Synthesis and structure of [(η⁶-p-cymene)Ru(2-anthracen-9-ylmethylene-N-ethylhydrazinecarbothioamide)Cl]Cl; biological evaluation, topoisomerase II inhibition and reaction with DNA and human serum albumin^†

Abstract

We have synthesized and evaluated the biological properties of a compound of the type [η⁶-p-cymene)Ru(EtATSC)Cl]Cl (1) where EtATSC = 2-anthracen-9-ylmethylene-N-ethylhydrazinecarbothioamide, a thiosemicarbazone. The complex has been characterized by elemental analysis, spectroscopically (NMR, UV-Vis, and IR) and structurally by XRD. The in vitro anticancer activity of 1 has been evaluated against two human colon cancer cell lines. The IC₅₀ value for activity against HCT-116 was 224 ± 7 μM and 205 ± 5 μM against the Caco-2 cell line. The proficiency of 1 as an antibacterial agent was also evaluated against six bacterial strains. The minimum inhibitory concentration for Bacillus cereus was determined to be 5 μM and for Enterococcus faecalis it was 20 μM. At the maximum concentration tested the complex showed no activity against the Gram-negative strains. The complex binds strongly to human serum albumin with a binding constant of 1.37 ± 0.02 M⁻¹ at 308 K on a single binding site. It is also a strong binder to DNA with an apparent binding constant of 2.82 × 10⁵ M⁻¹ at 308 K. 1 shows very good activity as a catalytic inhibitor of human topoisomerase II at concentrations as low as 20 μM.

Graphical Abstract

Synthesis of an organometallic complex bearing a thiosemicarbazone ligand establishes topoisomerase II as a viable cellular target.

1. Introduction

Ruthenium complexes of various types are actively studied as metallodrugs as they are believed to have low toxicity and good selectivity for tumors.¹ Organometallic compounds exhibit different and novel structural motifs and organometallic ruthenium(ii)-arene complexes (of the type [(η⁶-arene)Ru(LL)Xl]⁺, LL = ligand) are also currently being investigated as anticancer compounds.² Such organometallic complexes offer much scope for optimization of biological activity by variations of the three main building blocks-the monodentate ligand X, the bidentate ligand LL and the arene – to fine-tune the pharmacological properties of these complexes.³ As it relates to the chelating ligand the structure–activity relationships for these complexes is somewhat complex. It was originally believed that a NH on the chelating ligand was required (based on the parent complex for this class of compounds), but complexes with other ligands such as acetylacetone or the monodentate phosphine 1,3,5-triaza-7-phosphaadamantane (pta)⁴ have shown good activity. In our research group we focus on thiosemicarbazones as the chelating ligand. Thiosemicarbazones are of considerable pharmacological interest since a number of derivatives have shown a broad spectrum of chemotherapeutic properties. The wide range of biological activities possessed by substituted thiosemicarbazones includes cytotoxic, antitumor,⁵ antibacterial,⁶ and antiviral⁷ properties. The biological properties of the ligands can be modified and in fact enhanced, by the linkage to metal ions.^8–10

While it is believed that DNA is a primary target for these complexes, since DNA replication is integral to the progression of these diseases, there is also the recognition that the observed biological activity is not always related to their DNA-binding ability.² The drug binding to plasma and tissue proteins are also fundamental factors in determining the overall pharmacological activity of a drug.¹¹ Among serum proteins, human serum albumin (HSA) and alpha 1-acid glycoprotein (AGP) play important roles in the binding of many drugs, acting as reservoirs for a long duration of action, and ultimately affecting drug absorption, metabolism, distribution and excretion, properties that are of key importance to drug development.¹² Since HSA serves as a transport carrier for drugs, it is important to study the interactions of potential drugs with this protein. Knowledge of interaction mechanisms between drugs and plasma proteins is of crucial importance to the understanding of the pharmacodynamics and pharmacokinetics of a drug or drug prospect.

In this paper we report on a study (synthesis, characterization, biophysical reactivity and biological activity) of an organometallic ruthenium complex containing a thiosemicarbazone as the chelating ligand (Fig. 1).

Fig. 1

Structural representation of the ligand, EtATSC, used in this study and the organometallic complex it forms.

2. Experimental

Material and methods

Analytical or reagent grade chemicals were used throughout. All the chemicals including solvents were obtained from commercial vendors and used as received. Bacterial cultures were obtained from Carolina Biologicals (Burlington, North Carolina, USA). Microanalyses (C, H, N) were performed by Columbia Analytical, Tucson, AZ). Proton and carbon nuclear magnetic resonance (NMR) spectra were recorded in DMSO-d₆ on a Varian Mercury 300 spectrometer operating at room temperature. The residual ¹H and ¹³C present in DMSO-d₆ (2.50 and 39.51 ppm respectively) were used as internal references. Infrared (IR) spectra in the range 4000–500 cm⁻¹ were obtained using KBr pellets or using the ATR accessory on a Nicolet 6700 FTIR spectrophotometer. Cyclic voltammetric (CV) data were collected on a Bioanalytical Systems, Episilon potentiostat on a C3 cell stand at 296 K. CH₂Cl₂ solutions (1 × 10⁻³M) containing 0.1 mol L⁻¹tetrabutylammonium hexafluorophosphate were saturated with nitrogen for 15 min prior to each run. A blanket of nitrogen gas was maintained throughout the measurements. The measurements were carried out with a three-electrode system consisting of a platinumworking electrode, a platinum wire auxiliary electrode and a Ag–AgCl reference electrode. Ferrocene was used as an internal standard. The working electrode was polished before each experiment with alumina slurry. Fluorescence spectra were recorded on a Varian Cary Eclipse spectrophotometer. The topoisomerase II inhibition drug-screening kit (part number 1009-2) together with human topo II (p170) (part number 2000H-2) was obtained from TopoGen Inc. (Port Orange, Florida, USA).

Synthesis

The ligand, EtATSC, was synthesized as previously described.¹³ The starting metal compound, [(η⁶-p-cymene)RuCl₂]₂ was synthesized as described in the literature.¹⁴

EtATSC : ¹³C NMR (75.463 MHz, DMSO-d₆); δ = 14.56 (C18), 38.57 (C17), 124.83–130.83 (anthracyl moiety), 141.42 (C1), 176.91 (C16).

