Synthesis, pH-induced “on–off–on” luminescence switching, and partially intercalative DNA-binding and DNA photocleavage properties of an β-d -allopyranoside-grafted ruthenium(II) complex

Journal of Inorganic Biochemistry 113 (2012) 66–76

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Synthesis, pH-induced “on–off–on” luminescence switching, and partially intercalative DNA-binding and DNA photocleavage properties of an β-D-allopyranoside-grafted ruthenium(II) complex Xiao-Long Zhao, Yan-Zi Ma, Ke-Zhi Wang ⁎ College of Chemistry, Beijing Normal University, Beijing 100875, P.R. China

a r t i c l e

i n f o

Article history: Received 11 February 2012 Received in revised form 28 March 2012 Accepted 29 March 2012 Available online 6 April 2012 Keywords: Ruthenium DNA Luminescence switch pH DNA photocleavage Singlet oxygen

a b s t r a c t A new ruthenium(II) complex [Ru(Happip)3](ClO4)2 {Happip = 2-(4-(β-D-allopyranoside)phenyl)imidazo [4,5-f][1,10]phenanthroline} was synthesized and characterized by elemental analysis, 1H NMR and matrixassisted laser desorption ionization mass spectrometry. Calf-thymus DNA-binding properties were studied by DNA viscosity measurements and spectroscopic methods of UV-visible (UV-vis) absorption and luminescence titrations, steady-state emission quenching, DNA competitive binding with ethidium bromide and DNA melting experiments, indicating that the complex partially intercalates into the DNA with a large binding constant greater than 10 6 M− 1. The pH effects on UV-vis absorption and emission spectra of the complex were studied, demonstrating that the complex acted as an excellent pH-induced “on–off–on” luminescence switch with large on–off intensity ratios of 88 and 50 with one of luminescence on/off switching actions occurring in near-physiological pH region (pKa2 = 7.33). The DNA photocleaving properties of [Ru(Happip)3] 2+ were also studied and compared with those of [Ru(bpy)2(Happip)] 2+ and [Ru(bpy)2(Hpip)] 2+{bpy = 2,2′-bipyridine, Hpip = 2-(4-phenyl)imidazo[4,5-f][1,10]phenanthroline}. © 2012 Elsevier Inc. All rights reserved.

1. Introduction Recent emphasis has been placed on developing transition metal complex-based reagents capable of structurally probing DNA and cleaving DNA, applicable as DNA footprinting and anticancer agents [1–7]. In this regard, ruthenium(II) polypyridyl complexes were extensively studied owing to their tunable photophysical, photochemical and redox properties [8–17]. A lot of ruthenium(II) complexes with extended π organic ligands were reported to be classic DNA intercalators, and such structure–property relationships as the effects of the charge, planarity of the intercalative moiety, ancillary ligand, the formation of intramolecular hydrogen bond, and the populations and energies of lowest unoccupied molecular orbital (LUMO) and/or some virtual orbitals near LUMO on the DNA binding properties have been well revealed [18]. But much less is known about structure–property relationships for grooving [18,19] and partially intercalative DNA binders [20–22], due mostly to the fact that a very limited number of these two families of DNA binders have been reported. The groove binding of Ru(II) complex to the DNA could be very strong, and could result in evident spectral changes. For example, we have recently observed a 26-fold emission enhancement as [Ru(phen)2(cdpq)]+ {phen=1,10-phenanthroline, cdpq=deprotonated

⁎ Corresponding author. Tel.: + 86 10 58805476; fax: + 86 10 58802075. E-mail address: [email protected] (K.-Z. Wang). 0162-0134/$ – see front matter © 2012 Elsevier Inc. All rights reserved. doi:10.1016/j.jinorgbio.2012.03.010

2-carboxyldipyrido[3,2-f:2′,3′-h]quinoxaline} groove-bound to DNA [18]. The DNA binding constant values could attain to as high as 2.0 × 10 5 to 2.9 × 10 6 M − 1 equivalent to those of high-affinity metallointercalators, as recently reported by Thomas and Das et al. for groove binder [Ru(bpy)2(mbeb)] 2+ {bpy= 2,2′-bipyridine, mbeb = 4[(1E)-2-(4′-methyl[2,2′-bipyridin]-4-yl)ethenyl]-1,2-benzenediol} [19]. [Ru(bpy)2(dppz)] 2+ {dppz = dipyrido[3,2-a:2′,3′-c]phenazine} and [Ru(phen)2(dppz)] 2+ are the most interesting classical intercalators which are termed as the DNA molecular light switches due to their absence of luminescence in the water buffer, but greatly enhanced photoluminescence in the presence of DNA [23,24]. DNA cleavage is a critical event for gene mutation and cancer genesis in biological systems as well as DNA binding [25]. The observation of DNA photocleavage by ruthenium(II) polypyridyl complexes, such as [Ru(bpy)3] 2+ and [Ru(phen)3] 2+, can be traced back to 20 years ago [26,27]. Since then, many ruthenium complexes have been investigated as effective DNA photocleavers [28–35] and potential photodynamic therapy agents [36]. The DNA was cleaved by radicals or radical ions generated via electron transfer or hydrogen abstraction processes (anaerobic type I mechanisms), or reactive oxygen species generated by energy transfer from excited Ru(II) complex to dioxygen (highly oxygen-dependent type II mechanisms) [37]. The reactive oxygen species such as singlet oxygen, hydroxyl radical and superoxide anion radical could play important roles via type II mechanisms in the Ru-based DNA cleavers, and in particular singlet oxygen generated through energy transfer from the excited

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

state of the complex to dioxygen is regarded to be a more crucial component that contributes to the cleavage [29–32,34,35,38,39]. The DNA photocleavage by [Ru(bpy)3] 2+ was known to arise from the oxidative damage to the deoxyribose backbone by the reactive oxygen species, and the quantum yield of photosensitized formation of singlet oxygen by [Ru(bpy)3] 2+ was reported to be 0.81 [32,39,40]. However, the low quantum yield of singlet oxygen formation by [Ru(bpy)2(dppz)] 2+ (0.09) in water due to its rapid radiationless decay rate of excited triplet state and in turn its low photoluminescent yield, resulted in its inefficacy in cleaving DNA in spite of its high binding affinity to DNA [31,32,35]. The bright luminescence of the Ru(II) complex we studied in this paper prompted us to examine its singlet oxygen formation yield and DNA photocleavage properties. On the other hand, ruthenium complexes have attracted considerable attention as pH sensors in recent years due to their relative long excited-state lifetimes, high luminescence quantum yields, and rather sensitivity of their ground- and excited state properties to environmental pHs [8–13,41–45]. There has been increasing need for monitoring pH values in many chemical and biological processes, and clinical and environmental-protection analyses [46]. Protonation/ deprotonation can be regarded as one of the simplest but very important chemical or even biochemical processes. Many life processes, such as enzymes, only work within a very narrow pH range, and their activity can be described as pH responsive “on–off” switches [47]. Attempts to mimic such switching behavior by constructing luminescence devices that are modulated by pH are thus of great interest. In addition, pH luminescence sensing/switching systems have been proposed as logical operators to perform molecular computing [48,49]. Although many ruthenium complexes have been documented to have interesting on/off or off/on luminescence switching properties with large emission enhancement factors [9–12], to the best of our knowledge, only a limited number of multi-state “on–off–on”, “off– on–off” type pH-induced emission switches have so far been reported [9,43–45]. Ji's [43–45] and our groups [9,50,51] reported some imidazole-containing Ru(II) complex-based multi-state pH luminescence switches, but luminescence on/off intensity ratios that were usually less than 70, need further improvements. As part of our ongoing studies aimed at DNA binding and cleavage properties as well as sensitive pH luminescent sensors or switches based on Ru(II) complexes, we presented here the synthesis, characterization, DNA binding and photocleavage properties, and pH luminescence multi-state switching of a novel ruthenium complex grafted with a biologically and physiologically active β-D-allopyranoside. β-DAllopyranoside is a main part of 4-formylphenyl-β-D-allopyranoside which is called helicidum. In China, helicidum is used in clinic in the treatment of neurasthenia and vascular headache with high efficiency and limited side effects and toxicity, and synthetic efforts for chemically modifying helicium have recently evidenced to show improved sedative activity [52,53]. Here, a polypyridyl ruthenium(II) complex carrying with helicidium, [Ru(Happip)3] 2+ {Happip = 2-(4(β-D-allopyranoside)phenyl)imidazo[4,5-f][1,10]phenanthroline}, was demonstrated to be a DNA partial intercalator and an excellent “on– off–on” luminescence switch with on/off emission intensity ratios of 88 and 50 with the former switching action occurring near physiological pH region.

