Conformational control of anticancer activity: the application of arene-linked dinuclear ruthenium(II) organometallics

Abstract

Dinuclear metal complexes have emerged as a promising class of biologically active compounds which possess unique anticancer activity. Here, we describe a novel series of arene-linked dinuclear organometallic Ru(II) complexes, where the relative conformation of the ruthenium centres is controlled by the stereochemical configuration of 1,2-diphenylethylenediamine linker moieties, as confirmed by X-ray crystallography. The reactivity and cytotoxicity of these compounds is compared to flexible dinuclear and mononuclear analogues, demonstrating in all cases the complexes can undergo aquation, coordinate to typical biological donor ligands and importantly, in the case of dinuclear analogues, crosslink oligonucleotide and peptide sequences. Differences in the conformation of the isomeric dinuclear compounds lead to significantly different levels of cytotoxicity against A2780, A2780cisR and HEK-293 cell lines; isomers with a closed conformation are significantly more cytotoxic than isomers with a more open conformation and they are also significantly less susceptible to acquired resistance mechanisms operating in the A2780cisR cell line. These rigid dinuclear compounds possess markedly increased cytotoxicity relative to the non-cytotoxic mononuclear analogues that does not appear to be related to differences in complex lipophilicity or cellular uptake, which, in general, remain similar in magnitude across the series. Thus, the molecular conformation of such dinuclear species may be crucial in determining the nature of the adducts formed on coordination to biological targets in a cellular environment, and opens up a novel route toward the development of more active metal-based anticancer agents.

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Introduction

Metal-based compounds offer considerable potential in medicinal chemistry where the careful choice of metal may afford compounds possessing geometrical, coordination and potentially catalytic properties not accessible through purely organic molecules. As a result, the judicious combination of a metal ion and associated ligands may result in complexes capable of unique biological activity. In the context of developing metal-based drugs for the treatment of cancer, considerable effort has been directed toward the development of novel platinum complexes,¹ stemming from the clinical success of cisplatin (and subsequent derivatives).

Apart from the platinum family of metallodrugs several other metals have been utilised in the development and identification of further classes of compounds that exhibit favourable medicinal attributes. Of these, ruthenium-based organometallic compounds constitute a rapidly developing field that continues to yield complexes exhibiting diverse biological activity.² Prominent examples include the [Ru(η⁶-arene)(en)Cl]⁺ family of organometallics that has yielded compounds with a comparable cytotoxicity to that of cisplatin in certain cell lines.³ Structurally related compounds based on the [Ru(η⁶-arene)(L)Cl]⁺ scaffold, with organic ligands (L) chosen because of their various biological activities, also exhibited significant antiproliferative activity against a range of cancer cell lines.^4–6 An alternative strategy, based on the development of kinetically inert ruthenium half-sandwich complexes as potent inhibitors of protein kinases, has also led to complexes exhibiting high cytotoxicity against the HCT-116 cell line.⁷ Ru(II)–arene complexes have also been incorporated into multinuclear systems and assessed for their anticancer activity. These have included metalla-cycles,^8–12 metalla-cages^13–15 and dendrimer-based systems.¹⁶ Of particular interest to us are the [Ru(η⁶-arene)Cl₂(PTA)] (RAPTA) series of organometallic ruthenium(II) compounds which display selective activity on metastatic tumours in vivo.^17,18 Recent work observed that proteins are a major intracellular target of the RAPTA compounds,¹⁹ including the histone proteins of the nucleosome core particle. Structural studies also show that RAPTA-C binds to the protein component of the nucleosome core particle in preference to the nucleic acid component.²⁰ In addition, subcellular localisation of RAPTA-T in A2780 and A2780cisR cells revealed higher metal to protein ratios in the particulate fraction (including the mitochondria) relative to the cytosol and nuclear fractions.¹⁹ Further accounts have described selective binding of RAPTA-C within protein mixtures²¹ and also potent enzyme inhibition by RAPTA compounds.^22,23 These studies offer an insight into the potential of the RAPTA compounds to exert their biological activity through protein interactions rather than DNA damaging mechanisms characteristic of the vast majority of platinum metallodrugs reported so far. We reasoned that dinuclear analogues of the RAPTA series may exhibit a different spectrum of biological activity compared to mononuclear RAPTA derivatives, potentially retaining the propensity of the mononuclear RAPTA compounds to bind proteins while acting via crosslinking of target biomolecules through long-range interactions rather than short range interactions typical of mononuclear species. In the field of platinum metallodrugs the formation of dinuclear analogues has been a successful route toward the development of compounds capable of unique binding modes and interactions with DNA.²⁴ This approach has led to compounds of increased potency compared to established mononuclear platinum compounds which retain high activity in cell lines resistant to cisplatin.^25,26 In contrast, dinuclear organometallic ruthenium(II) arene compounds are underexplored.²⁷ In the most comprehensive of these studies to date a series of dinuclear ruthenium compounds linked via maltol-derived ligands were investigated.^28–30 The cytotoxicity of these compounds was found to be tuned by modification of the chain length of the alkyl component of the linker ligand, with cytotoxicity correlating well to the experimentally determined lipophilicity of the resulting complexes.²⁸ Of notable interest is the observation that not only could these complexes crosslink two DNA duplexes but also that DNA–protein crosslinks could be formed, potentially a novel mode of action for such complexes.³⁰

We hypothesised that the use of linker ligands rigid enough to fix the relative orientation of the metal centres in a dinuclear RAPTA complex would allow access to a series of isomeric structures of differing conformation. Such complexes would allow an assessment of the effect of conformation on biological activity to be probed, including the manner in which isomeric complexes of different conformation interact with potential target biomolecules.

