Synthesis, structural characteristics, DNA binding properties and cytotoxicity studies of a series of Ru(III) complexes

Journal of Inorganic Biochemistry 102 (2008) 1644–1653

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Synthesis, structural characteristics, DNA binding properties and cytotoxicity studies of a series of Ru(III) complexes Caiping Tan, Jie Liu *, Lanmei Chen, Shuo Shi, Liangnian Ji * MOE Laboratory of Bioinorganic and Synthetic Chemistry, School of Chemistry and Chemical Engineering, Sun Yat-Sen University, Xingang Road, Guangzhou 510275, PR China

a r t i c l e

i n f o

Article history: Received 12 November 2007 Received in revised form 18 March 2008 Accepted 20 March 2008 Available online 1 April 2008 Keywords: Ru(III) complexes Anticancer drug DNA binding

a b s t r a c t Four related ruthenium(III) complexes, with the formula mer-[RuCl3(dmso)(NN)] (dmso = dimethyl sulfoxide; NN = 2,20 -bipyridine (1), 1,10-phenantroline (2), dipyrido[3,2-f:20 ,30 -h]quinoxaline (3) and dipyrido[3,2-a:20 ,30 -c]phenazine (4)), have been reported. Complexes 3 and 4 are newly synthesized and characterized by X-ray diffraction. The hydrolysis process of 1–4 has been studied by UV–vis measurement, and it has been found that the extension of the NN ligands can increase the stability of the complexes. The binding of these complexes with DNA has been investigated by plasmid cleavage assay, competitive binding with ethidium bromide (EB), DNA melting experiments and viscosity measurements. The DNA binding affinity is increased with the extension of the planar area of the NN ligands, and complex 4 shows an intercalative mode of interaction with DNA. The in vitro anticancer activities of these compounds are moderate on the five human cancer cell lines screened. Ó 2008 Elsevier Inc. All rights reserved.

1. Introduction Since the introduction of cisplatin (cis-[PtCl2(NH3)2]) into the clinic in 1979, intensive efforts have been channeled toward the search for cytotoxic compounds with more acceptable toxicity profiles but retentive or even expansive activity. The design of new platinum antitumor drugs has mainly concentrated on direct cisplatin analogues. More recently, abnormal structures that violate the empirical structure–activity relationships of platinum compounds, such as multinuclear complexes and transplatin derivatives, are developed to discover ‘‘non-classical” drugs that can act in a way different from cisplatin [1–4]. There have also been efforts to investigate drugs based on other transition metals [5,6]. Among the different metal complexes generating interests, ruthenium complexes have shown great potential and remain the subject of extensive drug discovery efforts [7,8]. [H2im][trans-RuCl4(dmsoS)(Him)] (NAMI-A; Him = imidazole, dmso = dimethyl sulfoxide) has entered clinical trials in 1999 due to its very good antimetastatic activity (Fig. 1) [9,10]. Two other complexes with similar structures, namely [H2im][trans-RuCl4(Him)2] (ICR; Him = imidazole) and [H2ind][trans-RuCl4(Hind)2] (KP1019; Hind = indazole), show excellent antitumor activity in various animal models (Fig. 1). KP1019 is introduced into phase I clinical trials against colon carcinomas and their metastases in 2003 [11–13]. Recently, several other ruthenium complexes have also shown promise as

* Corresponding authors. Tel.: +86 20 31745729; fax: +86 20 84035497 (J. Liu); tel./fax: +86 20 84035497 (L. Ji). E-mail addresses: [email protected] (J. Liu), [email protected] (L. Ji). 0162-0134/$ - see front matter Ó 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.jinorgbio.2008.03.005

anticancer agents, e.g. a-[Ru(II)(azpy)2Cl2] (azpy = 2-phenylazopyrine) [14,15], mer-[Ru(III)(terpy)Cl3] (terpy = 2,20 :60 200 -terpyridine) [16], [Ru(IV)(cdta)] (cdta = 1,2-cyclohexanediminetetraacetate) [17], and Ru(II) organometallic complexes of the type [(g6-arene)Ru(II)(en)Cl][PF6] (en = ethylenediamine) [18]. Although the mechanisms of action of both KP1019 and NAMI-A have not yet been fully understood, aquation is widely accepted as an important step for the function of such species. They appear to be precursors that hydrolyze rapidly in vivo forming a number of potentially active species, which can facilitate their binding with biomolecules, and the rate of this process can greatly influence their antitumor activities [19–23]. Besides aquation, the in vivo reduction of Ru(III) to the more reactive Ru(II) [24–26] and the ability of ruthenium to accumulate specifically in cancer tissues, possibly via the transferrin pathway [6], are also thought to be important for their modes of action, especially for KP1019 species. DNA is thought to be the primary target for platinum-based antitumor compounds [27], while the targets of ruthenium antitumor complexes have not yet been explicitly established. Though both NAMI-A and KP1019 can coordinate irreversibly to DNA [28], other targets such as plasma proteins and glutathione are also thought to be more important than DNA for their antitumor activities [29,30]. DNA binding seems to play a more important role for Ru(II)–arene organometallic complexes, which can interact with DNA by direct coordination to the bases, intercalation and stereospecific H-bonding [31]. Hydrophobic interactions have shown to be important in the biological activities of some platinum anticancer agents. Planar aromatic ligands such as acridine orange, 9-aminoacridine, and ethidium bromide have been incorporated into platinum com-