67

{Hpip =2-(4-phenyl)imidazo[4,5-f][1,10]phenanthroline}, was synthesized as detailed below. 2.2. Synthesis of [Ru(Happip)3](ClO4)2 A solution of RuCl3•2H2O (7.9 mg, 0.03 mmol) in methanol (5 mL) was treated with AgNO3 (15.3 mg, 0.09 mmol) for 2 h and filtered to remove the AgCl precipitate formed. To the filtrate was added Happip (43.0 mg, 0.09 mmol) and glycol (5 mL). The reaction mixture was microwave heated at 160 °C for 2 h under nitrogen. After cooling to room temperature, most of the solvent was removed under reduced pressure. The crude product was purified by chromatography on silica gel with saturated aqueous NaNO3-N,N-dimethylformamidemethanol (1:5:200, v/v/v) as eluent followed by reprecipitation with saturated NaClO4 aqueous solution. Yield: 18.0 mg, 32%. IR (KBr, cm− 1): 3396, 2921, 1655, 1612, 1480, 1454, 1385, 1365, 1240, 1182, 1080, 1039, 843, 807, 718, 622. 1H NMR (400 MHz, DMSO-d6): 14.22 (s, 3 H, Hq), 9.06 (d, J = 8.1 Hz, 6 H, Ha), 8.26–8.16 (dd, J1 = 8.4 Hz, J2 = 8.7 Hz, 6 H, Hc), 8.03 (d, 6 H, J2 b 1.9 Hz, Hd), 7.80 (m, 6 H, Hb), 7.28–7.03 (dd, J1 = 8.6 Hz, J 2 = 8.7 Hz, 6 H, He), 5.28 (d, J = 7.2 Hz, 3 H, Hf), 5.11 (d, J = 6.7 Hz, 3 H, H°), 5.01 (d, J = 3.5 Hz, 3 H, Hn), 4.70 (d, J = 7.2 Hz, 3 H, Hp), 4.51 (t, J1 = 4.9 Hz, J2 = 1.9 Hz, 3 H, Hm), 3.97 (d, J = 2.9 Hz, 3 H, Hj), 3.87 (m, 6 H, Hg,k), 3.46 ppm (m, 9 H, Hh,i,l). Anal. Calcd for C75H66Cl2N12O26Ru•8H2O: C, 48.24; H, 4.43; N, 9.00. Found: C, 48.29; H, 4.79; N, 9.19. Anal. Calcd for matrix-assisted laser desorption ionization mass spectrum m/z: 762.18 ([M-2ClO4−] 2+). Found: 762.17 ([M-2ClO4−] 2+). Caution: Perchlorate salts of metal complexes with organic ligands are potentially explosive, and only small amounts of the material should be prepared and handled with great care. 2.3. Instrumentations and methods Instrumentations and methods for spectroscopic measurements and nucleic acid binding studies are same as before [8–14], and shown in the Supplementary Materials. 2.4. DNA photocleavage experiments The photoinduced DNA cleavage by ruthenium(II) complex was examined by gel electrophoresis. Supercoiled pUC 18 DNA (0.2 μg) was treated with the ruthenium(II) complex in the buffer (50 mM Tris–HCl, 18 mM NaCl, pH = 7.2), and then the solution was irradiated at room temperature with UV light (360 nm, 16 mW/cm 2) for 2 h after incubation in the dark for 1 h. The samples were analyzed by electrophoresis for 1 h at 80 V on a 0.8% agarose gel in TAE buffer. The gel was stained with 1 μg/mL ethidium bromide and photographed under UV light. The percentage of cleavage (C) was calculated according to Eq. (1): C¼

DII þ 2DIII DI þ DII þ 2DIII

ð1Þ

where DI, DII and DIII are the integrated density values of Form I (supercoil form), Form II (nicking form) and Form III (linear form), respectively.

2. Experimental section

2.5. Quantum yield of 1O2 generation

2.1. Materials

The reaction of 1O2 with 1,3-diphenylisobenzofuran (DPBF) was adopted to measure the quantum yields of 1O2 generation for ruthenium(II) complex. A 3 mL of air-saturated methanol solution containing DPBF (20 μM) and a complex (5 μM), of which the absorbance at 480 nm originating from the absorption of complex was obtained from the UV-vis spectra, was charged into an open 1 cm path fluorescence cuvette and illuminated with light of 480 nm (obtained from a Shimadzu RF-5301PC fluorescence spectrophotometer, 3 nm of

1,10-Phenanthroline-5,6-dione [54], cis-Ru(bpy)2Cl2•2H2O [55] and Happip [56] were prepared according to the literature methods. The other chemicals were obtained from commercial sources and used without further purification. [Ru(Happip)3](ClO4)2 {see Scheme 1 for molecular structure along with those for analogous complexes of [Ru(bpy)2(Happip)]2+, [Ru(bpy)2(Hpip)]2+ and [Ru(Hpip)3]2+}

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Scheme 1. Molecular Structures of Happip, [Ru(bpy)2(Happip)]2+, [Ru(bpy)2(Hpip)]2+, [Ru(Hpip)3]2+, and [Ru(Happip)3]2+ with proton-numbering scheme for 1H NMR assignments.

excitation slit width). The consumptions of DPBF were followed by monitoring its fluorescence-intensity decrease at the emission maximum (λex = 405 nm, λem = 479 nm) at different irradiation times [32,39]. [Ru(bpy)3] 2+ was used as standard, whose 1O2 generation quantum yield was taken to be 0.81 in air saturated methanol [32,39]. 3. Results and discussion 3.1. DNA binding studies 3.1.1. UV-visible(UV-vis) absorption spectra The absorption spectra of [Ru(Happip)3] 2+ in neutral aqueous solution mainly consists of two well-resolved bands in the range 200–600 nm. The peak at 287 nm is assigned to the intraligand (IL) π–π* transition, and the lowest energy band centered at 474 nm with a shoulder peak at 440 nm is attributed to the metal-to-ligand charge-transfer (MLCT) transition, on comparisons with π–π* absorption bands at 284, 283 and 282 nm, and MLCT bands at 461, 458 and 465 nm reported for analogous complexes of [Ru(bpy)2(Happip)] 2+ [56], [Ru(bpy)2(Hpip)] 2+ [57] and [Ru(Hpip)3] 2+ [58], respectively. Obviously, the fusion of allopyranoside group to [Ru(Hpip)3] 2+ induce a 9-nm bathchromic shift. The absorption spectra of [Ru(Happip)3] 2+ in the absence and presence of calf thymus DNA are shown in Fig. 1. Upon successive additions of the DNA, the MLCT band of the complex at 474 nm exhibited bathochromism of 5 nm and pronounced hypochromism H% {H% = 100 (Afree − Abound)/Afree} of 56% at a concentration ratio of [DNA]/[Ru] ≈ 10 at saturated binding, which is different from the unaffected spectral behavior previously observed for [Ru(Hpip)3] 2+ [58] and even significantly greater than the corresponding hypochromism values of ~ 20% for proven DNA intercalators of [Ru(bpy)2(Happip)] 2+ [56] and [Ru(bpy)2(Hpip)] 2+ [57]. In order to elucidate the binding strength of the complex to DNA, the intrinsic binding constant Kb was determined by monitoring the changes of absorbance at the MLCT band according to Eq. (2) [59]: ½DNA=ðε a –εf Þ ¼ ½DNA=ðεb –εf Þ þ 1=½K b ðεb –εf Þ

