis, we used plasmid pBR322 DNA, which is primarily in the supercoiled (SC) form and migrates rapidly, in addition to the linear form (LC), which exhibits intermediate migration, and the circular form (OC), which migrates more slowly than SC and LC. By incubating plasmid pBR322 DNA with different concentrations of one of the ruthenium complexes, we observed that the DNA migration rate remained largely unchanged in the presence of Ru2, Ru4, or Ru5 compared to the negative control (DNA only) (Fig. 6D). Therefore, these ruthenium complexes did not significantly alter the structure of plasmid pBR322 DNA. Additionally, Ru3 has been reported to interact with plasmid pBR322 DNA similarly to Ru2, Ru4, and Ru5.¹⁹

HSA is the most abundant protein in blood and plays a crucial role in carrying substances. Consequently, it can be an important carrier of metallodrugs through the bloodstream. We studied how Ru2–Ru5 interact with HSA, which exhibits intrinsic fluorescence due to amino acid residues, by fluorescence.

We prepared solutions containing HSA at a constant concentration and varying concentrations of one of the ruthenium complexes, to obtain different molar ratios. Fig. 5 presents the fluorescence spectra of HSA in the absence and presence of Ru2. The HSA fluorescence became less intense with increasing Ru2 concentration, indicating that Ru2 interacted with HSA. We verified the same behavior for Ru3–Ru5 (Fig. S39–S41†).

Quantitative analysis of the fluorescence quenching process allowed us to evaluate whether fluorescence was suppressed through a static or dynamic mechanism. Increasing temperature decreased the quenching constant (K_SV) of Ru2, Ru3, and Ru4, indicating that fluorescence was suppressed through a static mechanism. In this mechanism, HSA and the quencher (ruthenium complex) form a complex in the ground state, and increasing temperature destabilizes such complex, hence decreasing K_SV.

For Ru5, K_SV increased with rising temperature, indicating a dynamic mechanism. In this mechanism, a collision occurs between HSA in the excited state and the quencher (ruthenium complex), which returns to its ground state without emitting fluorescence. Increasing temperature induces more collisions, raising K_SV. However, k_q obtained for Ru5 was in the order of 10¹⁴ M⁻¹ s⁻¹, exceeding the maximum value for a dynamic mechanism (2.0 × 10¹⁰ L mol⁻¹ s⁻¹).²⁸ Moreover, k_q increased with rising temperature, suggesting that Ru5 interacted with HSA via both the dynamic and static mechanisms.

To evaluate the magnitude of the ruthenium complex–HSA interaction, we calculated the binding constant (K_b), which indicated that the interaction was moderate, as judged from the order of magnitude around 10⁴ M⁻¹ (Table 2). The number of binding sites was approximately 1, suggesting that the complexes bound to HSA in a 1:1 stoichiometry (Table S6†).

We assessed the thermodynamic parameters to investigate the type of interaction between the ruthenium complexes and HSA. ΔG indicated that the interaction was spontaneous at the studied temperature. Negative ΔH and positive ΔS, as observed for Ru2, Ru3, and Ru4, indicated electrostatic interactions, whereas positive ΔH and ΔS, as observed for Ru5, indicated hydrophobic interactions³⁹ (Table S5†).

The complexes that were structurally similar (Ru2, Ru3, and Ru4), with two phosphine ligands, interacted with HSA similarly, showing the same type of mechanism and interaction. Meanwhile, Ru5, bearing two bipyridines, exhibited different mechanisms and interactions. This characteristic could be related to the presence of the bipyridine rings, which arranged themselves more orderly around the metal center compared to the bidentate ligands. Such orderly arrangement potentially facilitated interaction of the bipyridine rings with hydrophobic regions of HSA.

Cytotoxicity assays

The cytotoxic activity of Ru2–Ru5 was investigated toward B16-F10 (murine melanoma), A-375 (human melanoma), and HaCat (non-tumoral human keratinocyte) cells. Table 3 lists IC₅₀ (concentration of a compound that can inhibit cell viability by 50%) at 24 h and selectivity indexes (SI).

