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Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I and IIα
Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I and IIα Yanmei Sun, Jia Li, Hong Zhao, Lifeng Tan PII: DOI: Reference:
S0162-0134(16)30110-6 doi: 10.1016/j.jinorgbio.2016.04.028 JIB 9986
To appear in:
Journal of Inorganic Biochemistry
Received date: Revised date: Accepted date:
28 December 2015 12 April 2016 18 April 2016
Please cite this article as: Yanmei Sun, Jia Li, Hong Zhao, Lifeng Tan, Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I and IIα, Journal of Inorganic Biochemistry (2016), doi: 10.1016/j.jinorgbio.2016.04.028
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ACCEPTED MANUSCRIPT Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I
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and IIα
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Yanmei Sun a,1, Jia Li a,1, Hong Zhao a, Lifeng Tan*,b a College of Chemistry, Xiangtan University, Xiangtan, P. R. China
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b Key Lab of Environment-friendly Chemistry and Application in Ministry of Education, Xiangtan
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University, Xiangtan, P. R. China
* Corresponding author at: College of Chemistry, Xiangtan University, Xiangtan 411105, PR China. Tel.: +86 731 58293997; Fax: +86 731 58293477. E-mail address: [email protected] (L.-F. Tan). 1
These two authors are both first authors.
ABSTRACT Two ruthenium(II) polypyridyl complexes containing 1,8-naphthalimide group as DNA binders, 1
ACCEPTED MANUSCRIPT photonucleases, and inhibitors of topoisomerases I and IIα are evaluated. The binding properties of [Ru(phen)2(pnip)]2+ {1; phen =1,10-phenanthroline; pnip = 12-[N-(p-phenyl)-1,8-napthalimide]-
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imidazo[4',5'-f][1,10]phenanthroline} and [Ru(bpy)2(pnip)]2+ (2; bpy = 2,2′-bipyridine) with calf
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thymus DNA increases with increasing the bulkiness and hydrophobic character of ancillary ligands,
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although the two complexes possess high affinities for DNA via intercalation. Moreover, photoirradiation (λ = 365 nm) of the two complexes are found to induce strand cleavage of closed
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circular pBR322 plasmid DNA via singlet oxygen mechanism, while complex 1 displays more
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effective photocleavage activity than complex 2 under the same conditions. Topoisomerase inhibition and DNA strand passage assay reflect that complexes 1 and 2 are efficient dual poisons of
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topoisomerases I and IIα.
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Keywords: Ruthenium(II) complexes; DNA binding; Photocleavage; Topoisomerase inhibition
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ACCEPTED MANUSCRIPT 1. Introduction DNA topoisomerases are essential for the processes of chromosomal segregation and relaxation
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of DNA during replication and transcription, and may also be required for recombination [1]. Two
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types of DNA topoisomerases have been isolated from prokaryotes and eukaryote. In general,
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topoisomerase I (Topo I) catalyzes the relaxation of superhelical DNA by generating a transient single strand nick in the duplex through cycles of cleavage and religation [2], while topoisomerase
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II (Topo II) mediates the ATP-dependent induction of coordinated nicks in both strands of the DNA
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duplex through which another DNA segment is passed [3]. Under normal conditions, the step of DNA relegation is much faster than that of DNA cleavage, which may be tolerated by the cell.