ð2Þ

where [DNA] is the concentration of DNA in base pairs, εa is the apparent absorption coefficient of the Ru(II) complex in the presence of the DNA, which was obtained by calculating Aabs/[Ru], and εf and εb are the extinction coefficients for the free ruthenium complex and the ruthenium complex in the fully bound form, respectively. In a plot of [DNA]/(εa − εf) versus [DNA], Kb is given by the ratio of the slope to the y intercept. As illustrated in the inset a of Fig. 1, an intrinsic DNA-binding constant of (1.00 ± 0.16) × 106 M− 1 was obtained, which is greater than the Kb values of (5.24 ± 0.8) × 105 M − 1 and 4.7 × 105 M − 1 for [Ru(bpy)2(Happip)] 2+ [56] and [Ru(bpy)2(Hpip)]2+ [60]. The Kb value of the complex could also be calculated by nonlinear regression analysis using Eqs. (3) and (4) [61]:  ðεa –εf Þ=ðεb –ε f Þ ¼

 1=2  2 2 b– b –2K b C t ½DNA=n =ð2K b C t Þ

b ¼ 1 þ K b Ct þ K b ½DNA=2n

ð3Þ ð4Þ

where εa, εf and εb have the same definition as in Eq. (2), Ct is the total ruthenium complex concentration, [DNA] is the DNA concentration in nucleotides, and n is the binding site size. As shown in the inset b of Fig. 1, n = 0.41 ± 0.01 and Kb = (1.49 ± 0.07) × 106 M− 1 were derived by monitoring the decay of the absorbance at 474 nm. This Kb value is in good agreement with that derived according to Eq. (2), and indicates the avid DNA-binding affinity of the complex which may be due to the potential hydrogen-bonding interaction between the hydroxyl of the allopyranoside and the DNA. 3.1.2. Emission spectra The emission spectral changes of [Ru(Happip)3] 2+ upon increasing DNA concentrations are shown in Fig. 2. In contrast to DNAinduced emission intensity enhancement previously reported for [Ru(bpy)2(Happip)]2+ and [Ru(bpy)2(Hpip)]2+, an almost unaffected luminescent behavior of [Ru(Hpip)3]2+ [56–58], the spectral changes of [Ru(Happip)3]2+ are dissected into two opposite processes, namely a DNA-induced on–off–on emission switching was observed. The

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

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Fig. 1. UV-vis spectra of [Ru(Happip)3]2+ (4 μM) in the absence and presence of increasing DNA concentrations (0–40 μM). Inset: plots of [DNA] / (εa − εf) vs. [DNA] and the linear fit (a), and (εa − εf) / (εb − εf) vs. [DNA] and the nonlinear fit (b).

luminescent intensities of complex at 597 nm were reduced by 74% along with a red shift of 5 nm upon increasing DNA concentrations to [DNA]/ [Ru]=0.83 (Fig. 2a), and was instead enhanced by 4.5 folds with a 8 nm hypsochromic shift as the DNA concentrations were further increased from [DNA]/[Ru]=0.83 to [DNA]/[Ru]=10 (Fig. 2b). This DNA driven on–off–on emission switching behavior is unusual as compared with two-stage UV-vis spectral changes and monotone “turn on” emission we recently observed for the binding of [Ru(bpy)2(btppz)]2+ {btppz = benzo[h]tripyrido[3,2-a:2′,3′-c:2′′,3′′-j]phenazine} to the DNA [8]. The transition from “turn off” to “turn on” emission observed above for the DNA binding of [Ru(Happip)3]2+ is indicative of changes in DNA binding modes, probably from electrostatic/hydrogen bonding to non-covalent intercalation or groove binding, since the hydrophobic environment inside the DNA helix reduces the accessibility of water molecules to the complex and the complex mobility is restricted at the binding site, leading to decreased vibrational relaxation in the excited state [62]. The steady-state emission quenching experiments using [Fe(CN)6] 4as quencher were also used to assess the binding of ruthenium with DNA. [Fe(CN)6] 4- could very efficiently quench the emission of the ruthenium complex which is free in solution but poorly quench the emission of the DNA tightly bound ruthenium complex, due to the high repulsion between the highly anionic [Fe(CN)6] 4- and the negative DNA phosphate backbone. As illustrated in Fig. S1, in the absence of DNA, the luminescence of [Ru(Happip)3] 2+ was efficiently quenched by [Fe(CN)6]4 − as described by a linear Stern–Volmer plot with a large Stern–Volmer constant Ksv of 1.36 × 106 M− 1 according to equation: I0/I = 1 + Ksv [Q], where I0 and I are the emission intensities in the absence and presence of [Fe(CN)6] 4-, respectively, and [Q] is the concentration of [Fe(CN)6] 4-. In contrast, addition of DNA decreased the quenching efficiency of [Fe(CN)6]4- significantly with a much less quenching constant Ksv of 7.3×103 M− 1. A ratio of the Ksv value in the absence to that in the presence of DNA, R, was found to be 186 for [Ru(Happip)3]2+, which is comparable to 178 for [Ru(bpy)2(Happip)]2+ [56] and about 8-fold that for [Ru(bpy)2(Hpip)] 2+ [63], indicating that [Ru(Happip)3] 2+ was effectively protected by the DNA from accessibility of [Fe(CN)6] 4-. For an alternative assessment of the DNA-binding affinity and mode of the complex, the ethidium bromide (EB) competitive binding study was carried out. As a typical indicator of intercalation [64], EB emits intense fluorescence in the presence of DNA due to strong

Fig. 2. Changes in emission spectra of [Ru(Happip)3]2+ (4 μM) upon successive additions of DNA (0–3.33 μM) (a) and (3.33–40 μM) (b).

intercalation of the phenanthridinium ring between the adjacent base pairs. An intercalative complex can usually almost fully displace EB from DNA-bound EB, and the fluorescence of EB–DNA solution will be quenched due to the fact that free EB molecules are readily quenched by the surrounding water molecules [65,66]. Previous reports indicated that EB has two DNA binding modes, primarily intercalatively and partially groove bound to the DNA [67–71]. EB was also reported to intercalate into DNA through interactions with the minor groove of the DNA, the displacement of EB by the titration of a drug is thus suggestive of an intercalative or minor groove binding. For example, the nonintercalating DNA groove binding agents of berenil, bisamidines, spermine and spermidine were reported to be capable of displacing EB from DNA, but the reduction of DNA-bound EB emission is usually modest for the addition of a groove binder [66]. However, non-intercalative drugs could displace EB from DNAbound EB as well, but not fully. We recently reported that only 40% of the DNA-bound EB molecules were displaced by the groove binder [Ru(phen)2(cdpq)] + [18]. DNA groove binding agents of berenil, bisamidines, spermine and spermidine were reported to be capable of displacing EB from DNA only modestly [66]. As shown in Fig. S2a, the addition of [Ru(Happip)3] 2+ to the EB–DNA system resulted in appreciable reduction in emission intensity at 602 nm, but only 75% DNA-bound EB was displaced, in contrast to almost entire displacement of the EB we previously observed for proven DNA intercalators of [Ru(bpy)2(bipp)](ClO4)2, [Ru(bpy)2(bopp)](ClO4)2 and [Ru(bpy)2(btpp)](ClO4)2 {bipp = 2-benzimidazoyl-pyrazino[2,3f][1,10]phenanthroline, bopp = 2-benzoxazolyl-pyrazino[2,3-f][1,10] phenanthroline, and btpp = 2-benzthiazolyl-pyrazino[2,3-f][1,10] phenanthroline} [51], similarly to the partial displacement (40% and 31.4%) of DNA-bound EB by groove binder of [Ru(phen)2(cdpq)]+ [18]