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Ru2, Ru3, and Ru4, which contained two phosphine ligands, were cytotoxic toward all the investigated cell lines, whereas Ru5, with two bipyridine ligands, was not cytotoxic even at the highest evaluated concentration (200 μM). Therefore, phosphine ligands coordinated to the ruthenium metal center enhanced the cytotoxicity of the ruthenium complexes. Increased cytotoxic activity of ruthenium complexes upon introduction of phosphine ligands has also been observed for other complexes.⁴⁰

We investigated the cytotoxicity of cisplatin, as well, for comparison purposes. Ru2, Ru3, and Ru4 showed lower IC₅₀ for all the evaluated cell lines. For example, Ru4 was 4, 2, and 24 times more active than cisplatin toward HaCat, A-375, and B16-F10 cells, respectively. Ru4 was selective for A-375 cells compared to HaCat cells: the SI was 4.61, which made Ru4 the most selective ruthenium complex studied herein. Thus, we selected Ru4 to investigate antiproliferative activity by the clonogenic efficiency assay.

The effect of free ligands should also be mentioned. Although their cytotoxicity was not studied here, the literature reveals the absence of anticancer activity for bipy and dppm (an analogue to other phosphine-based ligands) on B16-F10 cancer cells (IC₅₀ > 200).⁴¹

The clonogenic efficiency assay allows one to determine the ability of a cell to proliferate indefinitely and to form colonies comprised of at least 50 cells. To this end, we seeded HaCat, A-375, or B16-F10 cells and exposed them to different Ru4 concentrations for 24 h. After this period, we added fresh culture medium without Ru4 to the plates and maintained them in the incubator for 10 days. This allowed us to evaluate how Ru4 affects colony formation, size, and number.

According to Fig. 6, Ru4 reduced the number of colonies in a concentration-dependent manner. Additionally, HaCat cells formed a larger number of colonies than A-375 and B16-F10 cells. This agreed with the selectivity observed in the cytotoxic activity revealed by the XTT assay. Thus, Ru4 exhibited cytostatic effects, which prevent cell growth, development, and multiplication. Ru4 was also cytotoxic because, depending on the concentration, it completely inhibited cell growth. This behavior is similar than observed for cisplatin in A-375 cancer cells.⁴²

Moreover, we investigated how Ru4 affects A-375 cell morphology. For this purpose, we treated A-375 cells with different Ru4 concentrations and captured images immediately after treatment and after treatment for 24 h (Fig. 7). After treatment for 24 h, A-375 cells presented morphological alterations, especially at 5, 10, and 20 μM Ru4 A-375 cells lost adhesion and were less dense. In addition, dead cells became more evident compared to the control (1% DMSO).

Cell migration is a fundamental process during the natural development of an organism and is important for wound healing, tissue repair and development, and defense. However, cell migration can contribute to the appearance of metastases, when a tumor cell migrates from a primary tumor to another region of the body, where it undergoes adhesion processes and gives rise to a new tumor. Therefore, a compound that inhibits cell migration is key for directly inhibiting or controlling metastasis.^43,44

In this sense, we investigated whether Ru4 inhibits cell migration. To this end, we seeded A-375 cells, and, after they reached confluence, we made a scratch on the adherent cell monolayer with sterile pipette tip. We added culture medium containing different Ru4 concentrations to the cells and captured images of the scratch immediately after we added Ru4 and after treatment for 24 h. Fig. 8 shows that 0.2 and 0.4 μM Ru4 inhibited cell migration given that the wound was not completely closed compared to the control (1% DMSO). Hence, Ru4 could potentially inhibit cell migration.

Conclusions

We synthesized and characterized five ruthenium complexes. Four of the ruthenium complexes (Ru1–Ru4) contained two diphosphines and one 2-mercapto-4(3H)-quinazoline as ligands, whereas one ruthenium complex (Ru5) contained two bipyridines and one 2-mercapto-4(3H)-quinazoline as ligands. In addition, we investigated the cytotoxic activities of Ru2–Ru5 against melanoma cell lines. All the complexes inhibited melanoma cell growth (B16-F10 and A-375 cells) in a dose-dependent manner. Ru4, bearing two dppen, exhibited four times greater activity against A-375 tumor cells compared to non-tumor HaCat cells. Additionally, Ru4 inhibited colony formation in both HaCat and A-375 cells (with a more pronounced effect on the latter cells), altered A-375 cell morphology, and inhibited cell migration at concentrations of 0.2 and 0.4 μM. Furthermore, we evaluated the ability of the ruthenium complexes to interact with biomolecules such as DNA and HSA by various analytical techniques. The ruthenium complexes interacted with DNA weakly, possibly through the grooves, and they interacted with HSA moderately. The ruthenium complexes bearing phosphine and mercapto as ligands displayed promising cytotoxic properties against melanoma. Our results confirmed our previous data on the cytotoxic activity of ruthenium complexes containing phosphines as ligands, which were even better than similar complexes containing bipyridine ligands.⁴⁰

