Journal of Inorganic Biochemistry 130 (2014) 15–27

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Dual topoisomerase I and II poisoning by chiral Ru(II) complexes containing 2-thiophenylimidazo[4,5-f][1,10]phenanthroline derivatives Yu-Chuan Wang a,1, Chen Qian a,1, Zai-Li Peng a, Xiao-Juan Hou a,b, Li-Li Wang a, Hui Chao a,⁎, Liang-Nian Ji a,⁎ a MOE Key Laboratory of Bioinorganic and Synthetic Chemistry, State Key Laboratory of Optoelectronic Materials and Technologies, School of Chemistry and Chemical Engineering, Sun Yat-Sen University, Guangzhou 510275, PR China b Huaihua Medical College, Huaihua 418000, PR China

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Article history: Received 13 June 2013 Received in revised form 25 September 2013 Accepted 25 September 2013 Available online 7 October 2013 Keywords: Ru(II) complexes DNA-binding Topoisomerase inhibition Apoptosis

a b s t r a c t A series of chiral Ru(II) complexes bearing thiophene ligands were synthesized and characterized. Both Ru(II) complexes Δ/Λ-[Ru(bpy)2(pscl)]2+ (Δ/Λ-1) and Δ/Λ-[Ru(bpy)2(psbr)]2+ (Δ/Λ-2) (bpy = 2,2′-bipyridine, pscl = 2-(5-chlorothiophen-2-yl)imidazo[4,5-f][1,10]phenanthroline, psbr = 2-(5-bromothiophen-2-yl) imidazo[4,5-f][1,10]phenanthroline) showed antitumor activities against A549, HepG2 and BEL-7402 tumor cell lines, especially HeLa tumor cell line. Moreover, Δ enantiomers were more active than Λ enantiomers, accounting for the different cellular uptake. In addition, with the extension of time, these enantiomers could finally accumulate in the nucleus, suggesting that nucleic acids were the cellular target of these enantiomers. The DNAbinding behaviors of complexes were studied using spectroscopic and viscosity measurements. Results suggested that four complexes could bind to DNA in an intercalative mode but no obvious DNA-binding selectivity between the enantiomers was observed. Topoisomerase inhibition and DNA religation assay confirmed that four complexes acted as efficient dual topoisomerase I and II poisons, DNA strand breaks had also been observed from alkaline single cell gel electrophoresis (comet assay). Δ-1 and Δ-2 inhibited the growth of HeLa cells through the induction of apoptotic cell death, as evidenced by the Alexa Fluor® 488 annexin V staining assays and flow cytometry analysis. The results demonstrated that Δ/Λ-1 and Δ/Λ-2 acted as dual topoisomerase I and II poisons and caused DNA damage that could lead to cell cycle arrest by apoptosis. © 2013 Elsevier Inc. All rights reserved.

1. Introduction Topoisomerases (Topo) play a key role in the normal and orderly progression of essential cellular processes. These enzymes transiently break and religate DNA, changing the conformation of a DNA segment and resolving the torsional strain which develops during DNA unwinding and hampers processes such as replication and transcription. Topo I and Topo II are two types of these enzymes, each of which has different mechanisms of action and biological functions [1–5]. Despite variation per tumor tissue in the level of expression of Topo I and/or Topo II, topoisomerases remain a very promising drug target in oncology. During the normal function of topoisomerases, transient Topo–DNA complexes are formed in which the enzymes are covalently linked to DNA. Topo I and II-inhibiting drugs stabilize the Topo–DNA complexes which finally result in lethal DNA damage during replication [6–8]. Several classes of topoisomerase inhibitors have been introduced into cancer clinics as potent anticancer drugs [7,9–12], including camptothecin-based agents targeting Topo I, while anthracyclines, ⁎ Corresponding authors. Fax: +86 20 84112245. E-mail addresses: [email protected] (H. Chao), [email protected] (L.-N. Ji). 1 These authors contributed equally to this work. 0162-0134/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.jinorgbio.2013.09.015

epipodophyllotoxins, aminoacridines, and ellipticines inhibiting Topo II. However, simultaneous treatment of cells in culture with a Topo II (e.g., etoposide) and Topo I (e.g., camptothecin) inhibitor antagonizes cytotoxicity, which is due to the different kinetics and action of Topo I and II through the cell cycle [13–15]. Therefore, drugs as dual Topo I/II inhibitors are among the most active in the series of related compounds. Inhibiting both Topo I and II, these drugs could possibly circumvent development of resistance that arises after treatment with one single topoisomerase inhibitor [16–18]. Ru(II) complexes with polypyridyl ligands have received particular attention in bioinorganic chemistry, due to their combination of easily constructed rigid chiral structures and a rich photophysical repertoire, having prominent DNA binding properties. At present, DNA intercalators represent one of the most important classes of anticancer drugs. Some DNA–intercalating Ru(II) polypyridyl complexes exhibited inhibition activities on Topo II [19–23]. However, studies involving the dual inhibition of Topo I and II by Ru(II) polypyridyl complexes are rare. Only recently, [Ru(bpy)2(bfipH)]2+, [Ru(phen)2(bfipH)]2+ (phen = 1,10-phenanthroline, bfipH = 2-(benzofuran-2-yl)imidazo[4,5-f][1,10] phenanthroline) and Δ/Λ-[Ru(bpy)2(ipad)]2+ (ipad = 2-(anthracene9,10-dione-2-yl)imidazo-[4,5-f][1,10]phenanthroline) were found to act as efficient dual inhibitors of Topo I and II [24,25]. To further study the activity and mechanism of toposiomerase inhibition of Ru(II) polypyridyl