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However, conditions that significantly change either the physiological concentration or the lifetime
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of the breaks are responsible for DNA alterations, playing a crucial role in inhibiting cell cycle
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progression [4]. Therefore, Topoisomerases are the cellular targets of clinically important anticancer and antibacterial drugs, and inhibition of topoisomerases has been considered as an
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effective strategy for the design of many anticancer agents [5]. In recent years, numerous natural and synthetic compounds have been chosen to detect the inhibition of topoisomerase activity [6–10]. To date, some topoisomerase inhibitors, such as camptothecin (CPT) and its derivatives that target Topo I [11] and etoposide (VP-16) [12], doxorubicin [13] and mitoxantrone [14] which target Topo II, have been clinically used as potent anticancer drugs. These agents have achieved great success in clinical cancer chemotherapy, while single inhibitors of topoisomerases are limited by several important negative consequences [15]. Furhtermore, the emergence of resistance phenomena to Topo I inhibitors is often accompanied by a concomitant rise at the level of Topo II expression and viceversa, resulting in the failure of clinical therapies [15]. In this regard, a single compound able to inhibit both Topo I and II may 3
ACCEPTED MANUSCRIPT present the advantage of improving antitopoisomerase activity, with reduced toxic side effects, with respect to the combination of two inhibitors [16]. Several classes of dual topos I and II inhibitors
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have been identified and described over the last decades [17–21]. Some of these dual inhibitors,
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such as Tafluposide and Batracylin, have been evaluated in clinical trials. Most of such studies at
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present, however, have been mainly revolved around organic compounds and, to a far lesser extent, around metal complexes [22]. In comparison with organic molecules, the interest in the field of
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Ru(II) polypyridyl complexes−nucleic acid interactions has burgeoned due to Ru(II) polypyridyl
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complexes with rich photochemical properties and varied coordination forms [23]. However, surprisingly and in contrast to studies on Ru(II) polypyridyl complexes−nucleic acid interactions,
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investigations of the inhibition of topoisomerases activity by Ru(II) polypyridyl complexes and the
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relationship between the structures of Ru(II) complexes and the enzymatic inhibition activities are
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very scarce [24]. Therefore, studies on the interaction between polypyridyl-based Ru(II) complexes and topoisomerases are very important for developing novel antitumor drugs and elucidating the
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underlying molecular mechanism.
Recently, we reported the synthesis and RNA-binding properties of Ru(II) polypyridyl complexes with 1,8-naphthalimide group (Fig. 1), [Ru(phen)2(pnip)]2+ (1; phen = 1,10-phenanthroline, pnip = 2-[N-(p-phenyl)-1,8-napthalimide]imidazo[4',5'-f][1,10]phenanthroline) and [Ru(bpy)2(pnip)]2+ (2; bpy = 2,2′-bipyridine), finding that the two complexes displayed high binding affinities for the RNA triplex as well as increasing the stability of the Hoogsteen base-paired strand of the triplex obviously [25]. In addition, we note that the development of functional 1,8-naphthalimide and its derivatives as DNA binders, photonucleases and enzyme inhibitors is a fast growing area [26]. However, biological investigations of ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group have attracted little attention relatively [26c]. 4
ACCEPTED MANUSCRIPT Considering all the above, this paper deals with the DNA binding, photo-induced DNA cleavage, and topoisomerase inhibitory activity of complexes 1 and 2. We hope that this work will aid in
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advancing our knowledge of the interaction between polypyridyl-based Ru(II) complexes and DNA,
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as well as laying the foundation for the rational design of dual topoisomerases I and II inhibitors.
2. Experimental sections
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2.1. Materials
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Complexes 1 and 2 were synthesized according to the literature methods [25]. Calf thymus DNA (CT-DNA) approximately 200 base pairs in average length was purchased from the Sino-American
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Biotechnology Company. DNA topoisomerase I from calf thymus together with human
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topoisomerase IIa were purchased from TopoGen Inc. Tris–HCl buffer A (5 mM Tris–HCl, 50 mM
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NaCl, pH 7.0) (Tris = tris(hydroxymethyl)aminomethane) solution was prepared using doubly distilled water. A solution of CT-DNA in the buffer gave a ratio of UV (UV = ultraviolet)
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absorbance at 260 and 280 nm of ca. 1.8–1.9:1, indicating that the DNA was sufficiently free of protein [27]. The DNA concentration per nucleotide was determined by absorption spectroscopy using the molar absorption coefficient (6600 M-1 cm-1) at 260 nm [28].
2.2. DNA-binding experiments UV–vis spectra were collected using a PerkinElmer Lambda 25 spectrophotometer at 20 oC. A typical titration of each Ru(II) complex in buffer A was performed by using a fixed concentration of either complex 1 or 2, to which the DNA stock solution was gradually added up to saturation. After each addition, the solution was mixed and allowed to reequilibrate for 5 min before the absorption spectra were recorded. The intrinsic binding constant Kb to DNA was determined the changes of 5
ACCEPTED MANUSCRIPT MLCT (MLCT = metal to ligand charge transfer) band by using the following equation [29]:
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(1a)
(1b)
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b = 1+ K b C t + K b [DNA]/(2s)
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b b2 2 Kb2Ct DNA / s a f b f 2 KCt
where [DNA] is the concentration of DNA in base pairs and the apparent absorption
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coefficients εa, εf, and εb correspond to A obsd /[Ru], the extinction coefficient for the free Ru(II) complex, and the extinction coefficient for a Ru(II) complex in the fully bound form,
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respectively. Kb is the equilibrium binding constant in M-1, Ct is the total metal complex
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2.3. Thermal denaturation studies
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concentration, and s is the binding site size.