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and partial intercalator of [Ru(dmp)2(Hmip)]2+ {dmp = 2,9-dimethyl1,10-phenanthroline, Hmip = {2-(2,3-methylenedioxyphenyl)imidazo [4,5-f]1,10-phenanthroline} [21], indicating that [Ru(Happip)3] 2+ is a partial intercalator or groove binder rather than a classic intercalator. The quenching plot of I0/I vs. [Ru]/[DNA] (inset of Fig. S2a) is in good agreement with the linear Stern–Volmer equation of I0/I=1+KD[Ru]/ [DNA], and a Stern–Volmer constant KD was derived to be 1.73. A competitive binding model was used to calculate the apparent binding constants by using equation Kapp =KEB [EB]50%/[Ru]50% [65], where Kapp is the apparent DNA-binding constant of [Ru(Happip)3]2+, KEB is the DNA-binding constant of EB, and [EB]50% and [Ru]50% are the EB and [Ru(Happip)3]2+ concentrations at 50% EB displacement. In a plot of percentage of free EB (Fig. S2b), [EB]/([EB]+[EB–DNA]) vs. [Ru]/[EB], we can see that 50% of EB molecules were displaced from DNA-bound EB at a concentration ratio of [Ru]/[EB]=2.59. By using the average value of previously reported DNA-binding constants of EB, KEB =4.94×105 M− 1 [72], a Kapp value of 1.91×105 M− 1 of the complex was obtained. It is noteworthy that the KD and Kapp values of [Ru(Happip)3]2+ are smaller than KD =2.64 and Kapp = 2.31× 105 M− 1 we previously reported for [Ru(bpy)2(Happip)]2+ [56], indicating that the higher DNA-binding affinity of [Ru(bpy)2(Happip)]2+ than [Ru(Happip)3]2+ was derived from the EB competition experiment, which is opposite to the DNA affinity order {Kb ≈106 M− 1 for [Ru(Happip)3]2+ vs. Kb =(5.24± 0.8)×105 M− 1 for [Ru(bpy)2(Happip)]2+} derived from UV-vis spectroscopy [56]. This discrepancy may result from the difference in the DNA binding modes of [Ru(Happip)3] 2+ and EB, namely the partially intercalative or groove binding mode for [Ru(Happip)3] 2+ vs. primarily intercalative mode for EB. 3.1.3. Thermal denaturation experiments Thermal denaturation behaviors of DNA in the presence of complex can distinguish intercalative from electrostatic binding mode, and get insights into DNA binding thermodynamic parameters. It is well known that the double-stranded DNA gradually dissociates to single strands with increasing the temperature, and results in a hyperchromic effect at 260 nm due to the higher extinction coefficient of the single-strand form than the double-helical form at this wavelength [73,74]. The melting temperature Tm, which is defined as the temperature at which half of the total base pairs is unpaired, was used to characterize denaturation process of the DNA. According to the literature [75,76], intercalation of natural or synthesized organicand metallo-intercalators into the base pairs of DNA would stabilize the double strand of DNA and generally results in a considerably increased Tm, while electrostatic binders have little effects on Tm. The effects of additions of [Ru(Happip)3] 2+ on Tm of DNA is shown in Fig. 3. A Tm value of the DNA alone was found to be 67.1 °C, and to steadily increase upon successively increasing the complex concentrations (Fig. S3). The increase in Tm, ΔTm, was found to be 8.7 °C at a concentration ratio of [Ru]/[DNA] = 0.10, which is distinctly distinguishable from the electrostatic binder, e.g. b2 °C for [Ru(bpy)3] 2+ at [Ru]/ [DNA] = 0.10 [75], and falls into rather broad ΔTm range from 5.9 °C to 19.3 °C as listed in Table S1 for some DNA intercalators. The binding constant of complex [Ru(Happip)3] 2+ to DNA at Tm was determined by McGee's Eq. (5) [77]: 0

1=n

1=T m –1=T m ¼ ðR=ΔH m Þlnð1 þ KLÞ

standard enthalpy change ΔH0 of the binding of [Ru(Happip)3] 2+ to DNA was determined by van't Hoff's Eq. (6) [79]:   0 lnðK 1 =K 2 Þ ¼ ΔH =R ½ðT 1 –T 2 Þ=T 1 T 2  0

ΔGT ¼ –RTlnK 0

0

ΔGT ¼ ΔH –TΔS

ð6Þ ð7Þ

0

ð8Þ

where K1 and K2 are the DNA-binding constants of the complex at T1 and T2, respectively. The standard free energy change ΔGT0, and standard entropy change ΔS 0 of the binding reaction were calculated according to Eqs. (7) and (8). By using K1 = 1.49 × 106 M− 1 (T1 = 298 K) and 0 K2 = 2.10 × 10 5 M− 1 (T2 = 349 K), the values of ΔH0, ΔG298K and ΔS 0 −1 at 25 °C were derived to be –33.33 kJ mol , –35.22 kJ mol− 1 and 0 6.34 J mol− 1 K − 1. The negative ΔG298K and ΔH0 values imply that the sum of the free energies and enthalpies of the complex and the DNA are higher than that of their adduct, and the binding reaction was driven enthalpically. The positive ΔS 0 value implies the higher entropy of the complex-DNA-binding adduct than the free DNA and the complex, which also benefit to the binding reaction. 3.1.4. Viscosity measurements The optical measurements addressed above only afford necessary but not conclusive evidences for DNA binding mode, helix lengthdependent DNA viscosities that are considered to be unambiguous evidences the for DNA binding mode, were measured. It is well accepted that a classical intercalation mode results in lengthening the DNA helix, as base pairs are separated to accommodate the binding ligand, leading to increase of DNA viscosity. The groove or electrostatic binding typically causes less-pronounced or no change in the DNA solution viscosities. A partial intercalation of a drug into DNA could bend or kink the DNA helix, reducing its effective length and its viscosity concomitantly [20–22]. The effects of [Ru(Happip)3] 2+ and EB on the viscosity of the DNA are shown in Fig. 4. As expected, EB increased the relative specific viscosity of the DNA due to lengthening of the DNA double helix resulting from intercalation. On the contrary, the increases in [Ru(Happip)3] 2+ concentrations were found to greatly decrease the relative viscosities of the DNA, in contrast to behaviors of proven DNA intercalators of [Ru(bpy)2(Happip)] 2+ [56] and [Ru(bpy)2(Hpip)]2+ [63], suggesting a partially intercalative binding mode to DNA. This may be related to the molecular structure of the complex. For [Ru(Happip)3] 2+, due to the steric hindrance of other two large Happip ligands, the Happip ligand is prevented from completely intercalating into the DNA. The partial intercalation may act as a “wedge” to pry apart one side of a base-pair stack apart, as

ð5Þ

0 where Tm and Tm are the melting temperatures of DNA alone and in the presence of complex, respectively, ΔHm is the enthalpy of DNA melting (ΔHm = 6.9 kcal mol − 1) [78], R is the gas constant, L is the free ruthenium complex concentration (approximated by the total complex concentration at Tm), and n is the binding site size. By taking n = 0.41 bp (bp = base pairs, approximated by the n value at 298 K) obtained from the absorption spectral titration data, K was determined to be 2.10 × 105 M − 1 at 75.8 °C, indicating that the complex still displays strong DNA-binding affinity at the melting point of DNA. The

Fig. 3. Thermal denaturation curves of the DNA ([DNA] = 53.0 μM) at different [Ru(Happip)3]2+ concentrations of [Ru]/[DNA]= 1/10, 1/20, 1/30, 1/40, 1/50 and DNA alone.