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complexes, here we designed and synthesized two planar aromatic ligands pscl, psbr and their chiral Ru(II) complexes. Thiophenes are a class of drugs typically used as antibiotics, antibacterials and anticancer agents [26,27]. Compounds with a thiophene core have been widely studied due to their ability to bind to DNA and many receptors with high affinity, some of them can also be used as enzyme inhibitors [28–32]. Therefore, we expect that the introduction of the thiophene moiety will result in a higher DNA-binding affinity and inhibition activity towards topoisomerases. As the enantiomers of metal complexes may have distinct DNA-binding properties, both the Δ- and Λ-enantiomers of [Ru(bpy)2(pscl)]2+ and [Ru(bpy)2(psbr)]2+ were synthesized. Results suggested that four complexes bind to DNA in an intercalative mode. No obvious DNA-binding selectivity between the enantiomers was observed. They were demonstrated to efficiently inhibit both of Topo I and II, and consequently caused DNA strand breaks and induced the apoptosis of cell, which was investigated by 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT) assay, real-time cell growth and proliferation assay, comet assay, Alexa Fluor® 488 annexin V staining and cell cycle analysis. The different antitumor activity was in accord with the results of cellular uptake. 2. Experimental 2.1. Materials and measurements All solvents were of analytical grade except those employed in photophysical experiments which were of spectroscopic grade. The compounds 1,10-phenanthroline-5,6-dione [33], Δ-[Ru(bpy)2(py)2][O, O′-dibenzoyl-D-tartrate]·12H2O [34] and Λ-[Ru(bpy)2(py)2][O,O′dibenzoyl-L-tartrate]·12H2O [34] were prepared and characterized according to the literatures. Camptothecin (CPT), etoposide (VP-16), MTT, propidium iodide (PI), ethidium bromide (EB), 4′,6-diamidino-2phenylindoleand (DAPI) and calf thymus DNA (CT-DNA) were obtained from Sigma Aldrich and used without further purification unless specially noted. Other materials were commercially available and of their highest available purity. Tris–HCl buffer (5 mM Tris–HCl, 50 mM NaCl, pH 7.0) solution was prepared using doubly distilled water and used for absorption titration, luminescence titration, viscosity measurements, and CD spectra. A solution of calf thymus DNA in the buffer gave a ratio of UV absorbance at 260 and 280 nm of ca. 1.8–1.9:1, indicating that the DNA was sufficiently free of protein [35]. DNA concentration was determined by absorption spectroscopy using the molar absorptivity (6600 M−1 cm−1) at 260 nm [36]. Microanalysis (C, H, and N) was carried out with a Perkin-Elmer 240Q elemental analyzer. 1H NMR spectra were recorded on a Varian INOVA 500NB NMR spectrometer with (CD3)2SO as solvent at room temperature and TMS as the internal standard. All chemical shifts were given relative to TMS. Electrospray mass spectra (ES-MS) were recorded on a LCQ system (Finnigan MAT, USA) and the quoted m/z values were for the major peaks in the isotope distribution. UV–visible (UV–vis) spectra were recorded on a Perkin-Elmer Lambda 850 spectrophotometer. Emission spectra were recorded on a Perkin-Elmer L55 spectrofluorophotometer at room temperature. The CD spectra were measured on a JASCO-J810 spectrometer. 2.2. Synthesis 2.2.1. Synthesis of ligand pscl A mixture of 1,10-phenanthroline-5,6-dione (0.53 g, 2.5 mmol), 5cholrothiophene-2-carbaldehyde (0.51 g, 3.5 mmol), ammonium acetate (3.88 g, 50 mmol) and glacial acetic acid (15 mL) was refluxed with stirring for 4 h. The cooled solution was diluted with water (ca, 40 mL) and neutralized with concentrated aqueous ammonia. The yellow precipitate was collected and well washed with water, then dried in vacuo. Yield: 0.680 g, 81%. Anal. Calcd for C17H9ClN4S: C, 60.62; H, 2.69; N, 16.64. Found: C, 60.57; H, 2.61; N, 16.58. 1H NMR (500 MHz,

d6-DMSO): δ 8.99 (d, J = 9.5 Hz, 2H), 8.75 (d, J = 13.5 Hz, 2H), 7.78 (dd, J = 13.5, 7 Hz, 2H), 7.68 (d, J = 6.5 Hz, 1H), 7.26 (d, J = 6.5 Hz, 1H). FAB-MS: m/z = 338 (M + 1). 2.2.2. Synthesis of ligand psbr These ligands were synthesized in a manner identical to that described for ligand pscl, with 5-bromothiophene-2-carbaldehyde (0.66 g, 3.5 mmol) in place of 5-cholrothiophene-2-carbaldehyde. Yield: 0.703 g, 74%. Anal. Calcd for C17H9BrN4S: C, 53.56; H, 2.38; N, 14.70. Found: C, 53.63; H, 2.46; N, 14.56. 1H NMR (500 MHz, d6DMSO): δ 9.02 (d, J = 9.5 Hz, 2H), 8.78 (d, J = 13 Hz, 2H), 7.81 (dd, J = 13.5, 7 Hz, 2H), 7.68 (d, J = 6.5 Hz, 1H), 7.41 (d, J = 6.5 Hz, 1H). FAB-MS: m/z = 382 (M + 1). 2.2.3. Synthesis of Δ-[Ru(bpy)2(pscl)](ClO4)2·2H2O (Δ-1) A mixture of Δ-[Ru(bpy)2(py)2][O,O′-dibenzoyl-D-tartrate]·12H2O (0.130 g, 0.1 mmol), pscl (0.067 g, 0.2 mmol), ethylene glycol (18 mL) and water (2 mL) was refluxed for 8 h under argon. Upon cooling, the resulting solution was diluted with 60 ml water. After filtration, the solution was treated with a saturated aqueous solution of NaClO4 to obtain a red precipitate. The crude product was purified by column chromatography on alumina with acetonitrile–toluene (2:1, v/v) as an eluent. The deep red product was further recrystallized with acetonitrile-ether and dried in vacuo. Yield: 0.047 g, 50%. Anal. Calcd for C37H25Cl3N8O8RuS: C, 46.82; H, 2.65; N, 11.81. Found: C, 46.75; H, 2.58; N, 11.88. 1H NMR (500 MHz, d6-DMSO): δ 8.88 (s, 2H), 8.85 (d, J = 13 Hz, 2H), 8.81 (d, J = 14 Hz, 2H), 8.17 (t, J = 13 Hz, 2H), 8.06 (t, J = 13 Hz, 2H), 7.84 (d, J = 8.5 Hz, 4H), 7.74 (dd, J = 13, 9 Hz, 2H), 7.55 (t, J = 8.5 Hz, 5H), 7.34 (t, J = 10.5 Hz, 2H), 7.16 (d, J = 6.5 Hz, 1H). ES-MS [CH3CN, m/z]: 375 ([M-2ClO4 + H]+), 749 ([M-2ClO4]2 +). UV–vis (λ (nm), ε (104 M− 1 cm− 1)) (CH3CN): 461 (1.5), 341 (2.6), 288 (6.8). CD (λ (nm), Δε (M− 1 cm− 1)) (in water): 478 (− 7.8), 420 (9.6), 294 (− 78.2), 277 (23.3), 239 (13.5). 2.2.4. Synthesis of Λ-[Ru(bpy)2(pscl)](ClO4)2·2H2O (Λ-1) This complex was synthesized in a manner identical to that described for complex Δ-1, with Λ-[Ru(bpy)2(py)2][O,O′-dibenzoyl-Ltartrate]·12H2O (0.13 g, 0.1 mmol) in place of Δ-[Ru(bpy)2(py)2][O,O′dibenzoyl-D-tartrate]·12H2O. Yield: 0.058 g, 61%. UV–vis (λ (nm), ε (104 M−1 cm−1)) (CH3CN): 462 (1.6), 341 (2.8), 288 (7.5). CD (λ (nm), Δε (M−1 cm−1)) (in water): 478 (8.1), 420 (−9.7), 294 (79.2), 277 (−24.5), 239 (−15.7). The 1H NMR, elemental analysis and mass spectral data of Λ-1 were tested to be quite same as its Δ enantiomer within the experimental errors. 2.2.5. Synthesis of Δ-[Ru(bpy)2psbr](ClO4)2·2H2O (Δ-2) These complexes were synthesized in a manner identical to that described for complex Δ-1, with psbr (0.076 g, 0.2 mmol) in place of pscl. Yield: 0.058 g, 58%. Anal. Calcd for C37H25BrCl2N8O8RuS: C, 44.73; H, 2.54; N, 11.28. Found: C, 44.82; H, 2.59; N, 11.22. 1H NMR (500 MHz, d6-DMSO): δ 8.83 (m, 6H), 8.18 (t, J = 13 Hz, 2H), 8.06 (t, J = 13 Hz, 2H), 7.84 (d, J = 8.5 Hz, 4H), 7.73 (dd, J = 13.5, 8.5 Hz, 2H), 7.56 (t, J = 10 Hz, 5H), 7.34 (t, J = 11 Hz, 2H), 7.24(d, J = 6 Hz, 1H). ES-MS [CH3CN, m/z]: 397 ([M-2ClO4 + H]+), 795 ([M-2ClO4]2+). UV–vis (λ (nm), ε (104 M−1 cm−1)) (CH3CN): 462 (1.6), 339 (2.7), 287 (7.2). CD (λ (nm), Δε (M−1 cm−1)) (in water): 478 (−8.2), 421 (10.2), 294 (−82.2), 277 (23.6), 239 (15.6). 2.2.6. Synthesis of Λ-[Ru(bpy)2psbr](ClO4)2·2H2O (Λ-2) These complexes were synthesized in a manner identical to that described for complex Δ-1, with Λ-[Ru(bpy)2(py)2][O,O′-dibenzoyl-Ltartrate]·12H2O (0.13 g, 0.1 mmol) in place of Δ-[Ru(bpy)2(py)2][O,O′dibenzoyl-D-tartrate]·12H2O. Yield: 0.065 g, 65%. UV–vis (λ (nm), ε (104 M−1 cm−1)) (CH3CN): 462 (1.6), 339 (2.9), 288 (7.4). CD (λ (nm), Δε (M−1 cm−1)) (in water): 478 (8.7), 421 (−10.5), 294 (83.5), 277 (−25.5), 239 (−17.4). The 1H NMR, elemental analysis