Here we report the synthesis, binding studies with potential biological targets and in vitro biological evaluation of a series of dinuclear Ru(II)–arene compounds, where the conformation of the RAPTA units relative to each other is controlled by the stereochemical configuration of a 1,2-diphenylethylenediamine (DPEN) linker molecule. Compounds with a linking group possessing either an (R,R)- or (S,S)-configuration exhibit a more ‘closed’ conformation whereas those with a (R,S)-configuration possess an open conformation not dissimilar to that of a complex linked via a flexible linker (Fig. 1). Such conformational control could significantly influence the interactions of each complex with biomolecular targets, ultimately yielding a different range of adducts, and potentially controlling the cytotoxicity and selectivity.

Fig. 1 Newman projections of predicted conformations of dinuclear complexes 4a–6a and 4b–6b based on ligand steric demand.

In addition to the rigid dinuclear complexes described above a further dinuclear complex, linked together using the flexible ethylene diamine analogue, and a mononuclear analogue are also reported. Our approach was to develop dinuclear Ru(II)–arene compounds linked together through the arene ligand in order to retain the core coordination environment around ruthenium as in the original RAPTA series to avoid significantly perturbing the coordination mode of these dinuclear complexes with potential biological targets.

Results and discussion

Synthesis and characterisation

The synthesis of the ruthenium dimers, and the mononuclear analogue, proceeded via amide-forming reactions between the common intermediate complex, 1, bearing a carboxylic acid substituent on the arene ligand, and the selected amines (Scheme 1). Complex 1 includes a single chelating oxalato-moiety as a protecting group in order to suppress undesired reactions with the ruthenium centre during amide bond formation. Reactions between 1 and the desired amines were performed utilising the coupling agent O-(benzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium tetrafluoroborate (TBTU) with N,N-diisopropylethylamine (DIPEA) in DMF, to either yield 3a and 4a as DMF-insoluble yellow powders that were isolated by filtration, or monomer analogue 2a and dinuclear compounds 5a and 6a as crude yellow powders following precipitation with acetone. All compounds were further purified by recrystallisation until analytically pure (see ESI†). Each oxalato-protected ruthenium complex was successfully converted to its chlorido-analogue by dissolution in an anhydrous solution of HCl in methanol – the yellow solutions immediately turned red in colour followed by precipitation of the desired dinuclear compounds 3b–6b as their HCl salts, or in the case of monomer analogue 2b the product was obtained by precipitation with diethyl ether. All compounds (1–6b) are water soluble; the oxalato-protected compounds (2a–6a) possess low solubility in DMSO and other organic solvents. All compounds were characterised by ¹H, ³¹P and ¹³C NMR, high-resolution mass spectrometry and elemental analysis (see ESI†).

Scheme 1 Synthesis and numbering scheme of the mononuclear oxalato-complex 2a, dinuclear oxalato-complexes 3a–6a and chlorido-analogues 2b–6b.

The molecular structures of 1, 2a, 4a, 5a and 6a (Fig. 2 and S1†) were confirmed by single crystal X-ray crystallography on crystals grown via the slow diffusion of diethyl ether into a methanolic solution of the complex (2a), or the slow diffusion of acetone into a solution of the complex in H₂O and acetone (1, 4a, 5a and 6a). In each case the coordination environment around the central ruthenium ions is similar, adopting the characteristic “piano-stool” configuration. Structural data confirms the stereoisomeric complexes, 4a, 5a and 6a, occupy conformations predicted by the Newman projections of the DPEN linkers (Fig. 1). For the enantiomeric complexes 5a and 6a linked by (1S,2S)-DPEN or (1R,2R)-DPEN respectively, the structures are of a concave conformation with the trans-geometry of the phenyl rings of the linker backbone forcing the two RAPTA units into positions adjacent to one another.‡ In contrast, the conformation of the diastereomeric complex 4a linked by (1R,2S)-DPEN is found to be more linear with the two RAPTA substituents being related through a centre of inversion.

Fig. 2 Molecular structures of 1 (top left), 2a (top right), 4a (middle) and 5a/6a (bottom). Hydrogen atoms have been omitted for clarity. Atoms are colour coded: ruthenium – cyan, phosphorus – orange, oxygen – red, nitrogen – purple, carbon – grey. See ESI† for figures of the compounds including atom labels.

¹H-NMR spectra of the complexes in D₂O shows that the central linker significantly influences molecular conformation. The spectrum of 3a (Fig. S30†), in which the RAPTA-units are linked through a flexible ethylene diamine moiety, is relatively simple. Both sets of methylene protons and arene protons manifest as a single set of resonances. In contrast, the spectra of dinuclear complexes constructed with the stereoisomeric 1,2-diphenylethylenediamine linking groups are more complex (Fig. 3), the form of each being dependent on the stereochemical configuration of the linker used. The ¹H NMR spectra of the enantiomeric complexes 5a and 6a are identical and possess four sets of Ru-coordinated arene resonances which are slightly shielded relative to equivalent resonances observed in the spectra of 1–3a. The methylene protons give rise to a complex set of resonances between 2.4–2.6 ppm whereas resonances corresponding to the aromatic protons of the linker consist of two peaks between 7.1–7.3 ppm. For the diastereomeric complex, 4a, two of the four Ru-coordinated arene resonances overlap, with all arene resonances having a different chemical shift compared to those of 5a and 6a. Interestingly, one of the arene resonances of 4a (5.08 ppm) is significantly more shielded relative to the other arene resonances. In addition, the resonance set corresponding to the methylene protons is more shielded and occurs between 2.16–2.33 ppm. These observations correlate well with the observation in the X-ray structure of 4a that the corresponding protons are shown to reside above the plane of the linker phenyl groups – a magnetic environment which would provide shielding from the applied magnetic field. Furthermore, in the spectrum of 4a the linker phenyl protons give rise to a single resonance at 7.4 ppm. The increased complexity in the spectra of 4a–6a relative to the spectrum of 3a reflects the increased central rigidity present in the former complexes due to the DPEN linkers. In addition, the shielding observed in the spectrum of 4a provides direct evidence that the conformation observed in the solid state is also preserved in aqueous solution.