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2. Experimental

O NH

H N

N Cl

HN

Cl

Ru

Cl

S Ru

Cl

Me Cl

HN

NH

Cl N

Cl

Cl

2.1. Materials

Me

N NH

HN

Fig. 1. Structures of [H2ind][trans-RuCl4(Hind)2] (KP1019; left) and [H2im][transRuCl4(dmso-S)(Him)] (NAMI-A; right).

plexes as potential intercalators [32–34]. The antitumor activity of trans-[PtCl2(NH3)L] (L = planar aromatic N donor such as pyridine, thiazole, or quinoline) is much higher than that of transplatin, which has been attributed to the additional intercalation or hydrophobic interaction after covalent binding of Pt(II) to nucleobases [35,36]. It is also well known that the presence of intercalative ligands in coordinatively saturated octahedral Ru(II), Os(II), and Rh(III) complexes can increase their DNA binding affinity, and give rise to highly specific recognition of DNA base sequences via shape selection [37]. Since the discovery of antitumor and antimetastatic properties of KP1019 and NAMI-A, many complexes with similar structures have been synthesized [10,38,39], but halo-Ru(III) complexes with bidentate ligands screened for antitumor activity are still scarce. Recently, we have reported the differences in biological properties between two Ru(III) complexes, namely mer-[RuCl3(CH3CN)(dpq)] and mer-[RuCl3(dmso)(dpq)] (dpq = dipyrido[3,2-f:20 ,30 -h]quinoxaline) [40]. The present study reports the synthesis, characterization and biological properties of a series of Ru(III) complexes containing planar bidentate N–N ligands (Fig. 2; bpy = 2,20 -bipyridine, phen = 1,10-phenantroline, dppz = dipyrido[3,2-a:20 ,30 -c]phenazine). These complexes have both the potential leaving groups and the possible DNA intercalating ligands, so they may interact with DNA in a bifunctional mode including covalent binding to the nucleobases and non-covalent intercalation. The stability under physiological conditions and the DNA binding properties of these complexes have been studied in detail. A preliminary cytotoxic study shows that the cytotoxicities of these complexes are moderate against the five human cancer cell lines evaluated. The relationship between the chemical property and the biological activity of these complexes has also been analyzed to investigate the possible antitumor mechanism.

Me O

Me S

Cl

Ru

N

N

Cl

N Cl

N

bpy

N

N

N

N

N

N

N

N

dpq

N N phen 1: N-N = bpy 2: N-N = phen 3: N-N = dpq 4: N-N = dppz

dppz Fig. 2. Sketch of the structures of the complexes.

All materials and solvents were purchased commercially and used without further purification unless otherwise noted. Ligands dpq, dppz [41], complexes Na[trans-RuCl4(dmso-S)(Him)] (NAMI) [9], cis-[RuCl2(dmso)4] [42], [Ru(phen)2(dppz)](PF6)2 [43], mer[RuCl3(dmso)(phen)] [44], [H(dmso)2][trans-RuCl4(dmso)2] [45] and mer-[RuCl3(dmso)(bpy)] [46] were prepared according to the literature procedures. Calf thymus DNA (CT-DNA) was obtained from the Sigma Company. MTT (3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide) was purchased from the Sigma Company. RPMI-1640 medium was purchased from Invitrogen Co. (USA) and DMEM medium could be got from Life Technologies. Doubly distilled water was used to prepare buffers. A solution of CT-DNA gave ratios of UV–visible (UV–vis) absorbance at 260 and 280 nm of about 1.9:1, indicating that the DNA was sufficiently free of protein [47]. The concentration of DNA was determined by UV–vis absorbance using the molar absorption coefficient (6600 M1 cm1) at 260 nm [48]. The plasmid DNA, pBR322 (as a solution in Tris buffer), with a length of 4361 base pairs was purchased from Biopolymers Inc., and kept at 20 °C and used directly without further purification. Caution: Metal perchlorates are potentially explosive and should thus be handled with care. 2.2. General methods Microanalysis (C, H, and N) was carried out using a Perkin-Elmer 240Q elemental analyzer. Electrospray mass spectra (ESIMS) were recorded on a LCQ system (Finnigan MAT, USA). The spray voltage, tube lens offset, capillary voltage and capillary temperature were set at 4.50 kV, 55.00 V, 18.00 V and 275 °C, respectively, and the quoted m/z are for the major peaks in the isotope distribution. UV–vis spectra were recorded on a Varian Cary 300 spectrophotometer. Emission spectra were recorded on a Shimadzu RF-5301 PC spectrofluorophotometer at room temperature. The IR spectra were recorded in the 400–4000 cm1 region using KBr pellets and a Bruker EQUINOX 55 spectrometer. The buffers used were as follows: Buffer A: 50 mM NaH2PO4, 100 mM NaCl, pH 7.4; Buffer B: 0.01–0.20 M NaClO4, 1 mM Tris– HCl, 0.1 mM Na2EDTA (EDTA = ethylenediaminetetraacetic acid), pH 7.4; Buffer C: 5 mM Tris–HCl, 50 mM NaCl, pH 7.1; Buffer D: 50 mM Tris–HCl, 18 mM NaCl, pH 7.2; Buffer E (TBE): 89 mM Tris, 83 mM H3BO3, 2 mM EDTA, pH 8.3. 2.3. Synthesis of the complexes 2.3.1. mer-[RuCl3(dmso)(bpy)] (1) Complex 1 was synthesized by treating [H(dmso)2][transRuCl4(dmso)2] with 2,20 -bipyridine in equimolar amounts in ethanol at room temperature following the literature method [46]. Yield: 55%. ESI-MS (m/z): 442.6 for [RuCl3(dmso)(bpy)]. Elemental anal. Calcd. for C12H14Cl3N2ORuS: C, 32.63; H, 3.19; N, 6.34. Found: C, 32.62; H, 3.18; N, 6.35. IR data (KBr, cm1): 3106 (m, C–H), 3074 (m, C–H), 1604–1447 (d, C–H), 1312–1308 (d, C–H), 1107 (m, S–O), 1071 (m, S–O). 2.3.2. mer-[RuCl3(dmso)(phen)] (2) Complex 2 was synthesized following the literature method [44] with slight modifications. A bright yellow mixture contaning cis-[RuCl2(dmso)4] (0.20 g, 0.41 mmol), phen (74 mg, 0.41 mmol), 6 M HCl (10 ml), and CH3CH2OH (10 ml) was heated at 80 °C for 2 h, after which the