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and mass spectral data of Λ-2 are tested to be quite same as its Δ enantiomer within the experimental errors.

2.3. Cell lines and cell culture Human cancer cell lines, including cervical carcinoma HeLa, hepatocellular carcinoma HepG2, were purchased from American Type Culture Collection (ATCC, Manassas, VA). The human hepatocellular carcinoma BEL-7402, lung carcinoma A549 was obtained from the Cell Bank (Cell Institute, Sinica Academica Shanghai, Shanghai, China). All cell lines were cultured in 25 cm2 culture flasks in either RPMI-1640 (Roswell Park Memorial Institute 1640, Gibco, Gaithersburg, MD) or DMEM (Dulbecco's Modified Eagle's Medium, Gibco, Gaithersburg, MD) culture media supplemented with 10% fetal bovine serum (Hyclone, Waltham, MA), 100 units/mL penicillin and 50 units/mL streptomycin in a humidified incubator with an atmosphere of 95% relative humidity and 5% CO2 at 37 °C. Cells were grown to 70% confluence, trypsinized with 0.25% trypsin, and the cells were ready for the study until the cell growth was in a stable state and the logarithmic growth phase unless otherwise specified.

2.4. Cytotoxicity assays Cells were plated in 96-well microassay culture plates (Costar) and grown overnight at a density of 3 × 103 cells per well. Test compounds in different concentrations were then added to the wells (final DMSO concentration, 0.1% v/v). Control wells were prepared by the addition of 100 μL culture medium with 0.1% DMSO. Wells containing culture medium without cells were used as blanks. The plates were incubated at 37 °C in a 5% CO2 incubator for 48 h. Upon completion of the incubation, stock MTT dye solution (10 μL, 5 mg/mL) was added to each well. After 4 h incubation, the medium was aspirated and replaced with 150 μL DMSO per well to solubilize the MTT formazan. The optical density of each well was then measured on a microplate spectrophotometer iMark (Bio-Rad, CA, USA) at a wavelength of 595 nm. The IC50 value was determined from plots of % viability against dose of compound added. All data were from at least three independent experiments and presented as mean ± standard deviation (SD).

2.5. Real-time cell growth and proliferation assay Experiments were carried out using xCELLigence RTCA DP System Real-Time Cell Analyzer (Roche Diagnostics GmbH, Germany) which was placed in a humidified incubator maintained at 37 °C with 95% air and 5% CO2. Growth curves were constructed using 16-well plates (Eplate 16, Roche Diagnostics GmbH, Germany). The change in impedance caused by cell attachment and spreading is then expressed as the cell index (CI). CI is defined as (Rn −Rb)/15, where Rb is the background impedance of the well measured with edium alone and Rn is the electrode resistances (a component of impedance) of the well measured at any time in the presence of cells. CI is a relative value to indicate how many cells attached to the electrodes. The slope of the CI curve reflects the growth speed of cells [37,38]. Briefly, 80 μL of cell culture media at room temperature was added into each well of E-plate 16. After this, the E-plate 16 was connected to the system and checked in the cell culture incubator for proper electrical-contacts and the background impedance was measured during 24 s. Meanwhile, the cells were resuspended in cell culture medium and adjusted to 5 × 104 cells/mL. 100 μL of each cell suspension was added onto the surface of microelectronic sensors in the wells of E-plate 16. Approximately 24 h after seeding, when the cells were in the log growth phase, a wide range of concentrations of drugs were added and the cells was automatically monitored every 15 min over 48 h by the xCELLigence system. Data analysis was carried out using RTCA Software 1.2 supplied with the instrument.

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2.6. Cellular uptake Cellular uptake was measured by using flow cytometry. HeLa cells in growth medium were seeded in 35 mm tissue culture dishes (Corning) and incubated at 37 °C under a 5% CO2 atmosphere until 70% confluent. The culture medium was removed and replaced with a medium (final DMSO concentration, 1% v/v) containing the Ru(II) complexes at 50 μM. After incubation for 0.5, 3, 6, 12 and 24 h, the cell layer was trypsinized and washed twice with cold phosphate buffered saline (PBS). The samples were raised in 500 μL of cold PBS and analyzed by a FACSCanto II flow cytometer (BD Biosciences, USA) immediately [39]. The samples were collected in FL2 channel (excitation at 488 nm and emission at 585 (21 nm)), and the number of cells analyzed for each sample was 10,000.

2.7. Fluorescence microscopic studies The cellular uptake of Ru(II) complexes was also monitored qualitatively by fluorescence microscope. Briefly, treated cells cultured on microdishes (Thistle Scientific) till 70% confluence were incubated with 50 μM Ru(II) complexes (final DMSO concentration, 0.1% v/v) for various periods of time. Co-staining was performed by using DAPI (500 nm) for 10 min before the cells were washed with PBS and imaged [40,41]. Cultures were luminescently imaged on an inverted fluorescence microscope (Zeiss Axio Observer D1). Ru(II) complexes was excited at 458 nm and emission monitored at 600–640 nm (red) wavelengths. DAPI was excited at 359 nm emission detected at 462 nm (blue) wavelengths.