Fig. 3 ¹H NMR spectra of 4a (top) and 5a (bottom) (D₂O, 400 MHz, 298 K, 10 mM). Resonances are identified with coloured circles: linker phenyl – red, Ru arene – blue, CH – green, PTA – orange, CH₂ – black and CH₃ – purple.

Aquation and stability studies

Many metal complexes, including cisplatin and several families of organometallic ruthenium complexes, are activated on cellular internalization (low [Cl⁻]) by aquation of labile metal–chlorido bonds, the aquated species being significantly more labile toward donor groups of biological targets.³¹ Early work¹⁷ with a prototypical example of the RAPTA series, RAPTA-C, revealed that upon dissolution in pure water or 4 mM NaCl solution the complex rapidly undergoes exchange of both chlorido ligands for aqua ligands. In 100 mM saline solution this process is completely suppressed.

Oxalato- and chlorido-containing ruthenium compounds synthesised in this work were assessed for their stability and reactivity in phosphate buffer (10 mM) at pH 7.2 in the presence of either 5 mM or 100 mM NaCl at 298 K. The UV-vis spectra of the chlorido analogues 2b–5b immediately change upon dissolution in phosphate buffer (10 mM) containing NaCl (5 mM), indicative of a change in coordination environment at the Ru centre. The half-lifes of the chlorido complexes (2b–5b) were estimated from the change in absorbance with time (see Fig. S5–S9†). In phosphate buffer 3b possessed the shortest half-life (8.5 min) followed by 2b (10.5 min), 4b (13.6 min) then 5b (17.3 min), the more closed structure of the latter compound presumably slows the aquation process for this complex. For comparison, under these conditions RAPTA-C has a half-life shorter than all compounds described here (6.8 min). These results indicate that although the different structures and conformations of compounds 2b–5b affect the kinetics of ligand exchange at ruthenium, the ‘active’ aqua species of each complex may be readily formed on dissolution in low [Cl⁻] aqueous solution. In the presence of elevated NaCl concentrations (100 mM) no aquation of these compounds was observed, in accordance with previous results.¹⁷ The oxalato-protected compounds 2a–5a were also examined by ³¹P NMR spectroscopy in the presence of HEPES–phosphate buffer (5 mM or 100 mM NaCl), and in the presence of BSA, CT-DNA and glutathione. In each case the appearance of no new ³¹P-NMR signals were observed over a period of 72 h at 310 K, demonstrating that the oxalato complexes are inert towards aquation or ligand exchange under ambient conditions on the timescale of the experiment. In contrast, on incubation of 2a–5a in RPMI or DMEM media used for cell culture it was found that a new resonance at −32.4 ppm gradually appears in the ³¹P NMR spectra over 48 h, with incubation of the complexes in DMEM resulting in the largest transformation (Fig. S10–S16†). The new resonance at −32.4 ppm may be attributed to the exchange of the oxalato ligand for a carbonato ligand as sodium bicarbonate is present in the RPMI and DMEM media used at concentrations of 23.8 mM and 44.0 mM respectively. Subsequent experiments involving the incubation of 2a with NH₄CO₃H (23.8 mM) in phosphate buffer (pH 7.4, 50 mM) confirmed this hypothesis, as the ³¹P NMR spectrum after 24 h exhibits the same transformation as the spectra recorded in RPMI/DMEM. The oxalato-protected complexes 2a–6a are stable toward aquation and inert in the presence of typical biological ligands, whereas media containing bicarbonate at concentration levels typically found in biological systems (10–30 mM), results in the gradual exchange of the oxalato ligand for a carbonato ligand, providing a possible route toward the intracellular activation of the oxalato-protected compounds.

Amino acid and nucleotide binding studies

In order to assess the ability of the complexes to coordinate to and crosslink biological targets, binding studies were performed with guanosine 5′-monophosphate, L-histidine and a range of model oligonucleotides and peptides.