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resulting clear red solution was concentrated in a rotary evaporator and then dried in vacuo. The crude product was purified by column chromatography on a neutral alumina with dried CH2Cl2 and CH3OH (10:1, v/v) as eluent, which gave 3 as a dark red solid. Yield: 134 mg (70%). The structure is the same as that reported in the literature [44]. ESI-MS (m/z): 466.7 for [RuCl3(dmso)(phen)]. Elemental anal. Calcd. for C14H14Cl3N2ORuS: C, 36.10; H, 3.03; N, 6.01. Found: C, 36.07; H, 3.05; N, 6.03. 2.3.3. mer-[RuCl3(dmso)(dpq)] (3) The complex was obtained by using dpq (96 mg, 0.41 mmol) instead of phen in 2. The crude product was purified by column chromatography on a neutral alumina with dried CH2Cl2 and CH3OH (10:1, v/v) as eluent, which gave 3 as a dark red solid. Yield: 0.160 g (72%). Brown prism-shaped crystals were obtained by slow evaporation of the concentrated CH3OH or CH3CH2OH solution of 3 at room temperature. Yield: 0.114 g (50%). ESI-MS (m/z): 518.9 for [RuCl3(dmso)(dpq)]. Elemental anal. Calcd. for C16H14Cl3N2ORuS: C, 37.11; H, 2.73; N, 10.82. Found: C, 37.10; H, 2.76; N, 10.80. IR data (KBr, cm1): 3074 (m, C–H), 2920 (m, C–H), 1607–1483 (d, C– H), 1385–1308(d, C–H), 1111(m, S–O), 1083 (m, S–O). 2.3.4. mer-[RuCl3(dmso)(dppz)] (4) The complex was obtained by the exchange of the phen in 2 with dppz (112 mg, 0.41 mmol). The crude product was purified by column chromatography on a neutral alumina with dried CH2Cl2 and CH3OH (10:1, v/v) as eluent, which gave 4 as a dark red solid. Yield: 0.149 g (60%). Red needle-like crystals were obtained by slow evaporation of the concentrated CH3CN solution of 4 at room temperature. Yield: 0.087 g (35%). ESI-MS (m/z): 568.9 for [RuCl3(dmso)(dppz)]. Elemental anal. Calcd. for C20H16Cl3N4ORuS: C, 42.30; H, 2.84; N, 9.87. Found: C, 42.33; H, 2.82; N, 9.86. IR data (KBr, cm1): 3076 (m, C–H), 2938 (m, C–H), 1613–1489 (d, C–H), 1356–1309 (d, C–H), 1131 (m, S–O), 1076 (m, S–O), 3499 (m, N–H). 2.4. Crystal structure determination and refinement of complexes 3 and 4 X-ray diffraction measurements were performed on a Brucker Smart 1000 CCD diffractometer with Mo Ka radiation (k = 0.71073 Å) at 293 K for 3 and 4. All empirical absorption corrections were applied by using the SADABS program [49]. The structures were determined using direct methods, which yielded the positions of all non-hydrogen atoms. These were refined, first, isotropically and, then, anisotropically. All the hydrogen atoms of the ligands were placed in calculated positions with fixed isotropic thermal parameters and the structure factor calculations were included in the final stage of full-matrix least-squares refinement. All calculations were performed using the SHELXTL-97 system of the computer programs [50].

Ru(III) complexes, sodium ascorbate and reduced glutathione in buffer D. The mixture was incubated at 37 °C for 24 h. A covered heating block was used to prevent solvent evaporation during the experiment. A dye solution (bromophenol blue 0.05%, glycerol 5%, and 2 mM EDTA) was added to the reaction mixture prior to electrophoresis. The samples were analyzed by electrophoresis for 3 h at 70 V on a 1% agarose gel in buffer E. Then the gel was stained with 1 lg/ml EB (ethidium bromide) and photographed on an Alpha Innotech IS-5500 fluorescence chemiluminescence and visible imaging system. 2.6.2. Competitive binding with EB The experiments of DNA competitive binding with EB were carried out in buffer C by keeping [DNA]/[EB] = 5 and varying the concentrations of the Ru(III) complexes. The fluorescence spectra of EB were measured using excitation wavelength at 537 nm and the emission range was set between 550 and 750 nm. The control experiments were performed by keeping [EB] = 20 lM (in the absence of DNA, [Ru] = 0–20 lM). The spectra were analyzed according to the classical Stern–Volmer equation (1), where I0 and I are the fluorescence intensities at 590 nm in the absence and presence of the complexes, respectively, Ksv is the linear Stern–Volmer quenching constant, r is the ratio of the total concentration of the complexes to that of DNA ([Ru]/[DNA]). I0 =I ¼ 1 þ K sv r