[(η⁶-p-cymene)Ru(2-anthracen-9-ylmethylene-N-ethylhydrazinecarbothioamide)Cl][Cl] (1)

The starting ruthenium dimer, [(η⁶-p-cymene)RuCl₂]₂, (201 mg, 0.33 mmol) was dissolved in 10 mL of dichloromethane and the orange-red solution degassed with argon for 15 min. To this solution was added the solid thiosemicarbazone (under a flow of argon) and the red mixture stirred for 3 h at room temperature. The volume of the reaction mixture was reduced to ∼50% and ether (10 mL) was added. The suspension that resulted was filtered and the solid washed with ether and dried at the vacuum line. The orange-red solid was recrystallized from dichloromethane and ether. Crystals suitable for X-ray analysis were grown by slow diffusion of ether into a concentrated dichloromethane solution. Orange-red solid. Yield: 0.342 g (86%). m.p. 188 °C. Analysis; Calcd for C₂₈H₃₁Cl₂N₃SRu: C, 54.81; H, 5.09; N, 6.85. Found: C, 54.00; H, 4.90; N, 6.75. ¹H NMR (300.08 MHz, DMSO-d₆): δ = 11.67 (1H, N_aH), 9.27 (1H, H1), 8.70 (1H, N_bH), 8.49 (2H, H4, H14), 8.35 (1H, H9), 8.15 (2H, H7, H11), 7.55–7.63 (4H, m, H5, H6, H12, H13), 3.57 (H17), 1.10–1.22 (H18 and CH(CH₃)₂), 5.78 (4H, m, p-cymene ring), 2.84 (1H, wm, CH(CH₃)₂), 2.08 (3H, s, CH₃-cymene ring). ¹³C NMR (75.463 MHz, DMSO-d₆); δ = 14.56 (C18), 124.83–130.89 (anthracyl moiety), 141.52 (C1), 176.84 (C16), 85.55–106.38 (C₆H₄), 17.91 (CH₃–C₆H₄), 21.52 (CH(CH₃)₂), 29.77 (CH(CH₃)₂).

X-Ray crystallography

A suitable crystal of 1 was mounted on a thin glass fiber on the goniometer head of a Bruker APEX 2 diffractometer equipped with a SMARTCCD area detector. Data were collected at room temperature using the instrument working in ω scan mode. The radiation source was a Mo tube (Kα radiation; λ = 0.71073 Å) with a highly oriented graphitemonochromator. Intensities were integrated from four series of 364 exposures, each covering 0.5° in ω within 20 to 60 s of acquisition time and the total data set being a sphere.¹⁵ The space group determination was done with the aid of XPREP software.¹⁶ Absorption corrections were applied based on crystal face indexing obtained using actual images recorded by video camera. The data processing was performed using the SADABS program that was included in the Bruker AXS software package.¹⁷ The structure was solved by direct methods and refined by least squares on weighted F² values for all reflections using the SHELXTL program.¹⁶ All atoms received assigned anisotropic displacement parameters and were refined without positional constraints. Crystallographic data and data collection parameters are listed in Table 1, while bond lengths and valence angles are summarized in Table 2. Figures for the crystal structures of the complex were drawn using ORTEP 32 software¹⁸ at a 50% thermal ellipsoids probability level. The PLATON checks of crystallographic data and actual CIF files as well as a complete list of bond lengths and angles may be found in the Supplementary Information.†

Table 1

Selected crystallographic and refinement data for 1·CH₂Cl₂

Parameter	Value
Empirical formula	C29H33 Cl4N3ORuS
Formula weight	698.54
T/K	296(2) K
Wavelength (Å)	0.71073 Å
Crystal system	Triclinic
Space group	P1̄
Unit cell dimensions	a = 10.9976(7) Å α = 82.6790(10)°.
	b = 11.9803(7) Å β = 79.5180(10)°.
	c = 12.8925(8) Å γ = 73.7410(10)°.
Volume/Å3	1598.22(17) Å3
Z	2
Densitycalcd (Mg m−3)	1.408
Absorption coefficient (mm−1)	0.752 mm−1
Crystal size/mm	0.41 × 0.37 × 0.11 mm3
θ range	1.61–28.28°
Index ranges	−14 ≤ h ≤ 14, −15 ≤ k ≤ 15, −17 ≤ l ≤ 17
No. reflections collected	19 401
No. independent reflections	7853 [R(int) = 0.0185]
Absorption correction	Numerical
Data/rest/param	7853/0/360
GOF F2	1.048
Final R indices; [I > 2σ(I)]	R1 = 0.0308; wR2 = 0.0829
R indices (all data)	R1 = 0.0373; wR2 = 0.0891
CCDC	797492

Table 2

Selected bond lengths [Å] and angles [°] for 1·CH₂Cl₂

Ru(1)–C(19)	2.222(2)	Ru(1)–Cl(1)	2.4119(6)
Ru(1)–C(23)	2.198(2)	Ru(1)–N(1)	2.1457(18)
Ru(1)–C(24)	2.246(2)	Ru(1)–S(1)	2.3532(6)
Ru(1)–C(25)	2.252(2)	N(1)–Ru(1)–S(1)	81.38(5)
Ru(1)–C(26)	2.172(2)	N(1)–Ru(1)–Cl(1)	81.78(5)
Ru(1)–C(26)	2.169(2)	S(1)–Ru(1)–Cl(1)	87.19(2)

DNA-interaction studies

All experiments involving the interaction of 1 with DNA were carried out in Tris buffer (1) (5 mM, 50 mM NaCl, pH 7.20). Stock solutions of ct-DNA was prepared by dissolving commercial nucleic acids in buffer and stored at 4 °C for 24 h to attain homogeneity. After dilutions, DNA concentration per nucleotide phosphate was determined spectrophotometrically using the molar absorption coefficient of 6600 M⁻¹ cm⁻¹ at 260 nm.¹⁹ A solution of ct-DNA in the buffer gave a ratio of UV absorbance at 260 and 280 nm of ≥1.8 indicating that the DNA was reasonably free from protein²⁰ and did not need further purification. The DNA stock solutions were stored at 4 °C and used within 4 days after their preparation. Doubly purified water used in all experiments was from a Milli-Q system.