To establish whether each ruthenium ion of the dinuclear compounds 3b–6b could coordinate simultaneously to target ligands, binding studies were performed initially with guanosine 5′-monophosphate and L-histidine – both being good ligands for RAPTA-type compounds as constituents of DNA or proteins, respectively.^20,32 For each dinuclear complex 3b–6b, following incubation in the presence of 2 equivalents of 5′-GMP (72 h, 310 K, pH 4.5 unbuffered), electrospray ionization mass spectrometric analysis revealed the presence of a range of 2 : 1 and 1 : 1 5′-GMP–metal complex adducts. Interestingly, for the complexes with the closed conformation, 5b and 6b, and the flexible ethylene diamine linker, 3b, these adducts include examples where all four labile chlorido ligands are lost and replaced with a single 5′-GMP molecule – indicating both ruthenium centres are bridged via coordination to the same 5′-GMP ligand. ¹H and ³¹P NMR spectroscopy indicated the most likely coordination sites are N7 and phosphate oxygen (see below). For the more open complex, 4b, the conformation of which is expected to be less favourable for intramolecular bridging coordination to occur, the corresponding peaks in the mass spectrum are of a relatively low intensity, the peaks corresponding to 2 : 1 5′-GMP–metal complex adducts are of higher relative intensity (Fig. S17†). Studies with the monomer analogue 2b reveal the formation of only 1 : 1 5′-GMP–metal complex adducts, even in the presence of 2 equivalents 5′-GMP. For incubations of dinuclear compounds 3b–5b with 2 equivalents of 5′-GMP at pH 7.5 (310 K, 60 mM ammonium acetate) only 1 : 1 adducts were detected. Adducts include examples where all four chlorido ligands are replaced by a single 5′-GMP ligand, alongside similar adducts also containing a coordinated acetate ligand. It is likely that at pH 7.5 the deprotonation of the 5′-GMP phosphate (pK_a of 6.49 reported for PO₃H of 5′-GMP³³) facilitates the formation of intramolecularly bridged 1 : 1 adducts in preference to the formation of 2 : 1 5′-GMP–metal complex adducts. Complementary binding studies between 5′-GMP and 3b were also monitored by ¹H and ³¹P NMR spectroscopy over 24 h (pD = 7.5, 200 mM HEPES buffer, 310 K) (Fig. S18†). Spectra are complex due to the formation of diastereoisomeric adducts upon coordination with 5′-GMP. Multiple new 5′-GMP H8 resonances in the region 8.00–8.85 ppm were identified by ¹H NMR spectroscopy corresponding to adducts involving coordination of N7 of 5′-GMP to the ruthenium ion. Concurrent analysis with ³¹P NMR spectroscopy revealed the appearance of new 5′-GMP resonances (−2.90 to 8.45 ppm) indicative of phosphate coordination to the ruthenium ion alongside N7 coordination. This data complements the observation of intramolecular crosslinking of these dinuclear RAPTA analogues by 5′-GMP as observed in the mass spectrometric studies. A further distinctive proton resonance set, centred at 7.05 ppm, also gradually appeared in the aromatic region of the ¹H NMR spectra. This was deduced to be due to formation of the free arene ligand, indicating that upon coordination of 5′-GMP to 3b, potentially through both phosphate and N7 coordination, the arene ligand is lost from the adduct. In addition, the colour of the solution changed from yellow to black over the 24 h incubation period indicative of complex decomposition. Similar 5′-GMP–Ru adduct(s) were not identified during the mass spectrometric studies although the free arene ligand was detected by ESI-MS in the incubations performed at pH 7.5. The loss of the arene ligand is not unprecedented with similar loss of the arene ligand being reported previously in binding studies of RAPTA-C with a 14-mer oligonucleotide.³⁴

Binding experiments were also performed with 2 equivalents of L-histidine at pH 7.1 and monitored by ¹H and ³¹P NMR spectroscopy and ESI-MS. As with 5′-GMP, ESI-MS revealed the formation of a range of 1 : 1 adducts between L-histidine and dinuclear compounds 3b–6b. Bridging of L-histidine between the two ruthenium ions is observed for all the dinuclear compounds; characterised by peaks corresponding to the loss of all four chlorido ligands in exchange for a single histidine, although the intensity of these peaks are relatively low compared to the intensity of other 1 : 1 adduct peaks (Fig. S19†). Peaks corresponding to 2 : 1 L-histidine–metal complex adducts are also observed in all the spectra of the dinuclear compounds, but are of low relative intensity (<15%). For the mononuclear analogue 2b, in the presence of 2 equivalents of L-histidine, only peaks corresponding to 1 : 1 adducts are observed. ¹H NMR spectroscopy indicates that loss of the arene does not take place during the 24 h incubation, and, unlike the 5′-GMP incubation, the colour of the solution remains yellow.

As these small-molecule binding experiments confirm all dinuclear complexes 3b–6b readily coordinate to appropriate donor ligands simultaneously through both ruthenium atoms, binding studies were expanded to include a short model peptide sequence and a single-stranded 13-mer oligonucleotide.

Oligonucleotide and peptide binding studies

Oligonucleotide binding studies were performed on the 13-mer sequence 5′-ATACATCGTACAT-3′ in unbuffered aqueous solutions (72 h, 310 K, pH = 4.5, 0.2 mM complex and 2 eq. oligonucleotide). The reaction mixture was then diluted and directly analysed using ESI-MS in negative ionization mode. In higher mass regions of the spectra peaks attributable to 1 : 1 oligonucleotide–metal complex adducts are observed in which the metal complexes have lost their chlorido ligands. No higher order adducts were detected. However, the predominant species in these spectra correspond to oligonucleotide–metal complex adducts where the arene ligand is lost leaving the Ru–PTA fragment coordinated to the oligonucleotide. These results correlate with the NMR binding studies of 5′-GMP with 3b where loss of the arene ligand was observed.