ð1Þ

2.6.3. Thermal denaturization of CT-DNA Thermal denaturization experiments were performed on a UV–vis spectrophotometer equipped with a Peltier temperature controller in buffer B. The temperature of the cell containing the cuvette was ramped from 50 to 100 °C at 1 °C/min rate, and the absorbance at 260 nm was measured every 1 °C. The Tm values were determined from the maximum of the first derivative or tangentially from the graphs at midpoint of the transition curves. DTm values were calculated by subtracting Tm of the nucleic acid with complex from Tm of the free CT-DNA. 2.6.4. Viscosity measurements Viscosity measurements were carried out using an Ubbdlodhe viscometer maintained at a constant temperature of 30.0 ± 0.1 °C in a thermostatic bath. CT-DNA samples with an approximate average length of 200 base pairs were prepared by sonication in order to minimize complexities arising from DNA flexibility [51]. Flow time was measured with a digital stopwatch. Each sample was measured at least three times and an average flow time was calculated. Data were presented as (N/N0)1/3 versus binding ratio ([Ru]/ [DNA]) [52]. Viscosity values were calculated from the observed flow time of DNA-containing solutions corrected for the flow time of buffer alone (t0), N = tt0. 2.7. Cell culture and cytotoxicity assay

2.5. Solution chemistry and stability studies The absorption spectra in the UV–vis region were recorded at 25 °C. The Ru(III) complexes were first dissolved in a minimum amount of dmso (0.5% of the final volume), and then diluted with phosphate buffer (buffer A) or Tris–HCl buffer (buffer C). The stability studies were carried out by monitoring the electronic spectra of the resulting mixtures over 24 h. 2.6. DNA interactions 2.6.1. DNA cleavage study The cleavage reactions were carried out in a total volume of 10 ll containing pBR322 DNA (0.1 lg) and different amounts of

Cells were supplied by Center of Experimental Animal Sun YatSen University (Guangzhou, China). Cells were routinely kept in RPMI-1640 medium or DMEM medium supplemented with 10% fetal bovine serum, penicillin G (100 U/ml) and streptomycin (100 lg/ml) at 37 °C in a humidified atmosphere containing 5% CO2. After a confluent cell layer was formed, the cells were harvested from the adherent cultures using 0.125% trypsin + 0.01% EDTA in phosphate buffered saline or D-Hank’s buffer for 5 min. Suspensions were adjusted to cell densities of 5  104 cells/ml in order to ensure exponential growth throughout drug exposure. Aliquots of these suspensions were seeded into 96-well microcultures (100 ll/well). After incubation for 24 h cells were exposed to the tested compounds of serial concentrations. The compounds were

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dissolved in dmso and diluted with RPMI-1640 or DMEM to the required concentrations prior to use. After incubation for 20 h, 20 ll of aqueous MTT solution (5 mg/ml) was added to each cell, and the cells were incubated continually for another 4 h, the medium and MTT mixtures were removed and the formazan crystals were dissolved in 100 ll dmso/cell. The absorbance of each cell at 450 nm was determined by analysis with a microplate spectrophotometer (Thermo Lab Systems, Helsinki, Finland), and the percentage cell viability was determined by dividing the average absorbance of each column of Ru(III)-treated wells by the average absorbance of the control wells. The IC50 values were determined by plotting the percentage viability versus concentration on a logarithmic graph and reading off the concentration at which 50% of cells remain viable relative to the control. Each experiment was repeated at least three times to get the mean values. 3. Results and discussion 3.1. Synthesis and description of molecular structures Numerous ruthenium complexes with two or three N–N ligands are known, but examples of mono (N–N) complexes remain scarce, because it is difficult to prevent ligand redistribution during synthesis [42]. Complexes 1 [46] and 2 [44] are prepared following the literature method. 3 and 4 are obtained by using cis-[RuCl2(dmso)4] as the starting material. As proposed by Cingi et al., the Ru(II) precursor is oxidized to Ru(III) by dmso when HCl is present [53].

Fig. 3. Atomic displacement plot (30% probability level) of mer-[RuCl3(dmso)(dpq)] (3). H atoms are omitted for clarity.

The structures of 3 and 4 are confirmed by X-ray diffraction. The crystal structures are shown in Figs. 3 and 4 together with atomic numbering scheme. Relevant details about the structural refinements are listed in Table 1. Selected bond lengths and angles for 3 and 4 are listed in Tables S1 and S2 (Supplementary material), respectively. For complexes 3 and 4, the coordination sphere around the ruthenium center constitutes a distorted octahedron with coordination of one N–N ligand. The S-bonded dmso and one of the three chloride atoms are trans to one of the nitrogen atoms, respectively. Compared with other reported complexes with the same formula of mer-[RuCl3(dmso)(N–N)], the Ru–S bond lengths in the newly synthesized complexes (3: 2.3151(11) Å; 4: 2.3222(17) Å) fall into the same range (2.3236(5) Å [46], 2.2971(8) Å [44], 2.2912(8) Å [54]). The S–O bond lengths (3: 1.452(4) Å; 4: 1.435(5) Å) are shorter than that in free dmso (1.531(5) Å) [55], showing a considerable increase of double-bond character of the S–O bond on

Table 1 Crystallographic data for 3 and 4 Complex

3

Empirical formula Formula weight Temperature Wavelength (Å) Crystal system Space group Unit cell dimensions (Å, °)