Viscosity measurements

Viscosity studies were done using a Cannon-Manning semi-micro-dilution viscometer (type 75, Cannon Instruments Co., State College, PA, USA) immersed vertically in a thermostated water bath maintained at 31.0 ± 0.1 °C. The viscosity of DNA solutions was measured in the presence and absence of the metal complexes. The DNA concentration was maintained at 10 μM, while the complex concentration was varied from 0–90 μM. Data are presented as (η/η₀)^1/3vs. 1/R, where R = [DNA]/[complex], η is the viscosity of DNA in the presence of the complex and η₀ is the relative viscosity of DNA alone. Relative viscosity values were calculated from the observed flow time of DNA solution (t) and corrected for the flow time of buffer alone (t₀), using the expression η₀ = (t − t₀)/t₀. Flow time was measured with a digital stopwatch and each sample was measured three times and an average flow time was used.

Ethidium bromide and Hoechst 33258 displacement experiments

In the ethidium bromide (EB) fluorescence displacement experiment 3 mL of a solution, that is 10 μM DNA and 0.33 μM EB (saturated binding levels²¹), in Tris buffer was titrated with aliquots of a concentrated solution of the complex producing solutions with varied mole ratios of complex to ct-DNA. After each addition the solution was stirred at the appropriate temperature for 5 min before measurement. The fluorescence spectra of the solution were obtained by exciting at 520 nm and measuring the emission spectra from 530–700 nm using 5 nm slits. Temperature was controlled using a single-cell Peltier accessory. The procedure was the same for the Hoechst 33258 reactions using the following conditions: working solutions were 20 μM DNA and 2 μM Hoechst 33258; λ_ex = 338 nm and λ_em = 400–550 nm (with λ_max = 464 nm).

Chemical nuclease activity

The DNA unwinding and cleavage ability of the complex was evaluated by agarose gel electrophoresis of supercoiled pBR322 DNA. The experiments were done in the dark or with exposure to long-radiation UV light. Samples of pBR322 DNA (0.1 μg μL⁻¹) were incubated with the complexes (in concentrations ranging from 10 to 300 μM) in Tris buffer (50 mM Tris, 18 mM NaCl, pH 7.2) at room temperature for 1 h in the dark or subjected to 365 nm light. Samples containing sodium formate (400 mM), potassium iodide (400 mM) and sodium azide (100 mM) were included in the experiments. The reactions were quenched by addition of 3 μL of loading buffer (0.25% bromophenol blue and 15% Ficoll in water). Samples of the reaction mixtures were loaded onto a 1% agarose gel in TBE buffer (89 mM Tris, 89 mM boric acid, 2 mM EDTA, pH 8.2). The gels were subjected to electrophoresis for 1 h at 70 V, followed by staining with 0.5 μg mL⁻¹ethidium bromide for 30 min. The bands on the gel were visualized under UV light and photographed using a GEL Logic 440 Imaging System with a Kodak Molecular Imaging Software.

Effect on topoisomerase II activity

The capability of the ruthenium complexes to affect topoisomerase II activity was analyzed using gel electrophoresis. The reaction mixture was a total of 20 μL. Tris buffer (50 mM Tris, 18 mM NaCl, pH 7.2) was used as the reaction buffer for the complexes. The “complete buffer” was made fresh daily by adding equal volumes of 10× Topo II Assay Buffer A (0.5 M Tris–HCl (pH 8), 1.50 M NaCl, 100 mM MgCl₂, 5 mM dithiothreitol and 300 μg BSA mL⁻¹) to the 10× ATPBuffer B (20 mM ATP in water) and was used at 1/5 volume. The metal complexes were added to yield varying concentrations (20 μM, 50 μM, 100 μM, and 300 μM) from a 3 mM stock solution made fresh in a 1 : 1 solution of DMSO and Tris buffer. Etoposide used as a control poison was added from a 10 mM stock solution in DMSO to create a 1 mM concentration in the reaction. Also a control lane, in which DMSO was added to DNA at the highest concentration it appears in a single reaction, was included. DNA (pHOT-1) was used at a concentration of 12.5 ng μl⁻¹. Human topoisomerase IIα (3 units, 1.5 μl) was added and the reactions were immediately transferred from the ice bath to a 37 °C hot water bath for 30 min. The reactions were terminated by adding 1/10 volume of 10% sodium dodecyl sulfate. Proteinase K was added at a concentration of 50 μg ml⁻¹. The reactions were incubated in the hot water bath for an additional 15 min. 1/10 volume of 10× gel loading buffer was added and the reactions removed from the water bath. They were cleaned up via a CIAextraction. An equal volume of CIA (chloroform : isoamyl alcohol 24 : 1) was added, the mixture vortexed and then briefly centrifuged. The upper blue layer was split in half and placed into corresponding wells of two 1% agarose gels in Tris–Borate–EDTAbuffer (TBE). One gel contained 0.2 μg ml⁻¹ethidium bromide. The gels were run at 70 V for 2 h in TBE. The EB gel was run with TBE containing 0.5 μg ml⁻¹ethidium bromide. This gel was then de-stained in DNA-free water for 30 min while the other gel was stained in an ethidium bromide/buffer solution for 30 min. It was then de-stained as mentioned before. The bands on the gel were imaged under UV light using a GEL Logic 440 Imaging System with Kodak Molecular Imaging Software.

Reaction with human serum albumin (HSA)

For the fluorescence titration, a similar procedure to the EB displacement experiments was done. Solutions of HSA were prepared in Tris buffer (50 mM Tris, pH 7.40, 100 mM NaCl) and stored in the dark at 4 °C. The protein concentration was determined spectrophotometrically using the molar absorptivity of 36 000 M⁻¹ cm⁻¹ at 280 nm.²² In the experiments, a 3.0 mL solution of HSA (3 μM) was placed in a quartz cuvette and titrated with various amounts of a concentrated solution of the complex producing solutions with varied mole ratios of complex to HSA. The complex concentration ranged from 3–30 μM. The fluorescence spectra of the solutions were obtained by exciting at 295 nm and measuring the emission spectra from 300–500 nm.