Peptide binding studies utilised a fragment of amyloid β-protein (residues 1–16, H-Asp-Ala-Glu-Phe-Arg-His-Asp-Ser-Gly-Tyr-Glu-Val-His-His-Gln-Lys-OH). This 16-mer contains three histidine residues as well as one lysine and two glutamate residues – all of which have been observed to coordinate to RAPTA compounds in crystallographic studies.²⁰ Incubations were performed in unbuffered aqueous solutions (72 h, 310 K) in a 1 : 1 peptide–metal complex ratio. Using ESI-MS in all cases a range of 1 : 1 adducts were observed with no 2 : 1 or higher order complex–peptide or peptide–complex adducts detected. As with the small-molecule binding studies, 1 : 1 dinuclear metal complex–peptide adducts are detected where all four labile chlorido ligands are lost (and, in some cases, loss of PTA ligands is also observed) and substituted by a single peptide. The loss of two or three ligands at each Ru centre implies that each metal of the dinuclear compounds must be coordinated to one or more amino acid residues of the peptide. It is likely that crosslinked species where each metal centre is bound to a different amino acid residue form a proportion of these 1 : 1 adducts. ESI mass spectra of peptide adducts of 4b, 5b and 6b reveals a similar peak distribution (Fig. S20†). To probe the nature of the binding within these 1 : 1 adducts electron-transfer dissociation (ETD) fragmentation studies were performed on selected ions.

Electron-transfer dissociation (ETD) fragmentation and ion mobility-mass spectrometry (IM-MS) studies

ETD fragmentation of peptides is an established technique used to randomly fragment the N–C_α bonds of a peptide backbone. This technique often preserves post-translational modifications of the peptide side chains, such as phosphorylation and glycosylation, allowing their identification and localisation in a peptide sequence to be determined.³⁵ More recently, this technique has been used to identify drug metallation sites in peptide³⁶ and protein samples.³⁷ We used ETD to probe the binding of 2b–6b to the 16-mer peptide H-Asp-Ala-Glu-Phe-Arg-His-Asp-Ser-Gly-Tyr-Glu-Val-His-His-Gln-Lys-OH (see ESI† for a description of data analysis, see Table 1 for adducts analysed and location of metallated residues). Analysis of the ETD spectra of the 1 : 1 [peptide + 2b + 2H − 2Cl]⁴⁺ and [peptide + 2b + 2H − 2Cl − PTA]⁴⁺ adducts revealed complex binding at the His⁶, His¹³ and His¹⁴ sites. These observations could correspond to crosslinking of sites His⁶, His¹³ or His¹⁴ through metallation by a single Ru complex where crosslinking then breaks apart upon ETD fragmentation, or could correspond to a population of three metallated peptides within the sample, where the site of modification is solely at His⁶, His¹³ or His¹⁴.

Table 1 Peptide–complex adducts subjected to electron-transfer dissociation fragmentation and identified sites of metallation

Adduct	Identified site of metallation
[Peptide + 2b + 2H − 2Cl]^4+	His^6, His^13 and His^14
[Peptide + 2b + 2H − 2Cl − PTA]^4+	His^6, His^13 and His^14
[Peptide + 3b + H − 4Cl]^5+	His^6 and His^13
[Peptide + 3b − PTA − 4Cl]^4+	His^6, other sites unidentified
[Peptide + 3b − 4Cl]^4+	His^6, other sites unidentified
[Peptide + X + H − 4Cl − PTA]^5+ (X = 4b, 5b or 6b)	His^6 and His^13 or His^14
[Peptide + X + H − 4Cl]^5+ and [peptide + X − 4Cl − PTA]^4+ (X = 4b, 5b or 6b)	Central amino acid residues (no metallation observed at terminal residues)

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For dinuclear complexes 3b–6b, ETD spectra provided direct evidence that each of the complexes was able to crosslink the peptide through simultaneous coordination of each ruthenium centre to one or more different histidine residues at His⁶, His¹³ and His¹⁴ (Fig. 4). Further peptide adducts of each metal complex were analysed where either a single metallated histidine site was identified or no metallated amino acid sites were clearly identified (although metallation at the terminal amino acid residues was discounted). These spectra provide only a partial picture of the sites of metallation in these adducts but, given the 1 : 1 peptide–complex stoichiometries of the adducts, they also provide evidence towards crosslinking of the peptide through the histidine residues. It was noticeable that ETD spectra with 4b–6b are very similar for each particular adduct, exhibiting virtually identical fragment distributions in each case (Fig. S21 and S22†). These results suggest that although 4b and 5b/6b possess different conformations there is sufficient flexibility in these peptide adducts to allow crosslinking through the same histidine positions. To probe these adducts further complementary binding studies were performed using ion mobility-mass spectrometry (IM-MS). IM-MS has been recently used to probe conformational changes in the protein ubiquitin on platination with cisplatin³⁸ and also in combination with fragmentation techniques to identify drug metallation sites on peptides.³⁶

Fig. 4 ETD LTQ Orbitrap FTMS of the [H-Asp-Ala-Glu-Phe-Arg-His-Asp-Ser-Gly-Tyr-Glu-Val-His-His-Gln-Lys-OH + 3b + H − 4Cl]⁵⁺ adduct (top) and the [H-Asp-Ala-Glu-Phe-Arg-His-Asp-Ser-Gly-Tyr-Glu-Val-His-His-Gln-Lys-OH + 5b + H − 4Cl − PTA]⁵⁺ adduct (bottom) after a 100 ms interaction period with fluoroanthene radical anions. Fragments corresponding to metallation at His⁶ are labelled in black whereas fragments labelled in red are those corresponding to metallation at His¹³. Fragments labelled with * correspond to a metallated fragment.