4

C16H14Cl3N4ORuS C20H16Cl3N4ORuS 517.79 567.85 293(2) 293(2) 0.71073 0.71073 Monoclinic Orthorhombic P2(1)/n Pbca a = 7.7761(11) 8.5164(9) b = 22.215(3) 19.707(2) c = 10.6271(15) 25.067(3) a = 90.00 a = 90.00 b = 95.013(2) b = 90.00 c = 90.00 c = 90.00 3 Volume (Å ) 1828.7(4) 4207.0(7) Z 4 8 Density (calculated; g cm3) 1.881 1.793 Absorption coefficient 1.424 1.247 Reflections collected 15346 22869 Independent reflections 3993 4597 Observed reflections 2896 3098 Rint 0.0365 0.0421 R1 (all) 0.0630 0.0902 l (cm1) 1.424 1.247 R1a [I > 2r(I)] 0.0376 0.0575 w R2a 0.0945 0.1518 b GOF 1.041 1.057 n h i Ph io1=2 P P a R1 ¼ kF 0 j  jF c k= jF 0 j; wR2 ¼ R wðF 20  F 2c Þ2 = . wðF 20 Þ2 nP h io1=2 2 2 2 b GOF ¼ wðF 0  F c Þ =ðn  pÞ where n is the number of data and p is the number of parameters refined.

Fig. 4. Atomic displacement plot (30% probability level) of mer-[RuCl3(dmso)(dppz)] (4). H atoms are omitted for clarity.

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coordination to ruthenium through the sulfur atom. The bonds between Ru(III) ion and the N–N bidentate ligands are similar to those found in other ruthenium(II) polypyridine complexes [56]. 3.2. Solution chemistry The electronic spectra of 1–4 in acetonitrile all feature two bands with absorption maxima (Fig. 5). The electronic absorption spectra of Ru(II) and Ru(III) complexes with N, N-coordinated ligands contain bands that in most cases can be interpreted as arising from one or more of the following types of transitions: intraligand, metal to ligand charge transfer, ligand to metal charge transfer and p–p* ligand excitations. The intense bands at the higher energy showing a much greater dependence on the nature of the N–N ligands are ligand localized. The remaining less intense bands are attributed to charge transfer transitions from the chlorides to Ru(III) [57,58]. The UV–vis absorption spectra of 1–4 have also been measured in other solvents such as aqueous solution, CH3OH, and dmso (Supplementary material; Figs. S1–S4). It can be seen from these graphs that all of these bands are sensitive to the nature of the solvents, with 1 and 4 providing clear examples of solvatochromism. Though substitution reactions may take place at higher temperatures, especially in solvents possessing potential coordination abilities, complexes 1–4 are stable for several days when dissolved in these organic solvents at room temperature as monitored by UV–vis measurements. No steady-state emission is observed for all the complexes in both organic and aqueous solvents. Although the stability of Ru(III) complex can be measured by NMR spectroscopy, considering Ru(III) complexes are paramagnetic, the UV–vis absorption method is more convenient, especially when buffered solutions are used. The solubility of 1–4 in aqueous media follows the order 1 > 2 > 3 > 4. The various compounds are first dissolved in dmso. All four compounds are highly soluble and stable within this medium. Afterward, concentrated dmso solutions of each Ru(III) compound (1  102 M) are diluted in buffer A, to final concentrations of 1  104 M, and the samples are analyzed spectrophotometrically over 24 h at 25 °C. The resulting spectral profiles for 1–4 are shown in Fig. 6. In freshly prepared buffered solution of 1, two absorption bands are observed in the visible region, one at 296 nm and the other at 381 nm. In the first 30 min after dissolution of 1, the band lying at the higher energy region increases (ca. 29%) accompanied by a 4 nm blue shift of the absorption maximum, while at the same time the band at 381 nm decreases until it totally disappears, during which another band at 447 nm is formed. The formation of the

neat isosbestic points at 350 nm and 419 nm is an indication of the equilibrium between the starting compound and the first hydrolysis product (Fig. 6a). Consecutively, the spectra recorded in the course of a day provide evidence of the emergence of a second process, during which slow but steady decrease of the higher energy band (ca. 18%) with an 8 nm blue shift can be observed, and the newly formed band at 447 nm decreases as well, accompanied by the formation of another isosbestic point at 394 nm (Fig. 6b). No distinct spectra changes can be observed in the following 2 days. The hydrolysis profile of 2 is much similar to that of 1 (Fig. 6c and d). While for 3 (Fig. 6e) and 4 (Fig. 6f), only one process can be observed during 24 h, and the neat isobestic points clearly indicate the formation of only one new product for each compound. By choosing 2 as the model complex, the stability of these compounds under both acidic (buffer A, pH 3) and basic (buffer A, pH 11) conditions has also been investigated. When 2 is dissolved in acidic phosphate buffer, steady decrease of the whole spectra is observed (Supplementary material; Fig. S5), which may be ascribed to the slow precipitation of the compound. The decrease in absorption is not due to the formation of a new compound, because in that case, neat isobestic points would be expected, and the color of the solution stays yellow over 24 h monitored. When 2 is dissolved in basic conditions, the first hydrolysis step, which is completed within 10 min, is much faster than that observed in physiological conditions, and consecutively, in the course of a day, a second hydrolysis step also occurs, during which the color of the solution changes from yellow to green, and no further spectra change is observed over the following 48 h (Supplementary material; Fig. S6). Considering phosphate is a nucleophilic and potentially bidentate chelating ligand, control experiments using Tris–HCl buffer (buffer C) have also been carried out, and the hydrolysis profile observed in Tris–HCl buffer is almost the same as that in phosphate buffer. It can be concluded that: (1) the stability of 1–4 in buffered solutions is increased by extension of the surface areas of the N– N ligands. It is shown that the release of the leaving groups by 1 and 2 is much faster than that by 3 and 4, and the hydrolysis degree of 1 and 2 observed is higher than that of 3 and 4; (2) similar to that of NAMI and KP1019 type complexes, the hydrolysis of 1–4 is affected by the pH value. In acidic conditions, the complexes are stable, while in basic conditions, the hydrolysis process is accelerated, and the color change indicates that poly-oxo Ru species might be formed, as previously reported for NAMI [19] and NAMI-A [22]. 3.3. DNA interactions

1.2 1 2 0.8 A

3 4 0.4

0.0 300

400

500

wavelength / nm Fig. 5. UV–vis absorption spectra of 1–4 (2  104 M) in CH3CN (e = 5A  103 l mol1 cm1).