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

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observed for Δ-[Ru(phen)3] 2+ [78,80], but not fully separate the stack as required by the classical intercalation mode, causing a static bend or kink of the DNA strand, and a decrease in the viscosity of the DNA. This behavior is in contrast to that of analogous [Ru(Hpip)3] 2+, for which a primarily weak electrostatic binding mode to the DNA was supposed [58], since its absorption and emission spectra were almost unaffected by the presence of the DNA, indicating that allopyranoside group has a significant effect of the DNA binding affinity and binding mode, due most probably to the presence of hydrogen bonding between the phosphate backbone with the allopyranoside group in [Ru(Happip)3] 2+. 3.2. pH effects on absorption and luminescence spectra 3.2.1. UV-vis absorption spectra The UV-vis spectral pH titrations of [Ru(Happip)3]3+ were carried out over the pH range of 0–13.7. The spectral changes shown in Fig. 5 indicate that the complex underwent three ground-state protonation/deprotonation processes over the pH range studied. Upon increasing pH from 0 to 2.1 (Fig. 5a), the absorption band intensities centered at 282, 304, 428 and 457 nm decreased obviously, accompanying with appearance of three isosbestic points at 349, 387 and 471 nm, and a bathochromic shift of 5 and 9 nm for the 282 and 457 nm bands, respectively. These spectral changes observed were due to the dissociation of three protons on the protonated imidazole rings. Upon further increasing pH from 2.5 to 9.0 (Fig. 5b), the second deprotonation step took place, which was assigned to the deprotonation of two protons on the two neutral imidazole rings. The bands at 287 and 476 nm decreased sharply by 59% and 41%, and the band at 476 nm was red-shifted by 6 nm with appearance of an isosbestic point at 497 nm. The third deprotonation step occurred as pH increased from 9.6 to 13.7 (Fig. 5c), which was caused by the proton dissociation of the other one neutral imidazole ring, resulting in the following spectral changes: the absorption intensities for 287 and 476 nm bands increased considerably by 67 and 48%, respectively, and two new bands centered at about 335 and 382 nm appeared without any position shift. The above-mentioned three successive protonation/ deprotonation processes were summarized in Scheme 2. By sigmoidal fitting of the data in plots of the absorbance at 304 nm vs. pH (insets of Fig. 5), the negative logarithms of ground-state acid ionization constants values, pKa1 = 1.44, pKa2 = 7.33 and pKa3 = 12.20 were obtained. On comparison with pKa values previously reported for analogous complex [Ru(bpy)2(Happip)] 2+ (pKa1 = 2.48 ± 0.03, pKa2 = 8.94 ± 0.02) [56] and parent complex [Ru(bpy)2(Hpip)]2+ (pKa1 = 2.17, pKa2 = 8.82) [56] derived under identical experimental conditions, [Ru(Happip)3] 2+ behaves as a stronger acid with its pKa1 and pKa2 values about 1.0 and 1.5 pH units smaller than the Fig. 5. pH effects on the UV-vis spectra of [Ru(Happip)3]2+ (4 μM). (a) pH 0–2.1, (b) pH 2.5–9.4 and (c) pH 9.6–13.7. Arrows show spectral changes upon increasing pH.

corresponding values of [Ru(bpy)2(Happip)]2+ and [Ru(bpy)2(Hpip)]2+. It is interesting to note that the second protonation/deprotonation process for [Ru(Happip)3] 2+ was observed to occur near-physiological pH region (pKa2 = 7.33).

Fig. 4. Effects of increasing amounts of [Ru(Happip)3]2+ on the relative viscosities of the DNA in buffered 50 mM NaCl at 32.0 ± 0.1 °C.

3.2.2. Emission spectra The emission spectral changes of [Ru(Happip)3] 2+ as a function of pH are shown in Fig. 6. It can be seen that the emission spectra of the complex were strongly pH dependent. The insets of Fig. 6 clearly show that the profiles for the changes of emission intensities at 597 nm vs. pH, consist of three sigmoidal curves, indicative of three excited-state protonation/deprotonation processes, which are in accordance with the ground-state processes as revealed by UV-vis spectral spectroscopy. As pH increased from 0.4 to 1.7 (Fig. 6a), the emission maxima had a bathochromic shift from 597 to 613 nm, and the intensities decreased to original 40.5%. Upon increasing

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X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

pH from 2.5 to 9.4 (Fig. 6b), the emission maxima were almost unchanged, while a sharp decrease in the intensities was observed with a large “on–off” intensity ratio of 88. Subsequently, the emission intensities were found to increase significantly by more than 50 folds when pH was increased from 9.6 to 13.7 (Fig. 6c). In view of its switching type and on–off intensity ratios, pH-induced “on–off-on” luminescence switching properties observed for [Ru(Happip)3] 2+ are unusual as compared with “off–on–off” and “off–on–off–off” multi-state luminescence switching properties previously reported for Ru(II) complexes as listed in Table 1. Particularly, the acidic pHdriven luminescence on–off intensity ratio (88) compares favorably with the switching intensity ratios (1.9–20) for all the other Ru(II) complexes. The excited-state acid ionization constants of the complex, pKa*, were roughly evaluated based on the Föster cycle [81], which correlates pKa* with pKa thermodynamically by Eq. (9): pK a ¼ pK a þ ð0:625=T Þðν B –ν HB Þ

ð9Þ

in which νB and νHB are pure 0–0 transitions in cm − 1 for the basic and acidic species, respectively. In practice, νB and νHB are often difficult or even impossible to obtain, so a good approximation is to use

the emission band maxima of both protonated and deprotonated forms of the complex for νB and νHB [82]. Three pKa* values of pKa1* = 0.64, pKa2* = 7.33 and pKa3* = 12.20 were derived. All of these three excited-state pKa values are less or comparable to corresponding ground-state ones, indicating that the excited electrons are localized on the phenanthroline moiety rather than on the imidazole moiety in excited [Ru(H2appip)3] 5+*, [Ru(Happip)3] 2+* and [Ru(appip)2(Happip)]*.

3.3. DNA photocleavage studies The DNA photocleavage abilities of [Ru(bpy)2(Happip)] 2+, [Ru(Happip)3] 2+ and their parent complex [Ru(bpy)2(Hpip)] 2+ were examined by the agarose gel electrophoresis pattern of plasmid pUC 18 DNA. When plasmid DNA was subjected to electrophoresis, relatively fast migration is observed for the supercoiled form (Form I), whereas if scission occurs on one strand (nicking), the supercoils will relax to generate a slower moving open circular form (Form II); and if both strands are cleaved, a linear form (Form III) that migrates between Form I and Form II will be generated. Fig. 7 and S4 summarize the results. As shown in Fig. 7a, the control samples, i.e. the untreated

Scheme 2. Acid–Base Equilibria of [Ru(Happip)3]2+.

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

Fig. 6. pH effects on the emission spectra of [Ru(Happip)3]2+ (4 μM), (a) pH 0–2.1, (b) pH 2.5–9.4 and (c) pH 9.6–13.7. Arrows show spectral changes upon increasing pH.

DNA and treated DNA with the complexes in the dark did not show any cleavage (lanes 0 and 1). However, upon irradiation at 360 nm for 2 h, prominent DNA cleavage was observed with increasing concentrations of [Ru(bpy)2(Happip)] 2+ and [Ru(bpy)2(Hpip)] 2+, as