We used IM-MS to probe the different complex–peptide adducts analysed by ETD fragmentation and comparisons of their arrival time distributions (ATDs) have been made (Table 2, Fig. 5, 6 and S23–S26†). For the mononuclear complex-peptide adduct formed through loss of all chlorido and a PTA ligand [peptide + 2b + 2H − 2Cl − PTA]⁴⁺ a single peak was observed in the ATD indicating this adduct exists as a single isomer in the gas phase. For the equivalent adducts of the dinuclear complexes ([peptide + X − 4Cl − PTA]⁴⁺ (X = 3b–6b)) the ATDs are more complex and exhibit two peaks (Fig. 5) indicating these adducts all exist as two isomeric species in the gas phase. For 1 : 1 peptide–complex adducts, formed through loss of all chlorido ligands from Ru whilst retaining all their PTA ligands ([peptide + X + nH − nCl]⁴⁺ (X = 2b–6b)), the ATDs in each case consist of two peaks. These results show each adduct in this series also exists as two isomeric species in the gas phase. A comparison of the drift times at which each peak in the ATDs is centred revealed that peaks for the [peptide + X + nH − nCl]⁴⁺ adducts are all centred at equal or longer drift times than their corresponding [peptide + X + nH − nCl − PTA]⁴⁺ adducts (X = 2b–6b). This reflects the larger size of the former adducts due to their retention of the PTA ligand, and highlights the important role played by the ligand set around the Ru ion in determining the shape of the adduct formed. The split distributions observed in the ATDs of all the dinuclear Ru compound–peptide adducts are likely due to crosslinking of the peptide by the dinuclear compounds between His⁶–His¹³ and His⁶–His¹⁴ to yield two adducts of different size. In contrast, the single peak observed in the ATD of the adduct [peptide + 2b + 2H − 2Cl − PTA]⁴⁺ may be due to the metallation at the three histidine sites, as observed in the ETD analysis, yielding three adducts of identical size. An alternative interpretation is that the peptide is crosslinked by a single metal complex through sites His⁶, His¹³ and His¹⁴ to yield a single adduct; this interpretation correlates with the metal fragment of this adduct having lost two chlorido and one PTA ligand. Similar to the ETD analysis of the peptide adducts of the isomeric complexes 4b–6b no differences are observed in the ATDs of their peptide adducts, or between the ATDs of the [X − 4Cl + 2OH + 2H₂O]²⁺ (X = 4b–6b, Fig. S25†) ions themselves. Combined, these data indicate that despite the different conformations of the three complexes, as confirmed by crystallographic and ¹H NMR analysis, they have an identical size, at least in the gas phase. In addition, these data show that adducts formed between the peptide and the metal complexes are similar in terms of their size and in terms of their preferential binding sites.

Table 2 Data from IM-MS studies showing ion mobility drift times of peak maxima, measured monoisotopic masses of selected peptide adducts or complex ions and the corresponding theoretical values

Peptide adduct/ion	Ion mobility drift time(s) (ms)	Measured monoisotopic mass (m/z)	Theoretical monoisotopic mass (m/z)
[Peptide + 2b + 2H − 2Cl − PTA]^4+ (C97H140N28O29Ru)	2.93	565.77	565.73
[Peptide + 3b − 4Cl − PTA]^4+ (C112H159N32O30PRu2)	2.88, 3.15	666.78	666.74
[Peptide + 4b − 4Cl − PTA]^4+ (C124H167N32O30PRu2)	2.98, 3.42	704.80	704.76
[Peptide + 5b − 4Cl − PTA]^4+ (C124H167N32O30PRu2)	2.98, 3.42	704.80	704.76
[Peptide + 6b − 4Cl − PTA]^4+ (C124H167N32O30PRu2)	2.98, 3.42	704.80	704.76
[Peptide + 2b + 2H − 2Cl]^4+ (C103H152N31O29PRu)	2.98, 3.09	605.04	605.00
[Peptide + 3b − 4Cl]^4+ (C118H171N35O30P2Ru2)	2.93, 3.42	706.06	706.01
[Peptide + 4b − 4Cl]^4+ (C130H179N35O30P2Ru2)	3.04, 3.53	744.07	744.03
[Peptide + 5b − 4Cl]^4+ (C130H179N35O30P2Ru2)	2.98, 3.53	744.08	744.03
[Peptide + 6b − 4Cl]^4+ (C130H179N35O30P2Ru2)	2.98, 3.58	744.07	744.03
[4b + 2OH + 2H2O − 4Cl]^2+ (C46H66N8O6P2Ru2)	2.66, 2.88	546.12	546.13
[5b + 2OH + 2H2O − 4Cl]^2+ (C46H66N8O6P2Ru2)	2.66, 2.88	546.12	546.13
[6b + 2OH + 2H2O − 4Cl]^2+ (C46H66N8O6P2Ru2)	2.66, 2.88	546.12	546.13
[Peptide + 4H]^4+ (C84H123N27O28)	2.66	489.51	489.48

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Fig. 5 Comparison of the ATDs of the [peptide + 2b − 2Cl − PTA + 2H]⁴⁺ (bottom), [peptide + 3b − 4Cl − PTA]⁴⁺ (middle) and [peptide + 4b − 4Cl − PTA]⁴⁺ (top) adducts. Inset shows the isotope profile of the corresponding mass peaks in the mass spectrum.

Fig. 6 Comparison of the ATDs of the [peptide + 2b − 2Cl + 2H]⁴⁺ (bottom), [peptide + 3b − 4Cl]⁴⁺ (middle) and [peptide + 4b − 4Cl]⁴⁺ (top) adducts. Inset shows the isotope profile of the corresponding mass peaks in the mass spectrum.