3.3.1. Nuclease activity of the complexes Many metal complexes exert DNA cleavage activity depending on light, oxidizing or reducing agents, but both oxidative cleavage and photocleavage are unfavorable for therapeutic applications, because they cause diffusible and uncontrolled damages and produce non-natural fragments of DNA, which hampers the further enzymatic manipulation such as labeling and ligation [59,60]. Interestingly, though all of these complexes can not cleave DNA under irradiation, after being incubated with DNA for 24 h under physiological conditions, three of them show cleavage ability in the absence of the UV light or reducing agent. The gel electrophoretic separation showing the cleavage of plasmid pBR322 DNA induced by the complexes under identical conditions is shown in Fig. 7. Complex 1 shows no cleavage activity, and the observed DNA cleavage activity order for other complexes is 4 > 3 > 2 (Fig. 7a). Complexes 3 [40] and 4 can cleave DNA in a dose-dependent manner, and complex 4 with dppz as the intercalating ligand can almost completely cleave DNA from supercoiled from (FORM I) to nicked circular form (FORM II) at 40 lM (Fig. 7b).

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1.6

1.6

1h

0 min 1.5 min

1.2

2h

1.2

4h

0.8

18 min

A

A

9 min

8h

0.8

24 h

36 min 0.4

0.4

0.0 300

400

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0.0

600

300

Wavelength/nm 1.2

400 500 Wavelength/nm

1.2

A

0.8

0.4

27 min 1.75 h 3h 4.5 h 6h 10 h 24 h

0.8 A

0 min 3 min 6 min 9 min 18 min 27 min 51 min

600

0.4

0.0

0.0 300

400

500

300

600

400

500

Wavelength/nm

0 min 6 min 1.5 h 3h 4.5 h 6h 12 h 24 h

0 min 1.2

0.8

2h

A

6h

A

4h 0.8

8h 0.4

600

Wavelength/nm

0.4

24 h

0.0

0.0 300

400

500

Wavelength/nm

600

300

400

500

600

Wavelength/nm

Fig. 6. Time-dependent UV–vis absorption spectra of mer-[RuCl3(bpy)(dmso)] (a and b), mer-[RuCl3(phen)(dmso)] (c and d), mer-[RuCl3(dpq)(dmso)] (e) and mer-[RuCl3(dppz)(dmso)] (f) in buffer A at 25 °C recorded with time.

Fig. 7. Agarose gel electrophoresis mobility patterns for the cleavage of pBR322 DNA incubated for 24 h at 37 °C with Ru(III) complexes. (a) Lane 1, DNA alone; Lane 2, DNA + 20 lM 2; Lane 3, 20 lM 3; Lane 4, 20 lM 4. (b) Lane 1, DNA alone; Lane 2, DNA + 5 lM 4; Lane 3, 10 lM 4; Lane 4, 20 lM 4; Lane 5, 25 lM 4. (c) Complex 3 and (d) complex 4; Lane 1, DNA alone; Lane 2, 40 lM sodium ascorbate + DNA; Lane 3, 40 lM reduced glutathione + DNA; Lane 4, 20 lM complex + DNA; Lanes 5–7, 20 lM complex + sodium ascorbate (20 lM, 40 lM, 80 lM) + DNA; lanes 8–10, 20 lM complex + reduced glutathione (20 lM, 40 lM, 80 lM) + DNA.

C. Tan et al. / Journal of Inorganic Biochemistry 102 (2008) 1644–1653

3.3.3. Thermal denaturization of the DNA Intercalation of small molecules into the double helix is known to increase the helix melting temperature, the temperature at which the double helix denatures into single-strand DNA. The extinction coefficient of DNA bases at 260 nm in the double-helical form is much less than that in the single strand form, hence, melting of the helix leads to an increase in the absorption at this wavelength. Thus, the helix to coil transition temperature can be determined by monitoring the absorbance of the DNA bases at 260 nm as function of temperature [65–68]. The thermal-denatur-

2.5 K

2 2.0

sv

= 1.73

I0/ I

Emission Intensity

600

1.5 400 1.0 0.0

0.2 0.4 [Ru]/[DNA]