Author contributions

Nádija N. P. da Silva: writing – original draft, validation, methodology, investigation, formal analysis, data curation, conceptualization. Marcos V. Palmeira-Mello: writing – original draft, methodology, formal analysis, data curation. Nathália O. Acésio: methodology, formal analysis, data curation. Carlos A. F. Moraes: methodology, formal analysis, data curation. João Honorato: methodology, formal analysis, data curation. Eduardo E. Castellano: methodology, formal analysis, data curation. Denise C. Tavares: methodology, formal analysis, data curation. Katia M. Oliveira: writing – original draft, validation, methodology, investigation. Alzir A. Batista: writing – review and editing, writing original draft, visualization, validation, supervision, funding acquisition, formal analysis, conceptualization.

Data availability

The data supporting this article have been included as part of the ESI.†

Crystallographic data for Ru1, Ru2 and Ru5 has been deposited at the CCDC 2355448 (Ru1), 2355449 (Ru2) and 2355450 (Ru5).†

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

The authors are grateful for the financial support of the Brazilian Research Agencies CAPES, CNPq, FAPESP, and FAPDF. K. M. Oliveira would like to thank Fundação de Apoio à Pesquisa do Distrito Federal (FAPDF, process 00193-00002088/2023-42) and Decanatos de Pesquisa e Inovação e da Pós-graduação (DPI/DPG) da Universidade de Brasília (UnB) for the financial support provided. N. N. P. da Silva thanks Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq, process 130556/2021-1) for financial support. M. V. Palmeira-Mello thanks Fundação de Apoio à Pesquisa do Estado de São Paulo (FAPESP, process 2021/01787-0). Alzir A. Batista thanks FAPESP for financial support (Processes 2023/02475-8 and 2017/15850-0).