2.8. Ru–DNA binding experiments The absorption titration of Ru(II) complexes was performed by using a fixed complex concentration (12 μM) to which increments of each DNA stock solution were added until the absorbance did not change any more. Complex–DNA solutions were allowed to incubate for 5min before the absorption spectra were recorded. The intrinsic binding constants Kb of Ru(II) complexes to DNA was determined according to Eqs. (1a) and (1b) [42].

εa −ε f ¼ εb −ε f

qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi  ffi b− b2 −2K 2b C t ½DNA=s 2KC t

b ¼ 1 þ K b C t þ K b ½DNA=2s

ð1aÞ

ð1bÞ

where [DNA] is the concentration of DNA in base pairs, the apparent absorption coefficients εa, εf, and εb correspond to Aobsd/[Ru], the extinction coefficient for the free ruthenium complex, and the extinction coefficient for the ruthenium complex in the fully bound form, respectively. Kb is the equilibrium binding constant in M−1, Ct is the total metal complex concentration, and s is the binding size. Luminescence titration experiments were performed at a fixed metal complex concentration (20 μM) to which increments of a stock DNA solution (0–160 μM) containing the same concentration of the metal complex were added. After the addition of DNA to the metal complex, the resulting solution was allowed to equilibrate at 25 °C in the dark for 10 min before being excited by 380 nm light. Viscosity measurements were carried out at a constant temperature at 28.0 ± 0.1 °C in a thermostatic bath. Flow time was measured with a digital stopwatch, and each sample was measured three times, and an average flow time was calculated. Data are presented as (η/η0)1/3 versus binding ratio, where η is the viscosity of DNA in the presence of complex and η0 is the viscosity of DNA alone.

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2.9. Topoisomerase inhibition assay

2.12. Alexa Fluor® 488 annexin V staining [47]

DNA Topoisomerase I (Topo I) from calf thymus was purchased from MBI Fermentas. No further purification was performed. One unit of enzyme was defined as completely relaxes 1 μg of negatively supercoiled pBR322 DNA in 30 min at 37 °C under the standard assay conditions. The reaction mixture (20 μL) contained 35 mM Tris–HCl (pH 8.0), 72 mM KCl, 5 mM MgCl2, 5 mM dithiothreitol (DTT), 2 mM spermidine, 0.1 mg/ml bovine serum albumin (BSA), 0.25 μg pBR322 DNA, 2 Unit Topo I, and Ru(II) complexes. The reaction mixtures were incubated at 37 °C for 30 min, and the reaction was terminated by the addition of 4 μL of 5× stop solution consisting of 0.25% bromophenol blue, 4.5% SDS, and 45% glycerol. The samples were electrophoresed through 1% agarose in TBE buffer (89 mM Tris-borate acid, 2 mM EDTA, pH 8.3) at 30 V for 8 h. The gel was stained with 1 μg/mL EB and photographed under UV light. The concentrations of the inhibitor that prevented 50% of the supercoiled DNA from being converted into relaxed DNA (IC50 values) were calculated by the midpoint concentration for druginduced DNA unwinding. DNA Topoisomerase IIα (Topo II) from Escherichia coli containing a clone of the human Topoisomerase II gene was purchased from GE Healthcare Bio-Sciences, and no further purification was performed. One unit of the enzyme was defined as completely relaxes 0.3 μg of negatively supercoiled pBR322 plasmid DNA in 15 min at 30 °C under the standard assay conditions. The reaction mixture (20 μL) contained 10 mM Tris–HCl (pH 7.9), 50 mM NaCl, 50 mM KCl, 5.0 mM MgCl2, 0.1 mM EDTA, 15 μg/ml BSA, 1.0 mM ATP, 0.25 μg pBR322 DNA, 5 Unit Topo II, and Ru(II) complexes. The reaction mixtures were incubated at 37 °C for 15 min. Reactions were stopped, processed, and subjected to gel electrophoresis as above.

5×105 cells/well of HeLa cells in 6-well plate with various concentrations (25, 50, and 100 μM) of Ru(II) complexes were incubated in an incubator for 24 h. Cells were trypsinized, washed twice with ice-cold PBS and then re-suspended in 100 μL binding buffer (50 mmol/L HEPES/ NaOH, pH 7.4, 700 mmol/L NaCl, 12.5 mmol/L CaCl2) containing 5 μL of annexin V stock (Invitrogen, Paisley, UK). After incubation for 15 min at room temperature in a light-protected area, another 400 μL binding buffer was added and the specimens were quantified by flow cytometry on a FACSCanto II (BD Biosciences, USA).

2.10. DNA religation assay The DNA religation reaction of human Topo I and Topo II was monitored according to the procedure of Osheroff [43] with minor modifications. The reaction mixtures for Topo I and Topo II were the same as in topoisomerase inhibition assay. Reactions were started by the addition of Ru(II) complexes and incubated for 6 min at 37 °C to allow the cleavage/religation reaction of the enzyme to reach equilibrium. Religation was initiated by shifting reactions from 37 to 0 °C. Reactions were stopped at time points following the temperature shift by adding 4 μL of 5× stop solution. Samples were treated and analyzed as described for topoisomerase inhibition assay. Apparent first-order religation rates were determined by quantitating the loss of linear DNA. 2.11. Single cell gel electrophoresis The alkaline single cell gel electrophoresis (comet assay) is a sensitive, reliable, and rapid method for DNA double- and single-strand breaks as well as alkali-labile sites, is the method of choice to investigate DNA-damage induction in eukaryotic individual cells [44–46]. So, the single cell gel electrophoresis in alkaline condition was performed to measure the DNA strand breaks induced by Ru(II) complexes, CPT or VP-16 in the HeLa cells for 24 h. Briefly, 1 × 104 cells/mL of treated and untreated HeLa cells was mixed with 0.8% low-melting-point agarose at a ratio of 1:10 (v/v) and spread on slides precoated with 1% normal agarose. The embedded cells were lysed in a precooled lytic solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris base and 1% Triton X-100, 10% DMSO; the last two compounds were added fresh, pH 10) at 4 °C for 120 min, rinsed, and equilibrated in an alkaline electrophoresis buffer (0.3 M NaOH and 1 mM EDTA, pH 13). Electrophoresis was performed in an ice-bath at 25 V and 300 mA for 20 min, and the slides were neutralized with 0.4 M Tris–HCl (pH 7.5), stained with SYBR green solution (200 μg/mL) for 5 min in the dark and analyzed using an inverted fluorescence microscope (Zeiss Axio Observer D1). The DNA contents in the head and tail were quantified using “Comet score” from Autocomet.

2.13. Flow cytometric analysis HeLa cells in a density of 1× 106 cells/mL were treated with IC50 concentration of Ru(II) complexes and incubated for 12, 24, 36 and 48 h. Then cells were collected and fixed with 1 mL of 70% cold ethanol (−20 °C) overnight. After incubation, the cells were centrifuged and washed in 1mL PBS and resuspended in 0.4mL PBS. To a 0.5mL cell sample, 50 μL RNase A (1 mg/mL in PBS), was added and incubated for 30 min at 37 °C followed, after gentle mixing by 50 μL propidium iodide (500μg/mL in PBS) solution. The mixed cells were incubated in the dark at room temperature for 15min and kept at 4°C until measured. The cell cycle distribution was analyzed with FACSCanto II (BD Biosciences, USA). The data were acquired and analyzed with BD FACSDiva software v6.0.