200

0 550

600

650

0.6

700

0.8

750

Wavelength/nm 800 3.0 K

2.5

4 600

I0/ I

3.3.2. Competitive binding with EB By only measuring the ability of a complex to affect the EB fluorescence intensity in the EB–DNA adduct, fluorescence quenching method can be used to determine the affinity of the complex for DNA, whatever their binding modes may be. If a complex can replace EB from DNA-bound EB, the fluorescence of the solution will be quenched due to the fact that free EB molecules are readily to be quenched by the surrounding water molecules [63]. For all the complexes, no emission is observed either alone or in the presence of CT-DNA in buffer C. The control experiments show that there is almost no change in the fluorescence intensity of free EB (in the absence of DNA) on increasing concentration of complexes 1–4 (data not shown). The emission spectra of EB–DNA system in the presence and absence of Ru(III) are shown in Fig. 8. The addition of 1 to DNA, pretreated with EB, causes small reduction in emission intensity relative to that observed in the absence the complex at a [Ru]/[EB] = 1 (data not shown), while appreciable reductions in emission intensities are achieved by addition of 2 and 4. The quenching plots of I0/I vs. [Ru]/[DNA] (the inset of Fig. 8) are in good agreement with the linear Stern–Volmer equation with Ksv values of 1.73 and 4.47 for 2 and 4, respectively. By taking a DNA binding constant of 1.0  107 M1 for EB [64], apparent DNA binding constants of 3.43  106 M1 (KEB/2.91) for 2 and 8.62  106 M1 (KEB/ 1.16) for 4 are derived. It has been reported by us that the Ksv of 3 measured under the same conditions is 3.03 [40], and the DNA binding constant of 3 is 6.9  106 M1, so the order of the binding affinity is 1 < 2 < 3 < 4, which is consistent with the previous DNA cleavage results. The result shows that the strength of interactions between DNA and the complexes is enhanced by the increase of the planar area of the N–N ligand. The coordination of Ru(III) towards the nucleobases of DNA is obviously not the main cause of the competitive binding, because in that case 1 should exert the same quenching effect, so the competitive binding by the complexes is mainly caused by their intercalating or groove binding with DNA.

800

Emission Intensity

The influence of sodium ascorbate and reduced glutathione has also been investigated (Fig. 7c and d). No apparent cleavage can be detected using sodium ascorbate and reduced glutathione as controls (Lane 2 and Lane 3). The addition of sodium ascorbate does not enhance the cleavage abilities of all the complexes (Lanes 5– 7), and actually, the cleavage abilities are slightly inhibited by the addition of reduced glutathione (Lanes 8–10). It is obvious that the cleavage abilities are parallel with the planar areas of the ligands (1 < 2 < 3 < 4) involved. We suppose that the nuclease activity mainly comes from the intercalation of the planar ligands with the double helix instead of the covalent binding of Ru(III) with DNA. The possibility of the oxidative pathway is ruled out by the fact that the addition of sodium ascorbate makes no difference. These complexes may resemble KP1019 [61] and cisplatin [62], which have high affinities to sulfur-containing substrates such as cysteine, methionine, tripeptide glutathione and human serum albumin, so the slight quenching effect of glutathione is possibly due to the decreased bioavailability of Ru(III) for DNA binding caused by the interaction of the complexes with glutathione.

sv

= 4.47

2.0 1.5

400 1.0 0.0 200

0 550

600

0.1 0.2 [Ru]/[DNA]

650

700

0.3

0.4

750

Wavelength/nm Fig. 8. Emission spectra of EB bound to DNA in the presence of complexes 2 and 4. [EB] = 20 lM, [DNA] = 100 lM; [Ru]/[DNA] = 0–0.70; kex = 537 nm. The arrows show the intensity changes upon increasing concentrations of the complexes. Inset: plots of I0/I vs. [Ru]/[DNA] with j for experimental data points and full line for linear fitting of the data.

1.0 0.8

(A - A0 )/(Af - A0 )

1650

0.6 0.4 0.2 0.0 50

60

70

80

90

100

T /ºC Fig. 9. Thermal denaturization curves of CT-DNA (60.0 lM) at the concentration ratios of [Ru]/[DNA] = 1/10. (j for DNA alone; s for 1 and DNA; h for 2 and DNA; N for 3 and DNA [40];  for 4 and DNA).

ization profiles of CT-DNA in the absence and presence of complexes 1, 2, and 4 are shown in Fig. 9 with the profile of 3 [40] included for comparison purpose (salt concentration: 0.01 M

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C. Tan et al. / Journal of Inorganic Biochemistry 102 (2008) 1644–1653

Table 2 IC50 values of complexes NAMI, 1, 2, and 4 towards different cell lines determined by the MTT assay with those of 3 [40] included for comparison

1.3

Complex

BEL-7402

CNE-2

MCF-7

HELA

NAMI

746

735

>1000

699

537

1 2 3 4

695 584 721 655

705 626 611 466

955 803 700 745

691 679 580 705

699 585 636 512

0

(η /η )

1/3

1.2

1.1

1.0 0.00

0.04

0.08

0.12

0.16

[Ru]/[D Fig. 10. Effects of increasing amounts of EB (), Ru(phen)2(dppz)2+ (j), 1 (d), 2 (N), 3 (.), and 4 (w) on the relative viscosities of CT-DNA ([DNA] = 0.5 mM), shown as a function of the concentration ratios of [Ru]/[DNA]. g is the viscosity of DNA in the presence of compounds and g0 is the viscosity of DNA alone.

IC50 (lM) SCG-7901

increasing the amounts of complexes 3 and 4, the relative viscosity of DNA increases steadily similarly to the behavior of EB and Ru(phen)2(dppz)2+. The increased degree of viscosity, which may depend on the binding affinity to DNA, follows the order of EB > Ru(phen)2(dppz)2+ > 4 > 3, These results suggest that the principle mode of DNA binding by 4 involves base-pair intercalation, and the binding affinity of complex 4 is higher than that of complex 3, which is consistent with the above experimental results. 3.4. In vitro antitumor potency