Cell culture

Cell lines included two human colon cancer cells : HCT-116 (human colon carcinoma) and Caco-2 (human epithelial colorectal adenocarcinoma). In addition, normal human colon cells CCD-18Co (human colon fibroblasts), were included. All cell lines were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA) and maintained at the University of Rhode Island. Caco-2 cells were grown in EMEM medium supplemented with 10% v/v fetal bovine serum, 1% v/v nonessential amino acids, 1% v/v l-glutamine and 1% v/v antibiotic solution (Sigma). HCT-116cells were grown in McCoy's 5a medium supplemented with 10% v/v fetal bovine serum, 1% v/v nonessential amino acids, 2% v/v HEPES and 1% v/v antibiotic solution. CCD-18Co cells were grown in EMEM medium supplemented with 10% v/v fetal bovine serum, 1% v/v nonessential amino acids, 1% v/v l-glutamine and 1% v/v antibiotic solution and were used from PDL = 26 to PDL = 35 for all experiments. Cells were maintained at 37 °C in an incubator under a 5% CO₂/95% air atmosphere at constant humidity and maintained in the linear phase of growth. The pH of the culture medium was determined using pH indicator paper (pHydrion™ Brilliant, pH 5.5–9.0, Micro Essential Laboratory, NY, USA) inside the incubator. All of the test samples were solubilized in DMSO (<0.5% in the culture medium) by sonication and were filter sterilized (0.2 μm) prior to addition to the culture media. Control cells were also run in parallel and subjected to the same changes in medium with a 0.5% DMSO.

Cytotoxicity assay

The assay was carried out as described previously²³ to measure the IC₅₀ values for samples. Briefly, the in vitro cytotoxicity of samples were assessed in tumor cells by a tetrazolium-based colorimetric assay, which takes advantage of the metabolic conversion of MTS [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfenyl)-2H-tetrazolium, inner salt] to a reduced form that absorbs light at 490 nm. Cells were counted using a hemacytometer and were plated at 2000–5000 cells per well, depending on the cell line, in a 96-well format for 24 h prior to drug addition. Test samples and a positive control, etoposide 4 (mg mL⁻¹), were solubilized in DMSO by sonication. All samples were diluted with media to the desired treatment concentration and the final DMSO concentration per well did not exceed 0.5%. Control wells were also included on all plates. Following a 24 h, 48 h or 72 h drug-incubation period at 37 °C with serially diluted test compounds, MTS, in combination with the electron coupling agent, phenazine methosulfate, was added to the wells and cells were incubated at 37 °C in a humidified incubator for 3 h. Absorbance at 490 nm (OD₄₉₀) was monitored with a spectrophotometer (SpectraMax M2, Molecular Devices Corp., operated by SoftmaxPro v.4.6 software, Sunnyvale, CA, USA) to obtain the number of surviving cells relative to control populations. The results are expressed as the median cytotoxic concentrations (IC₅₀ values) and were calculated from six-point dose response curves using 4-fold serial dilutions. Each point on the curve was tested in. Data are expressed as mean ± SE for three replicates on each cell line.

Antibacterial activity

The in vitro antimicrobial activity of the ligand and 1 was investigated as the minimum inhibitory concentration (MIC) against a number of Gram-positive (Staphylococcus aureus, Bacillus cereus, Enterococcus faecalis) and Gram-negative (Escherichia coli, Pseudomonas aeruginosa, Salmonella typhimurium) bacteria strains. The bacteria were maintained in Mueller Hinton Broth and compounds were dissolved in DMSO. The minimal inhibitory concentration (MIC) was determined from a microdilution method. Approximately 30 mL of bacteria culture were incubated overnight at 37.0 °C to produce exponentially growing cells. This was then diluted to yield a suspension containing 1.5 × 10⁸ CFU/mL (based on comparison of the turbidity to a 0.5 McFarland standard). Subsequently, 250 μL of this bacteria mixture were then inoculated into a sterile 96-well microplate. Various amounts of the compounds were added to wells to give predetermined concentrations ranging from 2–50 μM. The well absorbances (at 600 nm) were recorded on a BioTek Synergy HT microplate reader immediately after inoculation. The microplates were incubated for 24 h at 37.0 °C, and then the absorbance recorded again after the 24 h. The MIC was defined as the lowest concentration of compound that did not produce any visible cell-growth or change in absorbance after incubation. Solvent, media and positive growth controls were included on each plate. Ampicillin or streptomycin was used as a standard comparison. Each plate contained three replicates of each concentration and two separate experiments were done.

3. Discussion

Synthesis and characterization

The ligand was synthesized by the acid-catalyzed condensation of 9-anthraldehyde with 4-ethyl-3-thiosemicarbazide.¹³ The complex 1 was synthesized by simply reacting the thiosemicarbazone with the ruthenium dimer at room temperature. Based on microanalytical and spectroscopic data we suggest that the complex is best formulated as [(η⁶-p-cymene)Ru(EtATSC)Cl]Cl (Fig. 1). The thiosemicarbazone ligand can undergo tautomerism and the proposal implies that it coordinates as the thione tautomer. This idea is supported by the infrared spectroscopic data. The amine region shows two peaks: one at 3412 cm⁻¹ corresponding to the amino N_b–H and the other at 3185 cm⁻¹ which is due to the hydrazinic N_a–H. This peak would have been absent had the thiol tautomer (as the ionic form) been the coordinated species. These peaks are shifted moderately to higher wavenumbers confirming the change in the electronic current as the ligand coordinates through the azomethine nitrogen and the thione sulfur. The coordination at the azomethine nitrogen is also obvious in the infrared spectra. The ν(C˭N) bands of thiosemicarbazones are sensitive to metal chelation providing evidence for metal coordination and the peak due to this chromophore shifts to 1597 cm⁻¹ from 1622 cm⁻¹ in the free ligand. The C–S peaks also change on coordination. The 8 cm⁻¹ shift (to lower wavenumbers) of the peak attributable to the thioamide IV ν(C˭S) band on complexation is usually interpreted as evidence for coordination through the neutral ligand.²⁴