73

evidenced by the disappearance of Form I and the appearance of Form II. When concentration reached 60 μM for [Ru(bpy)2(Happip)]2+ and 40 μM for [Ru(bpy)2(Hpip)] 2+, DNA was completely converted from Form I to Form II, indicating that [Ru(bpy)2(Hpip)] 2+ exhibited more effective DNA cleavage activity than [Ru(bpy)2(Happip)]2+ under the same experimental conditions. For higher loading of [Ru(Happip)3] 2+ resulting in complete sedimentation of the DNA [83,84], the cleavage experiments by [Ru(Happip)3]2+ could only be carried out over concentrations≤ 2.0 μM. As shown in Fig. S4, DNA cleavage activity (18.8% cleaving efficiency, Lane 4 in Fig. S4) of 2.0 μM of [Ru(Happip)3] 2+ was comparable to that (20.2%) (Lane 5) of [Ru(bpy)2(Happip)] 2+, while weaker than an efficiency of 40.5% (Lane 5) observed for Ru(bpy)2(Hpip)] 2+ under the identical conditions. In attempts to unravel the DNA photocleavage mechanism, a few control experiments were conducted in the presence of singlet oxygen scavengers (Histidine and NaN3) [85], hydroxyl radical scavengers (Mannitol and DMSO) [86,87] and superoxide scavenger (Tiron) [16,87]. Fig. 7b and c show the cleavage of DNA in the presence of the complexes and inhibiting agents, together with the bar diagram of the percentage of cleavage (C). It was observed that the photocleavage by the complexes was not practically inhibited by Mannitol, but significantly inhibited by histidine, NaN3, DMSO and Tiron. This suggests that 1O2 and O2•− that are produced by photochemical reactions of the Ru(II) complexes with O2, are responsible for the DNA photocleavage, similar to previously reported analogous complexes of [Ru(bpy)2(Hmaip)] 2+ {Hmaip=2-(3-aminophenyl)imidazo[4,5-f][1,10]phenanthroline} [29], [Ru(bpy)2(Hpaip)]2+ {Hpaip=2-(4-aminophenyl)imidazo[4,5-f][1,10] phenanthroline} [29], [Ru(dmb)2(Hfpp)]2+ {dmb=4,4′-dimethyl-2,2′bipyridine, Hfpp=2-(3′,3′-difluoro-3,4-methylenedioxyphenyl)imidazo [4,5-f][1,10]phenanthroline} [30] and [Ru(bpy)2(Hfpp)]2+ [30]. Since singlet oxygen plays an important role in the photoactivated cleavage of the plasmid DNA, we further investigated the 1O2 generation abilities of [Ru(bpy)2(Happip)] 2+, [Ru(Happip)3] 2+ and [Ru(bpy)2(Hpip)] 2+. The 1O2 generation quantum yields (ΦΔ) of the complexes were calculated according to Eqs. (10) and (11) [32,39,88], where Iin is the incident monochromatic light intensity, Φab is the light absorbing efficiency of the photosensitizer, Φr is the reaction quantum yield of 1O2 with DPBF, t is the irradiation time, I0 and It are the fluorescence intensity of DPBF before and after irradiation, k is the slope and superscript s stands for standard sample. With [Ru(bpy)3]2+ as standard (ΦΔ = 0.81) [32,39], and by substituting the k values of 0.02058 ± 0.00017, 0.01538 ± 0.00018, 0.04250± 0.00058 and 0.01431 ± 0.00016 for [Ru(bpy)3]2+, [Ru(bpy)2(Happip)]2+, [Ru(Happip)3] 2+ and [Ru(bpy)2(Hpip)] 2+ that derived from the data in Fig. 8, the Φab values of 0.0394, 0.0720, 0.150 and 0.0592 obtained based on the 480 nm absorbance of the complexes in sample solutions (Fig. S5), the ΦΔ values of [Ru(bpy)2(Happip)]2+, [Ru(Happip)3] 2+ and [Ru(bpy)2(Hpip)] 2+ in methanol were determined to be 0.33, 0.44 and 0.37, respectively, which are higher than 0.09 for [Ru(bpy)2 (dppz)]2+ [31,32,35], comparable to 0.36 for [Ru(bpy)2(nitatp)] 2+ {nitatp = 5-nitro-isatino[1,2-b]-1,4,8,9-tetraazatriphenylene} [31], 0.37 for [Ru(phen)2(ppd)] 2+ {ppd = pteridino[6,7-f] [1,10]phenanthroline11,13(10H,12H)-dione} [32], and 0.43 for [Ru(bpy)2(mitatp)]2+ {mitatp = 5-methoxy-isatino[1,2-b]-1,4,8,9-tetraazatriphenylene} [31],

Table 1 Comparison of pKa and pKa* Values and Luminescence on/off Intensity Ratios for Ruthenium(II) Complex-Based Multi-State Luminescence Switches. a

Switch type

pKa

pKa*

On/off ratio (pH region)

Ref

[Ru(bpy)2(H2pipip)]2+ [Ru(bpy)2(Hbopip)]2+ [(bpy)2Ru(H2bpib)Ru(bpy)2]4+ [Ru(Hecip)(Hdcbpy)(NCS)2][Ru(Hipdpa)(Hdcbpy)(NCS)][Ru(bpy)2(H2mpipip)]2+

off–on–off off–on–off off–on–off off–on–off off–on–off off–on–off–off

0.6, 4.7, 10.7 1,70, 5.23, 8.22 4.11, 7.84 0.79, 2.44, 4.29, 6.02, 10.63 1.33,1.98,4.73,10.36 1.23, 6.02, 10.24

0.7, 5.4, 11.4 3.06, 5.01, 8.22 4.34, 7.46 –, –, –, 8.09, 9.47 –, –, 6.78, 9.59 1.29, 6.13, 11.46

2.6 (0.08–1.9); 65.3 (1.9–13.0) 20 (1.0–3.0); 3.0 (3.2–9.4) 2.7 (2.0–6.0); 4.6 (6.0–9.0) 4.4 (2.0–8.0); 2.6 (8.0–13) 4.2 (3.5–8.0); 2.6 (8.5–12) 1.9(1.9–8.8), 8.9 (8.8–12.6)

[43] [10] [45] [12] [11] [44]

Complex

a

See Abbreviations section.

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X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

Fig. 7. (a) Photoactivated cleavage of pUC 18 DNA in the presence of different concentrations of [Ru(bpy)2(Happip)]2+ and [Ru(bpy)2(Hpip)]2+ after irradiation at 360 nm for 2 h. Lane 1: DNA treated with the complexes in the dark. (b) Photoactivated cleavage of pUC 18 DNA by [Ru(bpy)2(Happip)]2+ and [Ru(bpy)2(Hpip)]2+ in the absence and presence of different inhibitors (50 mM Histidine, 40 mM NaN3, 100 mM Mannitol, 200 mM DMSO, 10 mM Tiron) after irradiation at 360 nm for 2 h. (c) Bar diagram representation of the effect of the inhibitors on the DNA photoactivated cleavage.

and lower than 0.50 for [Ru(phen)(ppd)2] 2+ [32] and 0.51 for [Ru(btz)2(dppz)]2+ (btz = 4,4′-bithiazole) [35]. The high 1O2 generation efficiencies observed for [Ru(bpy)2(Happip)] 2+, [Ru(Happip)3]2+ and [Ru(bpy)2(Hpip)] 2+ contributed to their efficient photoactivated DNA cleavage.

−Δ½DPBF=t ¼ ðI0 –It Þ=t ¼ Iin Φab ΦΔ Φr

ð10Þ

s

k=ks ¼ Φab ΦΔ =Φab ΦΔ

s

4. Conclusion In summary, a new ruthenium(II) complex [Ru(Happip)3](ClO4)2 has been synthesized and characterized. The optical spectroscopy and DNA viscosities indicated that the complex bound to the DNA in a partially intercalative mode with binding constant value on 10 6 M − 1 order of magnitude. The complex underwent three-step protonation/deprotonation processes with one of the processes occurring near-physiological pH region (pKa2 = 7.33), and acted as an excellent “on–off–on” pH-luminescence switch, with two large luminescence “on–off” intensity ratios of 88 and 50. When irradiated at 360 nm, [Ru(Happip)3](ClO4)2 and its two analogous complexes of [Ru(bpy)2(Happip)] 2+ and [Ru(bpy)2(Hpip)] 2+ were found to be efficient plasmid pUC 18 photocleavers with 1O2 and O2•− being the active species. The high singlet oxygen generation quantum yields 0.33–0.44 in methanol can be rationalized as one of key factors to contribute to the efficient DNA cleavage observed. 5. Abbreviations

Fig. 8. The emission spectral changes of DPBF (20 μM) in the presence of [Ru(bpy)2(Happip)] 2+ (5 μM) as irradiated at 480 nm. Inset: the DPBF consumption percentage as a function of irradiation time in the air-equilibrated methanol solution of [Ru(bpy)2 (Happip)] 2+.