References

1. Q. Lin, X. Guo, B. Feng, J. Guo, S. Ni and H. Dong, Comput. Biol. Med., 2024, 108549 CrossRef .
2. S. Carr, C. Smith and J. Wernberg, Surg. Clin. North Am., 2020, 100, 1–12 CrossRef .
3. C. Liang, P. Wang, M. Li, R. Li, K. P. Lai and J. Chen, Heliyon, 2024, 10, e28616 CrossRef CAS .
4. C. M. Manzano, D. H. Nakahata and R. E. F. de Paiva, Coord. Chem. Rev., 2022, 462, 214506 CrossRef CAS .
5. B. L. V. Rosenberg, Nature, 1969, 224, 488–490 CrossRef .
6. K. D. Mjos and C. Orvig, Chem. Rev., 2014, 114, 4540–4563 CrossRef CAS .
7. Q. Peña, A. Wang, O. Zaremba, Y. Shi, H. W. Scheeren, J. M. Metselaar, F. Kiessling, R. M. Pallares, S. Wuttke and T. Lammers, Chem. Soc. Rev., 2022, 51, 2544–2582 RSC .
8. L. Zeng, P. Gupta, Y. Chen, E. Wang, L. Ji, H. Chao and Z.-S. Chen, Chem. Soc. Rev., 2017, 46, 5771–5804 RSC .
9. I. Yousuf, M. Bashir, F. Arjmand and S. Tabassum, Coord. Chem. Rev., 2021, 445, 214104 CrossRef CAS .
10. A. Casini and A. Pöthig, ACS Cent. Sci., 2024, 10, 242–250 CrossRef CAS PubMed .
11. X. Xiong, L.-Y. Liu, Z.-W. Mao and T. Zou, Coord. Chem. Rev., 2022, 453, 214311 CrossRef CAS .
12. S. Thota, D. A. Rodrigues, D. C. Crans and E. J. Barreiro, J. Med. Chem., 2018, 61, 5805–5821 CrossRef CAS .
13. S. Monro, K. L. Colón, H. Yin, J. Roque, P. Konda, S. Gujar, R. P. Thummel, L. Lilge, C. G. Cameron and S. A. McFarland, Chem. Rev., 2019, 119, 797–828 CrossRef CAS .
14. C. Sumithaa and M. Ganeshpandian, Mol. Pharm., 2023, 20, 1453–1479 CrossRef CAS .
15. S. M. Meier-Menches, C. Gerner, W. Berger, C. G. Hartinger and B. K. Keppler, Chem. Soc. Rev., 2018, 47, 909–928 RSC .
16. G. F. Grawe, K. M. Oliveira, C. M. Leite, T. D. de Oliveira, J. Honorato, A. G. Ferreira, E. E. Castellano, M. R. Cominetti, R. S. Correa and A. A. Batista, Dalton Trans., 2022, 51, 1489–1501 RSC .
17. A. E. Graminha, C. Popolin, J. Honorato, R. S. Correa, K. M. de Oliveira, L. R. Godoy, L. C. Vegas, J. Ellena, A. A. Batista and M. R. Cominetti, Eur. J. Med. Chem., 2022, 243, 114772 CrossRef CAS PubMed .
18. G. F. Grawe, K. M. Oliveira, C. M. Leite, T. D. de Oliveira, A. R. Costa, C. A. F. Moraes, J. Honorato, M. R. Cominetti, E. E. Castellano, R. S. Correa, S. P. Machado and A. A. Batista, J. Inorg. Biochem., 2023, 244, 112204 CrossRef CAS .
19. M. M. da Silva, G. H. Ribeiro, M. S. de Camargo, A. G. Ferreira, L. Ribeiro, M. I. F. Barbosa, V. M. Deflon, S. Castelli, A. Desideri, R. S. Corrêa, A. B. Ribeiro, H. D. Nicolella, S. D. Ozelin, D. C. Tavares and A. A. Batista, Inorg. Chem., 2021, 60, 14174–14189 CrossRef CAS PubMed .
20. M. V. Palmeira-Mello, A. R. Costa, L. P. de Oliveira, O. Blacque, G. Gasser and A. A. Batista, Dalton Trans., 2024, 53, 10947–10960 RSC .
21. A. A. Batista, L. A. C. Cordeiro, G. Oliva and O. R. Nascimento, Inorg. Chim. Acta, 1997, 258, 131–137 CrossRef CAS .
22. B. P. Sullivan, D. J. Salmon and T. J. Meyer, Inorg. Chem., 1978, 17, 3334–3341 CrossRef CAS .
23. M. T. Bautista, E. P. Cappellani, S. D. Drouin, R. H. Morris, C. T. Schweitzer, A. Sella and J. Zubkowski, J. Am. Chem. Soc., 1991, 113, 4876–4887 CrossRef .
24. B. P. Sullivan and T. J. Meyer, Inorg. Chem., 1982, 21, 1037–1040 CrossRef CAS .
25. G. M. Sheldrick, Acta Crystallogr., Sect. C: Struct. Chem., 2015, 71, 3–8 Search PubMed .
26. O. V. Dolomanov, L. J. Bourhis, R. J. Gildea, J. A. K. Howard and H. Puschmann, J. Appl. Crystallogr., 2009, 42, 339–341 CrossRef CAS .
27. J. R. Lacowicz, Principles of Fluorescence Spectroscopy, Kluver Academic/Plenum Publishers, New York, 1999 Search PubMed .
28. M. Ganeshpandian, R. Loganathan, E. Suresh, A. Riyasdeen, M. A. Akbarsha and M. Palaniandavar, Dalton Trans., 2014, 43, 1203–1219 RSC .
29. N. A. P. Franken, H. M. Rodermond, J. Stap, J. Haveman and C. van Bree, Nat. Protoc., 2006, 1, 2315–2319 CrossRef CAS PubMed .
30. K. Nakamoto, Infrared and Raman spectra of inorganic and coordination compounds, New York, 4th edn, 2009 Search PubMed .
31. G. H. Ribeiro, A. P. M. Guedes, T. D. De Oliveira, C. R. S. T. b. De Correia, L. Colina-Vegas, M. A. Lima, J. A. Nóbrega, M. R. Cominetti, F. V. Rocha, A. G. Ferreira, E. E. Castellano, F. R. Teixeira and A. A. Batista, Inorg. Chem., 2020, 59, 15004–15018 CrossRef CAS PubMed .
32. K. M. Oliveira, J. Honorato, F. C. Demidoff, M. S. Schultz, C. D. Netto, M. R. Cominetti, R. S. Correa and A. A. Batista, J. Inorg. Biochem., 2021, 214, 111289 CrossRef CAS PubMed .
33. A. Kellett, Z. Molphy, C. Slator, V. McKee and N. P. Farrell, Chem. Soc. Rev., 2019, 48, 971–988 RSC .
34. T. Biver, F. Secco and M. Venturini, Coord. Chem. Rev., 2008, 252, 1163–1177 CrossRef CAS .
35. Y. Wei and L.-H. Guo, Environ. Toxicol. Chem., 2009, 28, 940–945 CrossRef CAS .
36. S. U. Rehman, T. Sarwar, M. A. Husain, H. M. Ishqi and M. Tabish, Arch. Biochem. Biophys., 2015, 576, 49–60 CrossRef PubMed .
37. M. V. Palmeira-Mello, A. B. Caballero, P. Herrera-Ramírez, A. R. Costa, S. S. Santana, G. P. Guedes, A. Caubet, A. A. Batista, P. Gamez and M. Lanznaster, J. Inorg. Biochem., 2023, 248, 112345 CrossRef CAS .
38. B. N. Cunha, L. Colina-Vegas, A. M. Plutín, R. G. Silveira, J. Honorato, K. M. Oliveira, M. R. Cominetti, A. G. Ferreira, E. E. Castellano and A. A. Batista, J. Inorg. Biochem., 2018, 186, 147–156 CrossRef CAS PubMed .
39. P. D. Ross and S. Subramanian, Biochemistry, 1981, 20, 3096–3102 CrossRef CAS PubMed .
40. R. A. De Grandis, P. W. da S. dos Santos, K. M. de Oliveira, A. R. T. Machado, A. F. Aissa, A. A. Batista, L. M. G. Antunes and F. R. Pavan, Bioorg. Chem., 2019, 85, 455–468 CrossRef CAS .
41. A. P. Carnizello, M. I. F. Barbosa, M. Martins, N. H. Ferreira, P. F. Oliveira, G. M. Magalhães, A. A. Batista and D. C. Tavares, J. Inorg. Biochem., 2016, 164, 42–48 CrossRef CAS PubMed .
42. G. C. Segat, C. G. Moreira, E. C. Santos, M. Heller, R. C. Schwanke, A. V. Aksenov, N. A. Aksenov, D. A. Aksenov, A. Kornienko, R. Marcon and J. B. Calixto, Invest. New Drugs, 2020, 38, 977–989 CrossRef CAS PubMed .
43. S. A. Eccles and D. R. Welch, Lancet, 2007, 369, 1742–1757 CrossRef CAS .
44. H. Yamaguchi, J. Wyckoff and J. Condeelis, Curr. Opin. Cell Biol., 2005, 17, 559–564 CrossRef CAS .