Complex 1 is soluble in acetone, CH₂Cl₂ and DMSO but not as soluble in alcohols. There is evidence of chloridehydrolysis when 1 is dissolved in aqueous buffer. This hydrolysis reaction was studied by monitoring changes in the UV-Vis absorption spectrum of 1 dissolved in buffer 1. The appropriate amount of a concentrated solution of 1 in DMSO was quickly diluted to 10 mM with buffer and the changes in absorbance at 260 nm measured over a 60 min period. From this experiment we calculated the rate of aquation to be (1.18 ± 0.24) × 10⁻³s⁻¹ at an ionic strength of 50 mM NaCl at 298 K. Similar complexes, [(η⁶-p-X)Ru(en)Cl][PF₆], where X = biphenyl, 5,8,9,10-tetrahydroanthracene, and 9,10-dihydroanthracene and en = ethylenediamine, have been reported to have aquation rates of 1.23–2.59 × 10⁻³s⁻¹ at 298 K and 100 mM ionic strength.²⁵

The redox behavior of complex 1 has been studied by cyclic voltammetry in the anodic region at a platinum disc electrode in 1.0 mM dichloromethane solutions containing 0.1 M tetrabutylammonium hexafluorophosphate as the supporting electrolyte using a scan rate of 300 mV s⁻¹. The complex shows two consecutive irreversible oxidations at 0.975 V corresponding to the Ru(ii)/Ru(iii) couple and the second (unassigned) at higher potential (near 1700 mV vs.Ag/AgCl). This redox behavior is very similar to our previously reported compounds.²⁶

X-ray structure

The complex was obtained in a crystalline form suitable for X-ray diffraction by diffusion of ether into a concentrated CH₂Cl₂ solution. The complex belongs to the P1̄ space group and crystallizes with a molecule of dichloromethane that is disordered (see Supplementary information F1†). A summary of the crystallographic data for 1·CH₂Cl₂ is given in Table 1. The molecular structure of the cation of 1 is shown in Fig. 2. The compound adopts the piano-stool geometry that is common for ruthenium half-sandwich arene complexes. The metal is coordinated by the p-cymene which occupies three coordinated sites, a chloride and the chelating thiosemicarbazone ligand. The metal center has a slightly distorted octahedral coordination geometry with bite angles around the ruthenium involving the three legs of the piano-stool in the range of 81–87°. The relevant bond lengths and selected bond angles are given in Table 2. The Ru–Cl (2.4119(6)), Ru–N (2.1457(18)) and Ru–S (2.3532(6)) are comparable to other structurally characterized ruthenium arene complexes with similar N–S donor ligands.²⁶ Our values are on par with those obtained by Hamaker²⁷ for a series of complexes with an imine–thioether donor set. The average Ru–C(arene) bond of 2.209 Å is in line with our other characterized thiosemicarbazone complexes²⁶ as well as other ruthenium arene half-sandwich complexes.²⁸ The Ru–centroid distance is 1.699 Å which is almost exactly the same as our other thiosemicarbazone complexes and close to the values reported by the Hamaker group. The p-cymene ligand is also somewhat asymmetrically coordinated to the ruthenium.

Fig. 2

An OPTEP drawing at 50% thermal ellipsoids probability. A perspective view of the molecular structure and numbering scheme for the [η⁶-p-cymene)Ru(EtATSC)Cl]⁺ cation (1). Hydrogen atoms, the chloride counter-anion and disordered molecule of dichloromethane are not shown for clarity.

Interaction of 1 with ct-DNA

The mechanism of action for the biological activity of organometallic ruthenium compounds is not definitively known. It is likely that there are a number of biological targets including proteins. Still in step with most metal-based anticancer drugs, DNA seems an obvious and likely target. So we investigated the interaction of 1 with ct-DNA. Small molecules interact with DNAvia a number of mechanisms, namely intercalation, groove binding or electrostatic interactions. Considering the structure of the complex with the flat poly-aromatic moiety on the thiosemicarbazone ligand, we assumed that it could intercalate into DNA. It is also possible that covalent binding to the ruthenium does occur. The aquation rates for similar compounds are relatively high (though the rate depends on the chelating ligand) and the aquated complexes are generally believed to be more reactive. It is known that DNA binds very strongly (through a guanine residue) to such complexes with the aid of hydrophobic forces.²⁹

Viscometric studies

Among the most common methods used to investigate the possibility of an intercalative mechanism, hydrodynamic measurements (viscometry in particular) that are sensitive to DNA length changes are the most definitive. A classical intercalator will cause an increase in the viscosity of a DNA solution since the DNA helix must lengthen as base pairs are separated to accommodate the binding ligand.³⁰ On the other hand, complexes that bind exclusively in the DNA grooves by a partial or non-classical intercalation of the compound, (under the same conditions), can reduce its effective length, and concomitantly its viscosity by bending the DNA helix.³¹ The viscometric experiments involve measuring the viscosity of ct-DNA solutions (10 μM) containing various amounts of 1 (0–90 μM). The data were treated by plotting (η/η₀)^1/3vs. the binding ratio [1]/[ct-DNA] as shown in Fig. 3. It was observed that increasing the concentration of 1 led to an increase in the viscosity of the DNA solution followed by a leveling at higher concentrations of 1. From this figure we can conclude that the binding to DNA can occur via intercalation.

Fig. 3

Relative specific viscosities of ct-DNA in the presence of increasing amounts of 1 at 31.0 ± 0.1 °C. [DNA] = 10 μM.

Competitive binding experiments

To further investigate the binding mode between 1 and DNA, fluorescence competition experiments with EB and Hoechst 32258 were employed. EB is a planar cationic dye well-known to intercalate into the DNA helix. While EB is only weakly fluorescent, the EB–DNA adduct is a strong emitter on excitation near 520 nm. Quenching of the fluorescence may be used to determine the extent of the binding between the quencher (1) and DNA. One reason for the quenching is the reduction in the number of binding sites on the DNA that is available to the EB presumably because of competition with 1 which is non-emissive under the experimental conditions. As seen in Fig. 41 can compete effectively with EB for the DNA binding sites as there is a decrease in the fluorescence at 602 nm (by as much as 87% of the initial in some cases) as the amount of 1 is increased. This supports the idea that 1 can interact with DNA by the intercalative mode.