ð11Þ

bipp bopp bpy btpp btppz btz

2-benzimidazoyl-pyrazino[2,3-f][1,10]phenanthroline 2-benzoxazolyl-pyrazino[2,3-f][1,10]phenanthroline 2,2′-bipyridine 2-benzthiazolyl-pyrazino[2,3-f][1,10]phenanthroline} benzo[h]tripyrido[3,2-a:2′,3′-c:2′′,3′′-j]phenazine 4,4′-bithiazole

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

cdpq deprotonated 2-carboxyldipyrido[3,2-f:2′,3′-h]quinoxaline dmb 4,4′-dimethyl-2,2′-bipyridine dmp 2,9-dimethyl-1,10-phenanthroline DPBF 1,3-diphenylisobenzofuran dppz dipyrido[3,2-a:2′,3′-c]phenazine H2bpib 1,4-bis([1,10]phenanthroline[5,6-d]-imidazol-2-yl)benzene H2mpipip 2-(3-(1H-phenanthro[9,10-d]imidazol-2-yl)phenyl)-1Himidazo[4,5-f][1,10]-phenanthroline H2pipip 2-(4-(1H-phenanthro[9,10-d]imidazol-2-yl)phenyl)-1Himidazo[4,5-f][1,10]-phenanthroline Happip 2-(4-(β-D-allopyranoside)phenyl)imidazo[4,5-f][1,10] phenanthroline Hdcbpy mono-deprotonated 2,2′-bipyridyl-4,4′-dicarboxylic acid Hecip 2-(9-ethyl-9H-carbazol-3-yl)-1H-imidazo[4,5-f][1,10] phenanthroline Hfpp 2-(3′,3′-difluoro-3,4-methylenedioxyphenyl)imidazo[4,5f][1,10]phenanthroline Hipdpa 4-(1H-imidazo[4,5-f][1,10]phenanthrolin-2-yl)-N,Ndiphenylaniline Hmaip 2-(3-aminophenyl)imidazo[4,5-f][1,10]phenanthroline Hmip 2-(2,3-methylenedioxyphenyl)imidazo[4,5-f]1,10phenanthroline Hpaip 2-(4-aminophenyl)imidazo[4,5-f][1,10]phenanthroline Hpip 2-(4-phenyl)imidazo[4,5-f][1,10]phenanthroline mbeb 4-[(1E)-2-(4′-methyl[2,2′-bipyridin]-4-yl)ethenyl]-1,2benzenediol mitatp 5-methoxy-isatino[1,2-b]-1,4,8,9-tetraazatriphenylene nitatp 5-nitro-isatino[1,2-b]-1,4,8,9-tetraazatriphenylene phen 1,10-phenanthroline ppd pteridino[6,7-f] [1,10]phenanthroline-11,13(10H,12H)-dione UV-vis UV-visible Acknowledgments The authors thank the National Natural Science Foundation (nos. 21171022, 20971016, 90922004), Beijing Natural Science Foundation (2072011), the Fundamental Research Funds for the Central Universities, and Measurements Fund of Beijing Normal University for financial supports. Appendix A. Supplementary data Supplementary data to this article can be found online at doi:10. 1016/j.jinorgbio.2012.03.010. References [1] T.D. Tullius, Nature 332 (1988) 663–664. [2] N.Y. Sardesai, K. Zimmermann, J.K. Barton, J. Am. Chem. Soc. 116 (1994) 7502–7508. [3] C. Moucheron, A. Kirsch-DeMesmaeker, J.M. Kelly, Photochem. Photobiol. B 40 (1997) 91–106. [4] B. Armitage, Chem. Rev. 98 (1998) 1171–1200. [5] P.K.L. Fu, P.M. Bradley, C. Turro, Inorg. Chem. 40 (2001) 2476–2477. [6] B. de Souza, F.R. Xavier, R.A. Peralta, A.J. Bortoluzzi, G. Conte, H. Gallardo, F.L. Fischer, G. Bussi, H. Terenzi, A. Neves, Chem. Commun. 46 (2010) 3375–3377. [7] A.D. Kulkarni, S.A. Patil, V.H. Naik, P.S. Badami, Med. Chem. Res. 20 (2011) 346–354. [8] Y.M. Chen, Y.J. Liu, Q. Li, K.Z. Wang, J. Inorg. Biochem. 103 (2009) 1395–1404. [9] G.Y. Bai, K.Z. Wang, Z.M. Duan, L.H. Gao, J. Inorg. Biochem. 98 (2004) 1017–1022. [10] M.J. Han, L.H. Gao, Y.Y. Lu, K.Z. Wang, J. Phys. Chem. B 110 (2006) 2364–2371. [11] S.H. Fan, A.G. Zhang, C.C. Ju, L.H. Gao, K.Z. Wang, Inorg. Chem. 49 (2010) 3752–3763. [12] S.H. Fan, K.Z. Wang, W.C. Yang, Eur. J. Inorg. Chem. (2009) 508–518. [13] C.C. Ju, A.G. Zhang, C.L. Yuan, X.L. Zhao, K.Z. Wang, J. Inorg. Biochem. 105 (2011) 435–443. [14] X.L. Zhao, M.J. Han, A.G. Zhang, K.Z. Wang, J. Inorg. Biochem. 107 (2012) 104–110. [15] D.L. Arockiasamy, S. Radhikaa, R. Parthasarathia, B. Unni Nair, Chem. Commun. 44 (2009) 2044–2051. [16] M. Mariappan, B.G. Maiya, Eur. J. Inorg. Chem. (2005) 2164–2173. [17] J. Malina, M.J. Hannon, V. Brabec, Chem. Eur. J. 14 (2008) 10408–10414. [18] A.G. Zhang, Y.Z. Zhang, Z.M. Duan, K.Z. Wang, Inorg. Chem. 50 (2011) 6425–6436.