2.14. Statistical analysis All the data were expressed as mean ± SD. Differences between two groups were analyzed by two-tailed Student's t test. One-way analysis of variance (ANOVA) was used in multiple group comparisons. These analyses were carried out by SPSS 12.0. 3. Results and discussion 3.1. Synthesis and characterization The synthesis route was shown in Fig. 1. Similar to that described by Steck and Day [48], the ligands pscl and psbr were prepared by condensation reaction using 1,10-phenanthroline-5,6-dione and 5methylthiophene-2-carbaldehyde etc., with the presence of ammonium acetate and glacial acetic acid, giving these ligands a high yield. The corresponding chiral Ru(II) complexes were synthesized by treating the chiral precursors Δ-[Ru(bpy)2(py)2][O,O′-dibenzoyl-D-tartrate]·12H2O or Λ-[Ru(bpy)2(py)2][O,O′-dibenzoyl-L-tartrate]·12H2O with the ligands. In the case of metal complexes, only isotope peaks corresponding to [M–2ClO4–H]+ and [M–2ClO4]2+ were observed. The measured molecular weights were consistent with expected values. Δ-1 and Λ-1 gave the same and well-defined 1H NMR spectra, which permitted unambiguous identification and assessment of purity. Due to the shielding influences of the adjacent pscl and bpy, the bpy protons of complexes exhibited two distinct sets of signals. In addition, the proton resonance on the nitrogen atom of the pscl imidazole ring was not observed, because metal coordination causes electron deficiency in the ligand and, as a result, the imidazole proton becomes active and there is a fast proton exchange between the two imidazole nitrogen atoms [49,50]. Enantiomers Δ/Λ-2 exhibited identical characterization in 1H NMR as that described for Δ/Λ-1 above. The enantiomeric purity of these complexes was assayed by CD spectroscopy measurements. Δ-1 and Δ-2 exhibited two primary CD signals, with a positive peak at approximately 277 nm and a negative peak at approximately 294 nm. As expected, opposing signals were observed for the enantiomers of Λ-1 and Λ-2. The 1H NMR spectra and CD spectra are shown in Fig. S1–S4.

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Fig. 1. Synthetic route for ligands and its enantiomeric Ru(II) complexes.

3.2. Inhibition of cell proliferation and cytotoxicity The in vitro cytotoxic activities of the Ru(II) complexes were evaluated against HeLa, A549, HepG2 and BEL-7402 tumor cell lines. Table 1 shows the IC50 values of Δ/Λ-1 and Δ/Λ-2 by MTT assay after a 48 h treatment. The Ru(II) complexes demonstrated higher in vitro cytotoxicity against selected tumor cell lines than NAMI-A, but relatively low cytotoxicity compared to cisplatin. We can also found that the tested cancer cells, especially the HeLa cells, were susceptible to the complexes. The two enantiomers displayed difference in the antitumor activities, the Δ enantiomers were more active against the A549, BEL7402 and HepG2 tumor cell lines, especially HeLa cells. Thus, this HeLa

Table 1 IC50 values of four Ru(II) complexes towards different tumor cell lines.a Complexes

Δ-1 Λ-1 Δ-2 Λ-2 NAMI-A Cisplatin

IC50 (μM) HeLa

A549

BEL-7402

HepG2

41.8 ± 2.3 77.2 ± 6.5 54.6 ± 2.9 93.1 ± 6.4 515.7 ± 17.4 18.3 ± 1.4

58.1 ± 4.7 73.5 ± 3.2 185.4 ± 8.3 205.8 ± 5.6 573.2 ± 41.6 26.9 ± 1.7

150.5 ± 4.2 185.1 ± 6.3 226.3 ± 9.4 271.1 ± 6.5 635.9 ± 35.4 16.4 ± 0.7

151.7 ± 1.3 162.5 ± 8.6 156.1 ± 5.7 188.5 ± 8.9 548.7 ± 28.6 19.7 ± 1.1

cell line was used for further investigation regarding the underlying mechanisms. Most commonly used conventional in vitro assays for cell viability are end-point assays which eventually also require lysis of cells. Recently real-time monitoring of target cell status based on electrical impedance measurements has been introduced as a continuous and nonradioactive method for assessing cellular cytotoxicity in vitro [38]. 5,000 HeLa cells/ well concentration in the real-time cell growth and proliferation assay was used to examine the toxic effects elicited by Δ-1 and Δ-2, respectively. As shown in Fig. 2, kinetic profiles provided by the xCELLigence System indicated that the rate and dynamics of cytotoxicity were similarly between the complexes. Interestingly, cell killing kinetics were shown to be concentration-dependent. Cells treated at high doses of Ru(II) complexes (100 μΜ) can rapidly abolished continuous cellular proliferation, indicating that cells undergo irreversible senescence. Whereas, cells treated at low doses of Ru(II) complexes (25 μΜ) seem to overcome the cellular damage, reaching control cell index levels with a short delay. This is most likely due to a DNA damage-mediated cell cycle arrest, which is eventually overcome after the damage is repaired by the cellular DNA repair machinery [38]. The impedance-based CI values clearly revealed cytotoxic effects in real-time, allowing pinpointing when to perform biochemical endpoint assays in downstream applications addressing questions in proteomics and genomics. 3.3. Cellular uptake

a

Cells were treated with various concentrations of tested compounds for 48 h. Cell viability was determined by MTT assay and IC50 values were calculated as described in the Experimental section. Each value represents the mean ± SD of three independent experiments.

Because of their stability in aqueous solution and luminescence (the luminescence intensity was increased upon binding with DNA, Fig. 6), the cellular uptake properties of Ru(II) complexes can be studied

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Fig. 2. Kinetics of cytotoxicity responses for Ru(II) complexes in cells monitored by the xCELLigence system. (A) Δ-1 in HeLa, (B) Δ-2 in HeLa. 5,000 cells/well were plated in 16-well strips for the RT-CES cytotoxicity assay. Downward arrow, the time of compound addition.