Fig. 4

Fluorescence spectra of the EB-bound ct-DNA in aqueous buffer in the absence and presence of increasing amounts of 1, λ_ex = 520 nm, [EB] = 0.33 μM, [DNA] = 10 μM, [1] (μM): 0–35 in 2.5 μM increments. T = 298 K. The inset is Stern–Volmer quenching plots at 298 K, 303 K and 308 K.

We can also perform a quantitative assessment of the interaction from the EB titration. This may be done by carrying out a Stern–Volmer analysis of the data. According to the Stern–Volmer eqn (1), the relative fluorescence is directly proportional to the concentration of the quencher.

$$\frac{F_0}{F}=1+K_{\text{SV}}[Q]$$

1

Here F₀ and F are the fluorescence intensity of the EB–DNA adduct before and after the addition of the complex, K_SV is the Stern–Volmer quenching constant and [Q] is the concentration of the quencher (in this case the complex). For a homogeneously emitting solution eqn (1) predicts a linear plot of F₀/F vs. [Q] but for many systems the plots have been found to curve upward.³² As seen in Fig. 4 (inset) this is case for 1. There are a number of possible reasons for the non-linearity and positive deviation of the Stern–Volmer plot. It is sometimes proposed that in addition to the dynamic quenching which governs the Stern–Volmer plot, a second mechanism, static quenching, is happening. The static contribution may be calculated from the following equation:

$$\frac{F_0}{F}=(1+K_{\text{D}}[Q])(1+K_{\text{S}}[Q])$$

2

where K_D and K_S are the dynamic and static quenching constants. (K_S would actually represent the formation constant for a dark complex between the quencher and the chromophores). However a completely unambiguous assignment of K_D and K_S requires fluorescence lifetime measurements³³ (though varying temperature can be used to characterize the quenching as predominantly dynamic or static). As implied above since linearity of the Stern–Volmer plot is usually associated with the fluorophore possessing a single binding site or multiple accessible binding sites, we suggest that deviation from linearity is due to the existence of non-equivalent (accessible vs. non-accessible) binding sites and/or the occurrence of combined quenching. Using the data from the linear portion of the Stern–Volmer plots (see Supplementary information F2†), the K_SV values were determined (Table 3). We can use the temperature-dependence of these values to suggest that the quenching is predominantly dynamic since higher temperatures would lead to faster diffusion and more collisional deactivation resulting in an increase in the quenching constant. For the possibility of non-equivalent binding sites it should be noted that only the fluorescence from the accessible fluorophore has the potential to be quenched. Consequently the apparent binding constant (K_app) for 1 with ct-DNA was determined by using eqn (3),34  

$$K_{\text{app}}=\frac{K_{\text{EB}}[EB]}{{[complex]}_{50\%}}$$

3

where K_EB = 1.2 × 10⁶ M⁻¹, the binding constant of EB to DNA and [complex]_50% is the concentration of 1 at 50% of the initial fluorescence. The binding constants and thermodynamic parameters for the process are given in Table 3. The binding constant is on the order of 10⁴ which indicate a strong binding affinity though not as large as that for a classical intercalator.

Table 3

Quenching, binding and thermodynamic parameters for the interaction of 1 with ct-DNA

Temperature (K)	KSVa (104 M−1)	K (105 M−1)	ΔG° (kJ mol−1)	ΔH° (kJ mol−1)	ΔS° (J mol−1 K)
298	3.94	2.10	−30.4	22.6b	178b
303	6.53	2.58	−31.4
308	18.2	2.82	−32.1

a

Calculated using only the data in the linear range.

b

From a van’t Hoff plot.

We stated above that the reduction in fluorescence of the EB–DNA adduct is due to replacement of some of the EB molecules from the adduct by 1. It could also be due to a direct quenching interaction on the DNA itself. In an effort to investigate this possibility and also to further examine the possibility of more than one binding site, we also carried out another fluorescence competition involving Hoechst 32258. The Hoechst 32258 dye is a well-known DNA groove binder and similar to EB, the Hoechst 32258–DNA adduct is a strong fluorescence emitter when excited at 338 nm (λ_em = 464 nm) while Hoechst 32258 is only weakly fluorescent. It was observed that the fluorescence from the Hoechst 32258–DNA complex is very effectively quenched suggesting that 1 can also bind to DNA in the grooves (see Supplementary information F3†). The Stern–Volmer plot shows a similar upward curvature to the EB–DNA reaction and the reasons are likely the same. Taken together, the results from both competition experiments and the viscosity experiments do suggest the possibility of variegate modes of binding to DNA for 1.

Cleavage of pBR322 DNA

The chemical nuclease activity of the complex was studied by examining its ability to convert supercoiled pBR322 DNA from Form I to Form II by agarose gel electrophoresis in the dark as well as after UV irradiation under aerobic conditions. When circular plasmid DNA is probed by electrophoresis, relatively fast migration is normally observed for the intact supercoil form (Form I). If scission occurs on one strand (nicking), the supercoil will relax to generate a slower-moving open circular form (Form II). Fig. 5 shows the electrophoretic separation of the DNA after incubation with the complexes in the dark and after irradiation. It is evident that the complexes have the ability to cause cleavage of the DNA under both sets of reaction conditions. There is an increase in this activity as the concentration of the complexes is increased (up to 100 μM). It is also evident that the cleavage is more pronounced when the reactants are irradiated. We also investigated the process in the presence of hydroxyl scavengers HCOONa (Lane 5) and KI (Lane 6) as well as the singlet oxygen-like species scavenger NaN₃ (Lane 7). These species seem capable of inhibiting the nuclease activity of the complexes with sodium azide appearing to be the strongest inhibitor. This might indicate a role for reactive-oxygen species in the interaction of the complexes with DNA.