Evaluation of in vitro anticancer activity

The cytotoxicity of 2a–6a and 2b–6b was assessed in human ovarian carcinoma (A2780), human ovarian carcinoma with acquired resistance to cisplatin (A2780cisR) and human embryonic kidney (HEK 293) cell lines using the MTT assay (Table 3). As discussed earlier, the cytotoxicity of the original RAPTA series toward a range of cancer cell lines is low (IC₅₀ often >300 μM) and these high IC₅₀ values are mirrored by the values determined for mononuclear compounds 2a and 2b, which are >300 μM in the three cell lines. All dinuclear compounds 3a–6a and 3b–6b are significantly more cytotoxic against the A2780 cell line than the mononuclear analogues. Of these, the most cytotoxic compounds are those with the closed conformation, i.e.5a, 6a, 5b and 6b, with the chlorido-analogues (5b and 6b) being slightly more cytotoxic than the oxalato-analogues (5a and 6a). Interestingly, the diastereoisomeric complexes 4a and 4b are significantly less active, with IC₅₀ values increasing over 5-fold in the case of the chlorido-analogue and over 2-fold for the oxalato-analogue. The flexible dinuclear oxalato-complex, 3a, has a comparable cytotoxicity to that of 4a whereas the IC₅₀ value of the chlorido analogue, 3b, is significantly greater than that of 4b.

Table 3 In vitro anticancer activity of compounds 2a–6b in human ovarian carcinoma (A2780), human ovarian carcinoma cisplatin resistant (A2780cisR) and human embryonic kidney 293 (HEK-293) cell lines after 72 h exposure, octanol–water partition coefficients for compounds 2a–6a and cellular (A2780) uptake of ruthenium after exposure to 2a–6a for 5 h exposure at 300 μM

Compound	IC50 (μM)				Cellular uptake (pmol ruthenium per 10^6 cells)
	A2780	A2780cisR	HEK-293	log P
2a	>400	>400	>400	−1.57	459 ± 89
3a	23 ± 0.5	88 ± 10	29 ± 2.5	−1.26	474 ± 21
4a	25 ± 2.5	74 ± 6	19 ± 4	−1.38	391 ± 52
5a	10 ± 0.5	10 ± 1	6 ± 1	−1.41	332 ± 25
6a	8.5 ± 0.5	15 ± 0.5	6 ± 0.5	−1.36	901 ± 97
2b	>300	>300	>300	—	—
3b	64 ± 9.5	>300	35 ± 3	—	—
4b	20 ± 0.5	95 ± 6.5	24 ± 3.5	—	—
5b	3.7 ± 0.6	4.5 ± 0.2	8 ± 0.5	—	—
6b	3.7 ± 0.6	7.0 ± 0.5	5.9 ± 0.2	—	—
Cisplatin	1.5 ± 0.2	14.5 ± 2	7.5 ± 1	—	—

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In the A2780cisR cell line the IC₅₀ values of complexes 5a, 6a, 5b and 6b remain low, albeit, for complexes 6a and 6b, with the linker of (R,R)-configuration, with a slight increase in IC₅₀ values relative to those values obtained for the A2780 cell line. The IC₅₀ values of complexes 5a and 5b, with the linker of (S,S)-configuration, remain essentially unchanged. In contrast, the IC₅₀ values for 3a and 3b (linked by the flexible ethylenediamine moiety) and 4a and 4b (with the linker of (R,S)-configuration) increase dramatically. For example, 4a and 4b possess IC₅₀ values 7 and 21 times, respectively, greater than those of their most active diastereoisomers (5a and 5b) whereas 3a is 9 fold less cytotoxic than the most active oxalato-compound 5a, and 3b is essentially inactive in the A2780cisR cell line (IC₅₀ > 300 μM). In addition, compounds 5b and 6b are significantly more active than cisplatin in this cell line (over 3 fold in the case of 5b).

In the HEK-293 cell line, used as a model for non-tumorigenic cells, the activity of the compounds follows the pattern of activity of the compounds observed in the A2780 cell line. The cytotoxicity of 5a, 6a, 5b and 6b towards this cell line remains higher than that of 3a, 3b, 4a and 4b with no significant differences in IC₅₀ values between chlorido- and oxalato-analogues observed for each case. One exception is that in this cell line the activity of the chlorido complex 3b is greater than that observed in the A2780 and A2780cisR cell lines and comparable to the value of its oxolato analogue 3a, which in other cell lines is more active.