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Footnote

† Electronic supplementary information (ESI) available: Fig. S1 (infrared spectra of the Ru1–Ru5 complexes and H2mq ligand); Fig. S2 (cyclic voltammogram of the Ru1–Ru5 complexes); Fig. S3 and S4 (UV-vis spectra of the Ru1–Ru5 complexes); Fig. S5–S8 (31P{1H} NMR spectra of the Ru1–Ru5 complexes); Fig. S9–S28 (1H, COSY (1H–1H), 13C{1H}, HSQC (1H–13C), and HMBC (1H–13C) NMR spectra of the Ru1–Ru5 complexes); Fig. S29–S35 (31P{1H} NMR spectra of the Ru1–Ru5 complexes over time in acetone, DMSO, and culture medium); Fig. S36 (UV-vis spectra of Ru5 complex over time in DMSO/culture medium); Fig. S37 (circular dichroism spectra of CT-DNA in absence and presence of Ru3–Ru5 complexes); Fig. S38 (fluorescence spectra of CT-DNA-Hoechst and Ru3–Ru5 complexes); Fig. S39–S41 (fluorescence spectra of HSA in absence and presence of Ru3–Ru5 complexes); Table S1 (values of coupling constant 2JP–P of Ru1–Ru4 complexes of31P{1H} NMR spectra); Tables S2 and S3 (crystal data and structure refinement for Ru1, Ru2, Ru3 and Ru5 complexes); Table S4 (thermodynamic parameters of the Ru2–Ru5 complexes with HSA). CCDC 2355448 (Ru1), 2355449 (Ru2) and 2355450 (Ru5). For ESI and crystallographic data in CIF or other electronic format see DOI: https://doi.org/10.1039/d4dt02575j

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