Fig. 5

Agarose gel electrophoresis diagram for the cleavage of pBR322 DNA by 1 at ambient temperature in the dark and upon irradiation with 365 nm light under aerobic conditions. Irradiation time was 1 h and incubation time was 1 h. Lane DNA, DNA alone; Lane 1, DNA + 10 μM 1; Lane 2, DNA + 50 μM 1; Lane 3, DNA + 100 μM 1; Lane 5, DNA + 100 μM 1 + 0.40 M HCOONa; Lane 6, DNA + 100 μM 1 + 0.40 M KI; Lane 7, DNA + 100 μM 1 + 0.10 M NaN3. (Lane 4 contained the reaction mixture with ascorbic acid but that lane repeatedly ran anomalously).

Effect on topoisomerase II activity

Though DNA is implicated as the main target for anticancer ruthenium compounds, affecting the activity of type II topoisomerases (topo II) may also be an effective mode of anti-neoplastic activity. Topo II are a class of ubiquitous enzymes that modulate the topological problems associated with DNA replication, transcription, and other nuclear processes by introducing temporary single- or double-strand breaks in the DNA. Topo II is a major enzyme in neoplastic cells. Consequently topoisomerases have been one of the major molecular targets in anticancer drug development. The ability of 1 to affect the catalytic activity of human topo II was studied by a DNA relaxation assay. Supercoiled plasmid DNA was treated with topo II in the presence of varying concentrations of the complex and the products were examined by electrophoresis on agarose gel. Topo II-targeting agents control the topo II activity either by poisoning the enzyme or by inhibiting the enzyme. So depending on the nature of the products one can propose if the complex is a poison or an inhibitor. Topo II poisons act by increasing the concentration of covalent enzyme-cleaved DNA complexes. Catalytic inhibitors prevent topo II from carrying out its required physiological functions. Fig. 6 shows the gel electrophoretic diagram from the experiment. Enzyme activity is characterized by conversion of pHOT1DNA from the supercoiled conformation (SC, Form I, Lane 1) to the fully relaxed conformation (R, Form II; see lanes 2 and 3). The species migrating between the two forms are the various topoisomers. The complex can prevent the relaxation of the enzyme at concentrations as low as 20 μM (lanes 4–7) as evidenced by the absence of the topoisomers and the presence of only the supercoiled form of the DNA. So we propose that the complex can act as a strong catalytic inhibitor. There was no formation of linear DNA, which would be evidence for the formation of an enzyme-drug-DNA ternary complex, during the reaction which supports the preceding conclusion. Previously, strong anti-topoisomerase activity by arene-ruthenium complexes of the type [(benzene)Ru(DMSO)Cl₂] was reported.³⁵ Those complexes were observed to inhibit relaxation as well as promote formation of the cleavage complex or cross-linking with topoisomerase.

Fig. 6

Gel electrophorogram showing the effect of different concentration of 1 on the activity of human topo II. Lane 1: DNA (pHOT1); Lane 2: DNA & Enzyme; Lane 3: DNA, Enzyme & DMSO; Lanes 4–7: Ru concentrations 20, 30, 50, 100 μM; Lane 8: etoposide; Lane 9: Linear DNA Marker.

Reaction with HSA

Given the importance of HSA in moderating drug behavior we investigated the interaction of 1 with HSA. This was done by monitoring the decrease in the fluorescence intensity of HSA solutions on addition of 1. HSA has a well-known structure consisting of a single polypeptide chain. Of the amino acid residues in the chain, the single tryptophan (Trp 214) is responsible for the majority of the intrinsic fluorescence of the protein. HSA has a strong fluorescence emission with a peak near 350 nm upon excitation at 295 nm. Excitation at this wavelength is to reduce the effect of any tyrosinate emission. The emission is sensitive to the changes in the local environment of the tryptophan and so can be attenuated by binding of a small molecule at or near this residue.

Fig. 7 shows the changes in fluorescence intensity as increasing amounts of 1 are added. There is a significant reduction in the fluorescence from the initial amount with a blue shift to 329 nm (Δλ = 18 nm). These changes indicate that the conformation of the protein is affected by binding to 1. The binding may be quantitated starting with the Stern–Volmer eqn (1). As seen in Fig. 7, there is significant upward curvature of the Stern–Volmer plot (at the highest temperature). For protein solutions having homogeneous emission, as would be expected from HSA with its single tryptophan residue, upward curvature has been observed with diverse quenchers.³² In fact the fluorescence of most proteins is likely heterogeneous³² and in the case of HSA one cannot completely eliminate tyrosine emission.

Fig. 7

Emission spectra of the HSA in the absence and presence of increasing amounts of 1, λ_ex = 295 nm, [HSA] = 5.0 μM and [1] (μM): 0–35 in 2.5 μM increments. T = 301 K. The inset is Stern–Volmer quenching plots at 296 K, 301 K and 308 K.

In addition to heterogeneity, the upward curving Stern–Volmer plots may indicate both static and dynamic quenching and for proteins the quenching constants are nearly identical. Under these circumstances the binding constant can be obtained from a modified Stern–Volmer (MSV) analysis.³² In this analysis the following eqn (4) is used:

$$\frac{F_0}{\Delta F}=\frac{1}{fK[metalcomplex]}+\frac{1}{f}$$

4

where ΔF = F₀ − F (F₀ = initial fluorescence of the HSA solution). f is the fraction of HSA molecules that is initially accessible to 1 and K is the binding constant (assuming the decrease in fluorescence comes from the interaction of HSA with 1). f may be treated as the number of binding sites. It is obtained from the intercept of the linear plot of F₀/ΔF vs. 1/[Ru] (Fig. 8). The ratio of the intercept to the slope gives K. The binding constants obtained from this analysis along with their attendant thermodynamic parameters are given in Table 4.

Fig. 8

Plot of the modified Stern–Volmer equation: F₀/(F₀ − F) vs. 1/[Ru]. λ_ex = 295 nm, [HSA] = 5.0 μM and [1] (μM): 0–35.

Table 4

Binding and thermodynamic parameters for the interaction of 1 with HSA

Temperature (K)	Ka (104 M−1) (f)	R2	Kb (104 M−1) (n)	R2	ΔG° (kJ mol−1)a	ΔH° (kJ mol−1)	ΔS° (J mol−1 K)
296	6.31 ± 0.23 (1.1)	0.994	6.41 ± 0.12 (1.1)	0.979	−27.6	48.3c	255c
301	9.82 ± 3.42 (1.2)	0.995	11.4 ± 0.19 (1.1)	0.999	−28.8	43.2d	239d
308	13.7 ± 0.02 (1.1)	0.999	13.8 ± 0.02 (1.1)	0.999	−30.2

a

The values of K are from the modified Stern–Volmer analysis.

b

The values of K are from the Scatchard analysis.

c

From the van’t Hoff plot using modified Stern–Volmer values.

d

From the van’t Hoff plot using Scatchard values.