The differences in activity between diastereoisomeric complexes 4a–6a and 4b–6b are significant, with an apparent conformational dependence observed for both cytotoxic activity and also for susceptibility to resistance in the A2780cisR cell line. The complexes with the “closed” conformation consistently show high cytotoxicity against all cell lines examined, and in the case of 5a and 5b, with the linker of (S,S)-configuration, virtually no loss of activity against the A2780cisR cell line. In contrast, the compounds with the more open structure (4a and 4b incorporating the linker of (R,S)-configuration), or those with a flexible ethylenediamine linker that may also be expected to occupy an open conformation (3a and 3b), are significantly less cytotoxic and also show a dramatic decrease in cytotoxicity against the A2780cisR cell line. To probe whether the origin of these observations were due to the differential intracellular uptake of these complexes, A2780 cells were incubated for 5 h with 2a–6a (300 μM) and the intracellular ruthenium content was determined using ICP-MS. It was found that the level of internalised 4a (391 ± 52 pmol Ru per 10⁶ cells) is comparable to that observed for 5a (332 ± 25 pmol Ru per 10⁶ cells). Unexpectedly, under identical incubation conditions the level of 6a (901 ± 97 pmol Ru per 10⁶ cells) is significantly higher than that observed for 5a despite it having a very similar cytotoxicity profile. At present, the origins of this differential uptake are unknown, though a recent report has shown the chirality of substituents on the ruthenium arene ligand can have a significant influence on biological activity.³⁹ Compound 3a exhibits slightly higher cellular uptake levels (474 ± 21 pmol Ru per 10⁶ cells) than 4a and 5a whereas the mononuclear analogue 2a (459 ± 89 pmol Ru per 10⁶ cells) displays the highest level of internalised complex of all the compounds. These results indicate that while the levels of ruthenium cellular uptake do not correlate with the cytotoxicity profiles of the compounds at 72 h, all compounds, under the experimental conditions, readily internalise in A2780 cells and at comparable levels observed for RAPTA-T.¹⁹ In addition to these uptake studies the log P values for complexes 2a, 3a, 4a, 5a and 6a were determined using the shake-flask method.⁴⁰ These experiments reveal the hydrophilic nature of all complexes with little variation in log P within the series. Therefore it is reasonable to conclude that the differences in the cytotoxicity, at least in the A2780 cell line, must be due to cellular events associated with the complexes and their ability to crosslink target biomolecules as demonstrated in binding studies, and is not due to significant differences in complex hydrophilicity and cellular uptake.

Conclusions

We have developed a new route to arene-functionalised organometallic Ru(II) compounds, allowing access to flexible and conformationally rigid dinuclear organometallic Ru(II) complexes. Crystallographic studies and NMR spectroscopy revealed that the stereochemical configuration of the 1,2-diphenylethylenediamine (DPEN) linker controls whether the dinuclear complexes have an open or closed conformation. The range of adducts formed between these rigid dinuclear complexes, 4b–6b and the small molecules 5′-GMP and L-histidine appears to be governed by the conformation of the complex, and all dinuclear complexes 3b–6b are capable of crosslinking model peptide/oligonucleotide sequences. The mononuclear complexes, as observed for the original RAPTA series, are non-cytotoxic whereas dinuclear complexes are significantly more cytotoxic. Although the different conformations of 4bvs.5b/6b resulted in no differences in type and size of adducts formed in model peptide binding studies, an apparent conformational dependence on the cytotoxicity of these dinuclear complexes was observed. Those with the more closed conformation (5a–6b) are significantly more cytotoxic than those with the more open conformation (4a and 4b) and are unaffected by resistance mechanisms operating in the A2780cisR cell line. In contrast, 4a and 4b were significantly less cytotoxic toward this cell line than the A2780 cell line. However, for all dinuclear complexes, cytotoxicity toward the cancerous A2780 cell line and the non-cancerous HEK-293 cell line varied little. Further studies revealed the hydrophilicity of the compounds, as assessed by measurement of log P values, was similar within the series 2a–6a, and uptake experiments with A2780 cells revealed an appreciable level of complex association in each case. Combined, these observations lead to the conclusions that the different cytotoxicities of the mononuclear and dinuclear compounds is linked to the ability of the complex to crosslink biomolecular targets and that the cytotoxicity of the dinuclear compounds is significantly linked to their conformation.

Acknowledgements

This work was supported by a Marie Curie Intra-European Fellowship within the 7^th European Community Framework Programme (Project 273658-DINURU to B.S.M). We thank the Nestle Institute of Health Sciences for access to the Synapt G2-S quadrupole-time-of-flight HDMS mass spectrometer and Julien Bourquin for assistance in data collection and analysis. We thank Dr Luc Patiny for developing calculation tools for interpretation of ETD fragmentation spectra. We thank Baihua Ye for chiral HPLC analysis and Prof. Nicolai Cramer for access to these facilities.

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Footnotes

† Electronic supplementary information (ESI) available: Experimental procedures for all novel compounds and copies of nuclear magnetic resonance spectra, X-ray diffraction parameters, HPLC chromatograms for arene ligands of 5a and 6a, UV-vis spectra and selected mass spectra. CCDC 978401–978404. For ESI and crystallographic data in CIF or other electronic format see DOI: 10.1039/c4sc00116h

‡ Structural data obtained from single crystals of samples of 5a and 6a is identical, with each crystal found to contain a 50 : 50 mixture of 5a and 6a resulting in the solved structures exhibiting disorder in the region of the chiral centres of the DPEN linker molecule. As the (1S,2S)-DPEN and (1R,2R)-DPEN linkers utilised in the synthesis of 5a and 6a respectively were commercial samples with a stated enantiomeric excess of 98% and 99% respectively, we postulate the racemic composition of crystals of 5a and 6a are not reflective of the bulk sample, but are most likely a result of the co-crystallization of the desired and undesired enantiomers present in the samples. Chiral HPLC was used to determine the percentage of each enantiomer in bulk samples of 5a and 6a by analysis of the arene ligand following cleavage from the Ru ions (see Experimental section and Fig. S2–S4† for details). This analysis showed that 5a was obtained with an enantiomeric excess of 99.4% and 6a was obtained with an enantiomeric excess of 99.8%. These results confirm that the bulk samples of 5a and 6a are of a high level of enantiomeric purity and indicate that co-crystallization of the enantiomeric impurity in each sample with the bulk product is the source of the racemic crystals obtained.

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