75

[19] A. Ghosh, P. Das, M.R. Gill, P. Kar, M.G. Walker, J.A. Thomas, A. Das, Chem. Eur. J. 17 (2011) 2089–2098. [20] L.F. Tan, H. Chao, Inorg. Chim. Acta 360 (2007) 2016–2022. [21] L.F. Tan, H. Chao, K.C. Zhen, J.J. Fei, F. Wang, Polyhedron 6 (2007) 5458–5468. [22] J.G. Liu, B.H. Ye, Q.L. Zhang, X.H. Zou, Q.X. Zhen, X. Tian, L.N. Ji, J. Biol. Inorg. Chem. 5 (2000) 119–128. [23] A.E. Friedman, C.V. Kumar, N.J. Turro, J.K. Barton, Nucleic Acids Res. 19 (1991) 2595–2602. [24] R.M. Hartshorn, J.K. Barton, J. Am. Chem. Soc. 114 (1992) 5919–5925. [25] I. Haq, J. Ladbury, J. Mol. Recognit. 13 (2000) 188–197. [26] J.M. Kelly, M.J. Murphy, D.J. McConnell, C. OhUigin, Nucleic Acids Res. 13 (1985) 167–184. [27] M.B. Fleisher, K.C. Waterman, N.J. Turro, J.K. Barton, Inorg. Chem. 25 (1986) 3549–3551. [28] G.J. Ryan, S. Quinn, T. Gunnlaugsson, Inorg. Chem. 47 (2008) 401–403. [29] Y.J. Liu, C.H. Zeng, H.L. Huang, L.X. He, F.H. Wu, Eur. J. Med. Chem. 45 (2010) 564–571. [30] Y.J. Liu, C.H. Zeng, F.H. Wu, J.H. Yao, L.X. He, H.L. Huang, J. Mol. Struct. 932 (2009) 105–111. [31] H.J. Yu, S.M. Huang, L.Y. Li, H.N. Jia, H. Chao, Z.W. Mao, J.Z. Liu, L.N. Ji, J. Inorg. Biochem. 103 (2009) 881–890. [32] F. Gao, H. Chao, Y.F. Wei, Y.X. Yuan, B. Peng, X. Chen, K.C. Zheng, L.N. Ji, Helv. Chim. Acta 91 (2008) 395–410. [33] L.C. Xu, S. Shi, J. Li, S.Y. Liao, K.C. Zheng, L.N. Ji, Dalton Trans. (2008) 291–301. [34] L.F. Tan, X.L. Liang, X.H. Liu, J. Inorg. Biochem. 103 (2009) 441–447. [35] H.J. Yu, H. Chao, L. Jiang, L.Y. Li, S.M. Huang, L.N. Ji, Inorg. Chem. Commun. 11 (2008) 553–556. [36] M.R. Detty, S.L. Gibson, S.J. Wagner, J. Med. Chem. 47 (2004) 3897–3915. [37] Q.X. Zhou, W.H. Lei, C. Li, Y.J. Hou, X.S. Wang, B.W. Zhang, New J. Chem. 34 (2010) 137–140. [38] A. Hergueta-Bravo, M.E. Jiménez-Hernández, F. Montero, E. Oliveros, G. Orellana, J. Phys. Chem. B 106 (2002) 4010–4017. [39] H.Y. Ding, X.S. Wang, L.Q. Song, J.R. Chen, J.H. Yu, C. Li, B.W. Zhang, J. Photochem. Photobiol. A 177 (2006) 286–294. [40] P.K.L. Fu, P.M. Bradley, D. van Loyen, H. Dürr, S.H. Bossmann, C. Turro, Inorg. Chem. 41 (2002) 3808–3810. [41] M. Haga, T. Takasugi, A. Tomie, M. Ishizuya, T. Yamada, M.D. Hossain, M. Inoue, Dalton Trans. (2003) 2069–2079. [42] A. Quaranta, F. Lachaud, C. Herrero, R. Guillot, M.F. Charlot, W. Leibl, A. Aukauloo, Chem. Eur. J. 13 (2007) 8201–8211. [43] F. Gao, H. Chao, F. Zhou, B. Peng, L.N. Ji, Inorg. Chem. Commun. 10 (2007) 170–173. [44] F. Gao, X. Chen, F. Zhou, L.P. Weng, L.T. Guo, M. Chen, H. Chao, L.N. Ji, Inorg. Chim. Acta 362 (2009) 4960–4966. [45] H. Chao, B.H. Ye, Q.L. Zhang, L.N. Ji, Inorg. Chem. Commun. 2 (1999) 338–340. [46] E. Evangelio, J. Hernando, I. Imaz, G.G. Bardají, R. Alibés, F. Busqué, D. Ruiz-Molina, Chem. Eur. J. 14 (2008) 9754–9763. [47] L. Stryer (Ed.), Biochemistry, Freeman, New York, 1988. [48] D. Margulies, G. Melman, A. Shanzer, Nat. Mater. 4 (2005) 768–771. [49] A.P. de Silva, S. Uchiyama, Nat. Nanotechnol. 2 (2007) 399–410. [50] F.R. Liu, K.Z. Wang, G.Y. Bai, Y.G. Zhang, L.H. Gao, Inorg. Chem. 43 (2004) 1799–1806. [51] M.J. Han, Z.M. Duan, Q. Hao, K.Z. Wang, J. Phys. Chem. C 111 (2007) 16577–16585. [52] D. Ye, L. Fu, B. Liang, S.F. Yin, Y. Li, Chin. J. Org. Chem. 29 (2009) 1768–1773. [53] H.F. Chen, H.W. Chen, X. Bai, Y. Li, S.F. Yin, Chin. J. Org. Chem. 31 (2011) 231–234. [54] A.M. Pyle, M.Y. Chiang, J.K. Barton, Inorg. Chem. 29 (1990) 4487–4495. [55] M.N. Ackermann, L.V. Interrante, Inorg. Chem. 23 (1984) 3904–3911. [56] Y.Z. Ma, H.J. Yin, K.Z. Wang, J. Phys. Chem. B 113 (2009) 11039–11047. [57] J.Z. Wu, B.H. Ye, L. Wang, L.N. Ji, J.Y. Zhou, R.H. Li, Z.Y. Zhou, J. Chem. Soc., Dalton Trans. (1997) 1395–1401. [58] J.Z. Wu, L.N. Ji, Transition Met. Chem. 24 (1999) 299–303. [59] A. Wolfe, G.H. Shimer Jr., T. Meehan, Biochemistry 26 (1987) 6392–6396. [60] K.C. Zheng, H. Deng, X.W. Liu, H. Li, H. Chao, L.N. Ji, J. Mol. Struct. (THEOCHEM) 682 (2004) 225–233. [61] M.T. Carter, M. Rodriguez, A.J. Bard, J. Am. Chem. Soc. 111 (1989) 8901–8911. [62] Q.X. Zhou, B.H. Ye, Q.L. Zhang, J.G. Liu, H. Li, L.N. Ji, L. Wang, J. Inorg. Biochem. 76 (1999) 47–53. [63] Y. Xiong, X.F. He, X.H. Zou, J.Z. Wu, X.M. Chen, L.N. Ji, R.H. Li, J.Y. Zhou, K.B. Yu, J. Chem. Soc., Dalton Trans. (1999) 19–24. [64] W.D. Wilson, L. Ratmeyer, M. Zhao, L. Strekowski, D. Boykin, Biochemistry 32 (1993) 4098–4101. [65] I. Ortmans, B. Elias, J.M. Kelly, C. Moucheron, A. Kirsch-DeMesmaeker, Dalton Trans. (2004) 668–676. [66] B.F. Cain, B.C. Baguley, W.A. Denny, J. Med. Chem. 21 (1978) 658–668. [67] D.L. Boger, B.E. Fink, S.R. Brunette, W.C. Tse, M.P. Hedrick, J. Am. Chem. Soc. 123 (2001) 5878–5891. [68] R. Sarkar, S.K. Pal, Biopolymers 83 (2006) 675–686. [69] K.J. Breslauer, D.P. Remeta, W.Y. Chou, R. Ferrante, J. Curry, D. Zaunczkowski, J.G. Snyder, L.A. Marky, Proc. Natl. Acad. Sci. U. S. A. 84 (1987) 8922–8926. [70] B.C. Baguley, E.M. Falkenhaug, Nucl. Acids Res. 5 (1978) 161–171. [71] D. Banerjee, S.K. Pal, J. Phys. Chem. B 111 (2007) 5047–5052. [72] W.K. Mudasir, W.E. Tri, N. Yoshioka, H. Inoue, Biophys. Chem. 121 (2006) 44–50. [73] D.J. Patel, Acc. Chem. Res. 12 (1979) 118–125. [74] H.M. Berman, P.R. Young, Annu. Rev. Biophys. Bioeng. 10 (1981) 87–114. [75] J.M. Kelly, A.B. Tossi, D.J. McConell, C. OhUigin, Nucleic Acids Res. 13 (1985) 6017–6034.

76

X.-L. Zhao et al. / Journal of Inorganic Biochemistry 113 (2012) 66–76

[76] G.A. Neyhart, N. Grover, S.R. Smith, W.A. Kalsbeck, T.A. Fairley, M. Cory, H.H. Thorp, J. Am. Chem. Soc. 115 (1993) 4423–4428. [77] J.D. McGhee, Biopolymers 15 (1976) 1345–1375. [78] S. Satyanarayana, J.C. Dabrowiak, J.B. Chaires, Biochemistry 32 (1993) 2573–2584. [79] F. Leng, J.B. Chaires, M.J. Waring, Nucleic Acids Res. 31 (2003) 6191–6197. [80] S. Satyanarayana, J.C. Dabrowiak, J.B. Chaires, Biochemistry 31 (1992) 9319–9324. [81] E.V. Donckt, in: G. Poter (Ed.), Progress in Reaction Kinetics, 5th ed., Pergamon Press, Oxford, England, 1970, pp. 273–299. [82] G.Y. Zheng, Y. Wang, D.P. Rillema, Inorg. Chem. 35 (1996) 7118–7123.

[83] S.P. Foxon, T. Phillips, M.R. Gill, M. Towrie, A.W. Parker, M. Webb, J.A. Thomas, Angew. Chem. Int. Ed. 46 (2007) 3686–3688. [84] B. Sun, J.X. Guan, L. Xu, B.L. Yu, L. Liang, J.F. Kou, L. Wang, X.D. Ding, H. Chao, L.N. Ji, Inorg. Chem. 48 (2009) 4637–4639. [85] R. Nilsson, P.B. Merkel, D.R. Kearns, Photochem. Photobiol. 16 (1972) 117–124. [86] C.C. Cheng, S.E. Rokita, C.J. Burrows, Angew. Chem. Int. Ed. 32 (1993) 277–278. [87] S.A. Lesko, R.J. Lorentzen, P.O.P. Ts'o, Biochemistry 19 (1980) 3023–3028. [88] R.H. Young, K. Wehrly, R.L. Martin, J. Am. Chem. Soc. 93 (1971) 5774–5779.