The table also show the binding constants obtained from a Scatchard analysis (which is still a popular way of representing data) (Fig. 9). This analysis is based on eqn (5).

$$\frac{r}{C_{\text{f}}}=nK+rK$$

5

In this equation r is the number of moles of 1 bound per mole of HSA. C_f is the molar concentration of the free metal complex, n is the number of binding sites and K is the intrinsic binding constant.

Fig. 9

The Scatchard plot for the binding of 1 to HSA λ_ex = 295 nm, [HSA] = 5.0 μM and [1] (μM): 0–35.

Both analyses yielded similar results with the binding constant on the order of 10⁴–10⁵ M⁻¹, indicating strong binding, and a single binding site. The reaction is spontaneous with ΔG (calculated from ΔG = −RTlnK) being negative. The ΔH and ΔS values were obtained from a van't Hoff plot of the variation of K with temperature (see Supplementary information F4†). By considering the nature of the signs (both positive) of these parameters we can propose that hydrophobic interactions are keys to the reaction of 1 with HSA.³⁶

The blue shift that occurred during the fluorescence titration suggests that the environment of the tryptophan residue became more nonpolar. When the tryptophan is in a polar environment the emission maximum is near 350 nm. It is closer to 325 nm when the environment is more nonpolar. Such a transformation would require significant reorganization of the protein possibly beyond the usual stochastic structural fluctuations. This would likely involve the expulsion of water molecules, the nonpolar pocket is created and the system becomes more random. This idea might be implicit in the large value (255 J mol⁻¹ K⁻¹) that was obtained for ΔS. We can also suggest therefore that the process is mostly entropy driven.

Biological activity

Anticancer activity

The in vitro cytotoxicity of complex 1 against two human cancer cell lines, HCT-116 (colon carcinoma) and Caco-2 (epithelial colorectal adenocarcinoma) and a non-cancerous cell line, CCD-18Co (colon fibroblasts) was investigated using a tetrazolium-based (MTS) colorimetric assay. Etoposide, a potent anti-neoplastic drug, was used as a standard comparison treatment. The IC₅₀ values (Table 5), the median cytotoxic concentrations, were determined after 24, 48 and 72 h of drug exposure. Generally, the longer the exposure time the more cytotoxic is the complex with the 72 h exposure being ∼20% more effective compared to the 24 h exposure. As another generality, the complex is slightly more active against the Caco-2 cells when compared to the HCT-116cell line. At 72 h exposure the cytotoxic effect on the non-cancerous cell line is noticeably less than on the two cancer lines. The small effect is true even for the standard drug treatment with etoposide being about ∼2.5 times more lethal to the cancer cellsversus the non-cancerous cells. The IC₅₀ values are in the hundreds micromolar range indicating the weak cytotoxicity of the complex. This result is significantly different than what we discovered²⁶ for similar complexes against the same cell lines as well as breast cancer (MCF-7 and MDA-MB-231) cells. Under the reaction conditions it is 12 times less effective than etoposide against the cancer cell lines. This is in itself not a serious issue. It is known that some ruthenium arene complexes have low in vitro toxicity but show good in vivo characteristics. For example complexes of the type [(cymene)Ru(PTA)Cl]⁺ exhibited low activity against cancer cells but had very good anti-metastatic activity in vivo.⁴

Table 5

IC₅₀ values (μM) representing the anti-proliferative activity of 1 in panel of three human celllines-2 tumorigenic (HCT-116 and Caco-2) and one non-cancerous (CCD-18Co)

Assay time	IC50
	HCT-116	Caco-2	CCD-18Co
24 h	290 ± 10	256 ± 4	298 ± 9
48 h	248 ± 8	235 ± 6	271 ± 8
72 h	224 ± 7	205 ± 5	253 ± 6
Etoposide (72 h)	18.3 ± 1.4	16.5 ± 2.0	42.2 ± 1.0

Antimicrobial activity

The in vitro antimicrobial efficiency of the ligand and the complex against a number of Gram-positive and Gram-negative bacteria has been determined as the minimum inhibitory concentrations (see Supplementary information T1†). The compounds were tested at low concentrations due to solubility issues and to avoid excessive concentrations of the solventDMSO in the wells. The complex was more active against the Gram-positive bacteria than the Gram-negative species showing excellent activity against Bacillus cereus (IC₅₀ = 5 μM) and moderate activity against Enterococcus faecalis (IC₅₀ = 20 μM). The complex was also found to be more active than the free ligand. This is not unusual as it is known that complexation to a metal ion may induce improved biological activity^8,37 and in some cases even different biological activity. Such complexation could enhance the lipophilic character of the compound which favors its permeation through the lipid layers of cell membrane. Although the complex is active, at least against certain species, it typically did not achieve the effectiveness of the conventional antibacterial standards ampicillin or streptomycin.

Conclusion

We have synthesized an organometallic ruthenium complex that is biologically active. The complex shows weak in vitro anticancer activity and also shows moderate antibacterial cytotoxicity primarily against Gram-positive bacteria. The compound binds strongly to human serum albumin and is a moderate multi-mode binder to DNA. It also shows excellent potential as an anti-topoisomerase II agent. With respect to the poor anticancer behavior, the complex is not dissimilar from other ruthenium arene complexes under active investigation and this complex may still have the potential for drug development.

Acknowledgements

The project described was supported by Award Number P20RR16460 from the National Centre for Research Resources to FAB. The content is solely the responsibility of the authors and does not necessarily represent official views of the National Centre for Research Resources or the National Institutes of Health.

Footnotes

†

Electronic supplementary information (ESI) available: Molecular structure and numbering scheme for the [η6-p-cymene)Ru(EtATSC)Cl]⁺. CCDC reference number 797492. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c1mt00003a