using flow cytometry and fluorescence microscope [39,51,52]. First, flow cytometry was used to obtain semiquantitative data on the uptake of Ru(II) complexes into HeLa cells (Fig. 3). Data on the uptake of Ru(II) complexes into HeLa cells was determined after 0.5, 3, 6, 12 and 24 h of exposure to the complexes. Incubation with 50 μM complexes for 0.5 h caused a dramatic accumulation in HeLa cells and the following order in the mean luminescence intensity could be seen: Δ-1 N Λ-1 and Δ-2 N Λ-2. With the extension of time, the uptake levels for Δ/Λ-1 and Δ/Λ-2 did not increase remarkably, indicating that the four complexes were quick on the uptake, however, the uptake order could be seen in all of the test times. Additionally, the cellular uptake data obtained for Δ/Λ-1 and Δ/Λ-2 showed a clear correlation with their cytotoxicity. Flow cytometry cannot discriminate among membrane, cytoplasmic, and nuclear localization, while the cellular localization characteristics of anticancer drugs are fundamental to their efficacy [53,54], thus the cellular distribution Ru(II) had been studied by a fluorescence microscope. As shown in Fig. 4, Δ-1 (50 μM) accumulated in the cell membrane after 0.5 h of treatment and the cellular Δ-1 increased in a time-dependent manner after that. After 3 h of incubation with HeLa cells, Δ-1 gradually penetrated into the interior of the nucleus and showed diffuse cytoplasmic and nuclear fluorescence. Using DAPI as a marker of nucleus, we observed the overlay of Δ-1 fluorescence with nucleus (blue) fluorescence with the extension of time, and Δ-1 even excluded DAPI from the nucleus after 24 h, suggesting that nucleic acids were the cellular target of Δ-1. The other Ru(II) complexes

Fig. 3. Mean luminescence intensity determined by flow cytometry in HeLa cells after 0.5,3, 6, 12 and 24 h of exposure to Δ/Λ-1 and Δ/Λ-2 (50 μM). The results are the means of three independent samples and are expressed as the means ± SD.

exhibited the same cellular localization characteristics as Δ-1 (data not show). 3.4. DNA binding studies Previously, fluorescence microscopic studies showed that Ru(II) complexes could readily pass the cell membrane and penetrate into the nucleus (Fig. 4), so genomic DNA may serve as the target of these complexes. Binding of the four Ru(II) complexes with CT-DNA were monitored by absorption spectra titration, luminescence titration and viscosity measurements. There was no obvious DNA-binding selectivity between the enantiomers. The absorption spectra of four Ru(II) complexes in the absence and presence of CT-DNA at various complex concentrations are given in Fig. 5. Upon increasing concentrations of DNA, the hypochromism (H% = 100(Afree − Abound) / Afree) in the metal–ligand charge transfer (MLCT) band reached 10.8% and 12.7% at a ratio of [DNA]/[Ru] of 16.0 for Δ-1 and Λ-1, with a red shift of 7 nm and 7 nm, respectively. For Δ2 and Λ-2, under the same experimental condition, the hypochromism in the MLCT band reached 16.1% and 16.8%, with a red shift of 6 nm and 6nm at a [DNA]/[Ru] ratio of 16.0. In order to compare quantitatively the binding strength of the two complexes, their intrinsic binding constants with CT-DNA were obtained by monitoring the changes in absorbance of the MLCT band for both complexes, with increasing concentration of DNA. Intrinsic binding constants Kb of (6.18 ± 1.60) × 105 M−1, (4.28 ± 1.46) × 105 M−1 and (4.80 ± 1.21) × 105 M−1, (3.32±1.16)×105 M−1 were obtained for Δ/Λ-1 and Δ/Λ-2, respectively. DNA-binding results between Δ/Λ-1 and Δ/Λ-2 were close to each other, together with the experimental errors it was hard to distinguish which enantiomer binds to DNA more strongly. The result is similar to those of Δ-[Ru(bpy)2(uip)]2+ (5.8 × 105 M−1) [55], Λ-[Ru(bpy)2(uip)]2+ (2.9 × 105 M−1) [55], and much higher than [Ru(phen)3]2+ (5.5 × 103 M−1) [56] which interacts with DNA through a semiintercalation or quasiintercalation [57], indicating that the complexes in this work most likely intercalatively bind to DNA. In the absence of DNA, all the four complexes could emit luminescence in Tris buffer at an ambient temperature. Changes in emission spectra of the complex with increasing DNA concentrations are shown in Fig. 6. For Δ-1 and Λ-1, as DNA was successively added into the complex solution, the emission intensities of the complex were gradually increased until stable emission intensity. The intensities of the two complexes in the presence of DNA were increased to around 2.53 and 1.92 times as compared to those of the DNA-free complexes, respectively. Comparing to the well-studied [Ru(phen)2(dppz)]2+ (dppz = dipyridophenazine) (luminescent enhancement N104), these new complexes may not be considered as DNA molecular “light switch” [58]. However, this result implied that all these complexes could interact with DNA to some extent and be protected by DNA efficiently, since

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Fig. 4. Fluorescence microscope images of HeLa cells after having been treated with 50 μM for 0.5, 3, 6, 12 and 24 h. Δ-1 and the nucleus were visualized by red and blue fluorescence, respectively.

the hydrophobic environment inside the DNA helix reduced the accessibility of solvent water molecules to the complex and the complex mobility was restricted at the binding site, leading to the decrease of the vibrational modes of relaxation. However, upon addition of CTDNA, the emission intensities of Δ-2 and Λ-2 were first reduced and reached a saturated value at [DNA]/[Ru]= 9.48 and thereafter gradually increased upon further increasing [DNA]/[Ru] until a stable emission intensity was observed, with I/I0 = 1.24 and 1.33, respectively. These emission spectral characteristics implied the presence of at least two possible binding conformations: one was less emissive and was formed

at the low DNA concentrations; the other formed at the high DNA concentrations and was highly emissive which most probably resulted from intercalative interaction. This effect prevailed over the other emission quenching processes, e.g., the potential photo-induced electron transfer from the guanine base of DNA to the 3MLCT of the complex [59]. Optical photophysical probes provide necessary but not sufficient evidence to support the binding model of Ru(II) complexes with DNA. Hydrodynamic measurements that are sensitive to change in length of the DNA strands (i.e. viscosity and sedimentation) are regarded as the least ambiguous and the most critical tests of a binding model in

Fig. 5. Absorption spectra of Δ-1 (a), Λ-1 (b), Δ-2 (c) and Λ-2 (d) in Tris–HCl buffer upon addition of CT-DNA ([Ru] = 10 μM, [DNA] = 0–280 μM). Arrows indicate the change in absorbance upon increasing the DNA concentration. Inset: Plot of (εa − εf)/(εb − εf) vs [DNA] for the titration of DNA to Ru(II) complexes.

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Fig. 6. Emission spectra of Δ-1 (a), Λ-1 (b), Δ-2 (c) and Λ-2 (d) in 5 mM Tris–HCl and 50 mM NaCl buffer (pH = 7.0) in the absence and presence of CT-DNA ([Ru] = 10 μM, [DNA] = 0–280 μM). Arrows indicate the change in emission intensity upon increasing the DNA concentration.

solution in the absence of crystallographic or NMR structural data. A classical intercalation model demands that the DNA helix must lengthen as base pairs are separated to accommodate the binding ligand, leading to an increase of the viscosity of the DNA solution. In contrast, a partial and/or nonclassical intercalation mode reduces the effective length of the DNA molecule by bending (or kinking) the strand and, therefore, its viscosity. In addition, electrostatic and grooving binding have little effect on DNA viscosities [60,61]. To further elucidate the binding mode of the present complexes, viscosity measurements were carried out on CT-DNA by varying the concentration of the added complexes. The effects of Δ/Λ-1 and Δ/Λ-2, together with [Ru(bpy)3]2+ and EB on the viscosity of rod-like DNA were shown in Fig. 7. EB increases the relative specific viscosity for the lengthening of the DNA double helix through the intercalation mode. However, the [Ru(bpy)3]2+ complex,