NaClO4). In the absence of Ru(III) complexes, CT-DNA shows a main transition at Tm = 68.1 °C. No distinct increase in the melting temperature is observed in the presence of 1 and 2, while the Tm of CT-DNA is increased upon addition of 4. The increase is more pronounced for the addition of 4 (DTm = 10.8 °C) than that of 3 (DTm = 5.3 °C; [40]). Compared with some DNA metallointercalators, which give DTm values of 10–14 °C [69], the large increased DTm suggest an intercalative binding of 4 to DNA. As it has been reported that the effect of NAMI and KP1019 on Tm is dependent on the salt concentration [28], we have also investigated the DTm values caused by 4 under different ionic strength. Less profound increase of Tm is observed (DTm = 7.5 °C) in salt concentrations of 0.1 M NaClO4. At even higher salt concentrations (0.2 M NaClO4) the modification of DNA by 4 results in no change of Tm. Three factors may account for the thermal stability of DNA modified by metal complexes, including: stabilizing effects of the positive charge on the metal moiety and of DNA interstrand cross-links, and a destabilizing effect of conformational distortions such as intrastrand cross-links by platinum coordination [70], and the presence of potential intercalative ligands should also be taken into account in this case. We suppose that at high salt concentrations, the electrostatic effects of the Ru(III) complexes are lowered with increasing concentration of Na+ counter ions, and the relatively unchanged Tm is a consequence of competition between the stabilizing effects and the destabilizing effects. The solution behavior of the DNA adducts of 1–4 appears different from those of NAMI-A and platinum complexes, which merits further study. 3.3.4. Viscosity measurements Optical photophysical probes are not sufficient to support a binding model, while hydrodynamic measurements (i.e., viscosity and sedimentation) that are sensitive to the length change are regarded as the least ambiguous and the most critical tests of a binding model in solution when no crystallographic structural data can be achieved [71]. Intercalation is expected to increase the DNA viscosity; in contrast, a partial, non-classical intercalation of the ligand can reduce its effective length, and concomitantly, its viscosity by bending the DNA helix [72]. Complex 3 [40] and the established intercalator EB and [Ru(phen)2(dppz)]2+ are included as references. As can be seen from Fig. 10, both EB and Ru(phen)2(dppz)2+ can increase the relative viscosity of DNA greatly, and the extent of the viscosity increase caused by EB is more obvious, which is consistent with the literature reports [73,74]. Upon

Cytotoxicity tests are performed using the MTT assay, following exposure of five tumor cell lines (gastric cancer cells (SCG-7901), hepatic cancer (BEL-7402), human low differentiation nasopharyngeal carcinoma (CNE-2), breast cancer (MCF-7), and cervical cancer (HELA)) to the synthesized complexes at increasing concentrations for 24 h. Because the solubility of the compounds is limited in water, a dmso stock solution is used for all compounds to preform a proper comparison among the complexes, and blank samples containing the same amount of dmso are taken as controls. NAMI has been included as the control, and it shows moderate cytotoxicity, which is in well accordance with the literature reports [38]. These complexes exhibit dose-dependent growth inhibitory effect against the tested cell lines and the concentration–response curves of them is shown in Figs. S7–S11 (Supplementary material). Table 2 demonstrates the IC50 values obtained from non-linear regression analysis of dose response data for the compounds tested. The cytotoxicities of 1, 2, and 4 are rather moderate, with IC50 values ranging between 580 and 955 lM, which are comparable to those of NAMI, and no apparent difference in cytotoxicities of 1–4 can be detected. 4. Conclusions In summary, a new series of Ru(III) complexes have been synthesized and characterized, all of the newly synthesized complexes (1, 3, and 4) are obtained in the crystalline state and, thus, their structure can be securely resolved. UV–vis spectra changes show that all of them undergo ligand substitution under physiological conditions, and both the speed and degree of hydrolysis are varied. The complexes presented can cleave plasmid DNA and quench the fluorescence of EB bound to DNA to a considerable degree, both the experiments show that the binding affinity between DNA and 1–4 follows the order: 1 < 2 < 3 < 4. Further studies of thermal denaturization of CT-DNA and viscosity measurements confirm that similar to 3 [40], 4 binds to DNA through intercalation. Though it is well documented that multichloro Ru(III) complexes can damage DNA through covalent binding after they have lost one or more chlorides, non-covalent binding seems to play a more important role for 1–4 in their binding modes with DNA. Like other types of ruthenium complexes, cytotoxicity of 1, 2, and 4 are relatively low. Nevertheless, considering the moderate cell cytotoxicity of NAMI-A species in vitro [29,38], general toxicity

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are low, but the prospect of clinical applications can not be eliminated completely. Comparison of the cytotoxicities of 1–4 shows no apparent differences. The lack of correlation between cell growth inhibition, DNA binding affinity, and hydrolysis stability suggests that differences in cellular uptake, subcellular distribution, and additional biological targets may play even more important roles in their antitumor activity. The in vivo test and protein binding studies of these complexes are ongoing in our laboratory. 5. Abbreviations bpy phen dpq dppz dmso EB NAMI-A ICR KP1019 CT-DNA azpy terpy cdta en MTT RPMI DMEM EDTA

2,20 -bipyridine 1,10-phenantroline 1,10-phenanthroline dipyrido[3,2-f:20 ,30 -h]quinoxaline dipyrido[3,2-a:20 ,30 -c]phenazine dimethyl sulfoxide ethidium bromide [H2im][trans-RuCl4(dmso-S)(Him)] (Him = imidazole) [H2im][trans-RuCl4(Him)2] (Him = imidazole) [H2ind][trans-RuCl4(Hind)2] (Hind = indazole) calf thymus DNA 2-phenylazopyrine 2,20 :60 200 -terpyridine 1,2-cyclohexanediminetetraacetate ethylenediamine 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide Roswell park memorial institute medium Dulbecco’s modified Eagle’s medium ethylenediaminetetraacetic acid

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