Fig. 7. Effect of increasing amounts of EB, [Ru(bpy)3]2+ and the Ru(II) complexes in this work on the relative viscosity of CT-DNA at 28.0 ± 0.1 °C. The total concentration of DNA is 0.5 mM.

which has been known to bind with DNA through electrostatic interactions, exhibited essentially no effect on the viscosity of DNA. On increasing the amounts of Ru(II) complexes, the relative viscosity of DNA is increasing steadily, which is similar to the behavior of EB. The increased degree of viscosity, which may depend on its affinity to DNA, follows the order of EB N Δ-1 ~ Λ-1 N Δ-2 ~ Λ-2 N [Ru(bpy)3]2+. The results suggest that all these Ru(II) complexes in this work intercalate into the base pairs of DNA, which are consistent with our foregoing hypothesis. 3.5. Topoisomerase inhibition The results of Topo I inhibition assay by different concentrations of Δ/Λ-1 and Δ/Λ-2 are shown in Fig. 8. Both Δ-1 and Λ-1 inhibited the ability of Topo I to relax negatively supercoiled plasmid DNA (IC50 ≈ 8 μM). IC50 of Δ-1 and Λ-1 on Topo I activities were calculated as described in the experimental section and listed with some typical topoisomerase inhibitors (Table 2). These findings implied that Δ-1 and Λ-1 may block DNA strand passage event of the enzyme, and inhibited the overall activity of Topo I. Complexes Δ-2 and Λ-2 also showed Topo I inhibitory activity in a concentration-dependent style. For Δ/Λ-1 and Δ/Λ-2, all these complexes inhibited the activity of Topo I at a lower concentration, comparing with some classical topoisomerase inhibitors (Table 2). Moreover, Δ/Λ-1 and Δ/Λ-2 also displayed significant inhibition of Topo II activity. The results of concentration-dependent Topo II inhibition assay of Δ/Λ-1 and Δ/Λ-2 are shown in Fig. 9. IC50 values of these complexes (IC50 ≈ 10, 10, 10 and 12μM, respectively) are quite comparable with some classical topoisomerase inhibitors (Table 2). Topoisomerases are nuclear enzymes that regulate DNA topology. Topo I acts by making a transient break in one DNA strand allowing the DNA helix to swivel, while Topo II makes transient breaks in both strands of one DNA molecule allowing the passage of another DNA duplex through the gap. Therefore, both Topo I and Topo II regulating DNA topology contained a DNA cleavage–religation process. Here we

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Fig. 8. Effects of different concentrations of Δ/Λ-1 (a) and Δ/Λ-2 (b) on the activity of DNA Topo I. Form I: supercoiled DNA; form II: nicked DNA.

performed the DNA religation assay for topoisomerases taking advantage of the fact that the enzyme can rejoin cleaved nucleic acids but cannot mediate DNA scission at suboptimal temperatures (0 °C) [65]. Within 60 min, both Topo I and Topo II showed a trend of supercoiling the relaxed plasmid and could religate the cleaved DNA to some extent (Fig. S5 and S6). It also proved that the rate of topoisomerase-catalyzed DNA supercoiling in the presence of EB was much higher than the rate without it [66]. Drug that block the overall catalytic activity of topoisomerases can be divided into two classes. Topoisomerase poisons trap the Topo– DNA cleavage complex, generating high levels of DNA breaks, such as CPT and VP-16. The other class is topoisomerase inhibitors, which inhibit the overall catalytic activities without inducing DNA breaks, such as mervarone and bisdioxopiperazines. These two classes of drugs have different mechanisms of action as well as their clinical schemes [2]. To determine whether the complexes interfered with DNA relaxation reaction by inhibiting topoisomerase catalysis or functioned as topoisomerase poisons, the mechanism of Ru(II) complexes was preliminarily studied based on the DNA religation assay. As shown in Fig. 10, with the presence of different Ru(II) complexes, both Topo I and Topo II could no longer religate the cleaved DNA in 60 min, suggesting that all the Ru(II) complexes in this work might have acted as novel dual Topo I/II poisons which captured protein-linked DNA breaks. 3.6. Fragmentation of nuclear DNA by comet assay The inhibition of the religation step during the processing of DNA by topoisomerases is believed to be the molecular basis of the antitumor activity of their inhibitors. Stabilization of cleavable complexes by Table 2 Inhibitory effects of four Ru(II) complexes on topoisomerase I and II activities. Drug

Camptothecin Doxorubicin Novobiocin Etoposide Hoechst 33258 Topostatin Δ-1 Λ-1 Δ-2 Λ-2

Inhibitory activity (IC50)/μM Topoisomerase I

Topoisomerase II

17 N100 N100 N1000 30 17 8 8 8 10

N100 1 32 35 35 4 10 10 10 12

Ref.

[62] [62] [62] [63] [64] [62] This This This This

topoisomerase inhibitors can lead to an inhibition of cell division and to cell killing. The alkaline single cell gel electrophoresis gives an image of the changes that have occurred in the chromatin organization in a single cell, which is considered a more accurate way of detection of early nuclear changes in a cell population [67]. As shown in Fig. 11, exposure of the HeLa cells to known TopoI/II poisons, CPT (50 μM) and VP-16 (50 μM) induced significant DNA strand breaks. Treatment with 50 μM Δ-1 and Δ-2 led to the appearance of a “broom-like” tail indicating severe DNA damage. Δ-1 and Δ-2 were able to induce DNA strand breaks in the manner similar to well-known TopoI/II poisons, CPT and VP-16, which corroborated the results from topoisomerase inhibition and DNA religation assay, suggesting that Δ-1 and Δ-2 may be dual topoisomerase I/II poisons. 3.7. Apoptosis induction The expression phosphatidylserine (PS) was chosen as the target process to identify apoptotic cells. PS is a simple anionic phospholipid that is normally restricted to the inner leaflet of the plasma membrane [68]. The redistribution of PS from the inner to the outer leaflet of the plasma membrane has been found to be a general feature of apoptosis, occurring before membrane bleb formation and DNA degradation [69]. Annexin V, a calcium-dependent phospholipid-binding protein with a high affinity for PS was used to detect early stage apoptosis [70]. Fig. 12 shows Alexa Fluor® 488 annexin V staining in HeLa cells treated for 24h with different concentrations of Δ-1 and Δ-2. The result showed a concentration dependent increase in annexin V binding in HeLa cells. Δ-1 at 25 μM showed a distinct shift in annexin fluorescence intensity. The mean fluorescence intensity difference between control and Δ-1 treated cell was significant at 50 and 100 μM. Δ-2 exhibited similar results as that described for Δ-1 above. The cell deaths induced by Δ-1 and Δ-2 were associated with the externalization of PS in the outer surface of the plasma membrane suggesting that Δ-1 and Δ-2 induced cell death mainly through apoptosis. 3.8. Cell cycle analysis

work work work work

The effect of complex Δ-1 and Δ-2 on cell cycle was investigated by the fluorescence-activated cell sorting (FACS) analysis of the DNA content. The cell cycle progression was analyzed at 25 μM concentration of Δ-1 and Δ-2 for 12, 24, 36 and 48 h (Fig. 13). It is well known that in this analysis, the cells in the G0/G1 phase have unreplicated diploid (2n) DNA content, whereas the G2/M phase has replicated ploid (4n) DNA. Also, hypodiploidy (b2n) DNA content at the sub-G1 phase and

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Fig. 9. Effects of different concentrations of Δ/Λ-1 (a) and Δ/Λ-2 (b) on the activity of DNA Topo II. Form I: supercoiled DNA; form II: nicked DNA.

Fig. 10. Effects of Ru(II) complexes on DNA religation mediated by Topo I and Topo II at different time points. (a) Topo I group: DNA + Topo I at different time points,Δ-1 group: DNA + Topo I + Δ-1 (15 μM) at different time points,Δ-2 group: DNA + Topo I + Δ-2 (15 μM) at different time points; (b) Topo II group: DNA + Topo II at different time points,Δ-1 group: DNA + Topo II + Δ-1 (15 μM) at different time points, Δ-2 group: DNA + Topo II + Δ-2 (15 μM) at different time points. Form I: supercoiled DNA; form II: nicked DNA.

replication in the S phase are observable. Upon exposure of the cells to Δ-1, the percentages of cells decreased in the G0/G1 phase of the cell cycle (52.37 to 42.17%) but increased in the S phase of the cell cycle (29.19 to 46.63%) with the increment of time. For Δ-2, under the same experimental condition, parallel results could been seen, the percentages of cells decreased in the G0/G1 phase of the cell cycle (53.72 to 44.27%) and also increased in the S phase of the cell cycle (28.84 to 44.27%). The only distinction was that with increasing treatment time the values of sub-G1 varied from 0% to 39.46% for Δ-1 but from 0% to 23.83% for Δ-2. And the differences were in accord with their cytotoxicity. Compared with the untreated control, Δ-1 and Δ-2 disturb cell cycle strongly arrested at S phases and induced an obvious increase in sub-G1.

All of these observations revealed that the major mode of cell death induced by Δ-1 and Δ-2 was apoptosis, as the accumulation of hypodiploid cells in the sub-G1 phase is considered to be a marker for apoptotic cell death [71]. Several intracellular cascades such as activation of caspases, disruption of normal mitochondrial function, or both would be involved in the road map towards the final destination, that is, the apoptotic cell death [47]. 4. Conclusions In summary, two pairs of chiral Ru(II) complexes Δ/Λ[Ru(bpy)2(pscl)]2 + and Δ/Λ-[Ru(bpy)2(psbr)]2+ have been synthesized

Fig. 11. Drug-induced DNA strand breaks in HeLa cells. HeLa cells were treated with different concentrations of drugs for 24 h. (a) Control; (b) Δ-1 (50 μM); (c) Δ-2 (50 μM); (d) CPT (50 μM); (e) VP-16 (50 μM).

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25

Fig. 12. Δ-1 and Δ-2 induced apoptotic cell death as examined by the Alexa Fluor® 488 annexin V assay. HeLa cells were treated with different concentrations of Δ-1 and Δ-2 for 24 h. (a) In the presence of Δ-1 (25 μM); (b) in the presence of Δ-1 (50 μM); (c) in the presence of Δ-1 (100 μM); (d) in the presence of Δ-2 (25 μM); (e) in the presence of Δ-2 (50 μM); (f) in the presence of Δ-2 (100 μM). Unfilled curve, cultures treated with vehicle (0.1% DMSO); filled curve, cultures treated with Ru(II) complexes.

and characterized. The two enantiomers showed different antitumor activities to HeLa, MCF-7, HepG2 and BEL-7402 tumor cells which was in accord with the results of cellular uptake. Moreover, we studied the cellular distribution Ru(II) using a fluorescence microscope and we found that these Ru(II) complexes gradually penetrate into the interior of the nucleus, suggesting that nucleic acids were the cellular target of these enantiomers. The data from UV–vis absorption titration and luminescence titration measurements all supported that Ru(II) complexes could bind to DNA base pairs. Topoisomerase inhibition and DNA religation assay confirmed that four Ru(II) complexes acted as efficient dual poisons of topoisomerases I and II. These results demonstrated that Δ/Λ-1 and Δ/Λ-2 bind to DNA and stabilize the TopoI/II–DNA cleavable complex. DNA strand breaks had also been observed from alkaline single cell gel electrophoresis (comet assay). According to results from Alexa Fluor® 488 annexin V staining assays and flow cytometry analysis, Δ-1 and Δ-2 inhibited the growth of HeLa cells through the induction of apoptotic cell death. The results demonstrated that Δ/Λ-[Ru(bpy)2(pscl)]2+ and Δ/Λ-[Ru(bpy)2(psbr)]2+ acted as dual poisons of topoisomerases I and II and caused DNA damage that could lead to cell cycle arrest and/or cell death by apoptosis. We hope the results to be of value in further understanding the DNA-binding and topoisomerase inhibition by Ru(II) complexes, as well as laying the foundation for the discovery of new antitumor agents. Abbreviations bpy 2,2′-bipyridine phen 1,10-phenanthroline pscl 2-(5-chlorothiophen-2-yl)imidazo[4,5-f][1,10] phenanthroline psbr 2-(5-bromothiophen-2-yl)imidazo[4,5-f][1,10] phenanthroline bfipH 2-(benzofuran-2-yl)imidazo[4,5-f][1,10]phenanthroline ipad 2-(anthracene-9,10-dione-2-yl)imidazo[4,5-f][1,10] phenanthroline

dppz Topo MTT

dipyridophenazine topoisomerases 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide DAPI 4′,6-diamidino-2-phenylindoleand CT-DNA calf thymus DNA EB ethidium bromide CPT camptothecin VP-16 etoposide PI propidium iodide PBS phosphate buffered saline DTT dithiothreitol BSA bovine serum albumin

Acknowledgments This work was supported by the National Science of Foundation of China (Nos. 21071155, 21172273, 21171177), the National High Technology Research and Development Program of China (863 Program, 2012AA020305), the National Science of Foundation of Guangdong Province (9351027501000003), the Research Fund for the Doctoral Program of Higher Education (20110171110013), State Key Laboratory of Optoelectronic Materials and Technologies (2010-ZY-4-5) and Sun Yat-Sen University. Zai-Li Peng thanks the NSFC (J1103305) for their support. Xiao-Juan Hou thanks the Science and Technology Department of Hunan Province (2010FJ4090) for their support.

Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.jinorgbio.2013.09.015.

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Fig. 13. Δ-1 induced apoptotic cell death as examined by flow cytometric analysis. Cells were treated with Δ-1 and Δ-2 at 25 μM for 12, 24, 36 and 48 h. (a)–(d) Control for 12, 24, 36 and 48 h; (e)–(h) Δ-1 for 12, 24, 36 and 48 h; (i)–(l) Δ-2 for 12, 24, 36 and 48 h.

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