PerkinElmer Lambda 25
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The thermal denaturation studies were performed on a
spectrophotometer equipped with a water-circulating bath. The temperature was ramped from 50 to
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90 °C with an increasing rate of 1.0 °C min−1. The melting curves were obtained by measuring the absorption of DNA solution at 260 nm in the absence and presence of the ruthenium complex (dissolved in buffer A) as a function of temperature. The data were presented as (A − A0)/(Af − A0) versus temperature, where A, A0, and Af are the observed, the initial, and the final absorbance at 260 nm, respectively.
2.4. Viscosity studies Viscosity measurements were carried out using an ubbelohde viscometer maintained at 25.0 ± 0.1 °C in buffer A. The flow time was measured using a digital stopwatch, and each sample was measured three times. Relative viscosities for the triplex RNA in either the absence or presence of 6
ACCEPTED MANUSCRIPT metal complexes were calculated according to literature procedures reported earlierand an average
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flow time was calculated. Data are presented as (η/η0)1/3 versus the binding ratio [30].
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2.5. DNA photocleavage experiments
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For the gel electrophoresis experiment, supercoiled pBR322DNA (0.05 μg) was treated with different concentration of each Ru(II) complex in buffer B (50 mM Tris-HCl, 18 mM NaCl, pH 7.2),
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and then the solution was irradiated with a UV lamp (365 nm, 10 W) at room temperature. The samples were analyzed by electrophoresis for 1 h at 80 V in buffer C (89 mM Tris, 89 mM boron
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hydroxide, 2 mM EDTA) (EDTA = ethylene diamine tetraacetie acid) containing 0.75% agarose gel. The gel was stained with EB (EB = ethidium bromide;1 mg/mL), photographed under UV light and
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then analysed on the FluorChem FC2. The percentage of photocleavage (C) was determined according to the following equation:
DII 2 DIII 100% DI DII 2 DIII
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C
(2)
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Where DI, DII and DIII are the relative density of supercoiled pBR322 DNA existing in the form of supercoil form (Form I), nicking form (Form II) and linear form (Form III) in the absence and presence of either complex 1 or 2, respectively.
2.6. Topoisomerase inhibition assay The relaxation assay was performed in buffer D (35 mM Tris-HCl, 72 mM KCl, 5 mM MgCl2, 5 mM dithiothreitol, 2 mM spermidine, 0.1 mg/ml bovine serum albumin, pH 8.0) containing 0.05 μg pBR322 DNA and 1 unit of Topo I in the absence and presence of either complex 1 or 2. Control experiments contained either pBR322 DNA alone or pBR322 DNA treated with Topo I only. The reaction mixture (total volume = 10 μL) was incubated at 37 oC for 30 min and quenched by 7
ACCEPTED MANUSCRIPT addition of 4 μL of loading buffer containing 0.25% bromophenol blue, 4.5% SDS and 45% glycerol. The DNA was fractionated by agarose gel electrophoresis by 1% agarose in TBE at 80 V
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for 1 h. The gel was then stained with EB (1 µg/mL), visualized using a UV illuminator and
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eventually photographed under UV light. In addition, IC50 values were the concentrations of the
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inhibitor that prevented 50% of the supercoiled DNA from being converted into relaxed DNA, which were calculated by the midpoint concentration for either complex 1 or 2 induced DNA
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unwinding.
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Topo IIa functional activity was assayed in buffer E (10 mM Tris-HCl, 50 mM NaCl, 50 mM KCl, 5.0 mM MgCl2, 0.1 mM Na2H2EDTA, 15 μg/mL bovine serum albumin, 1.0 mM ATP, pH 7.9)
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containing 0.05 μg pBR322DNA and 4 units of Topo IIa in the absence and presence of either
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complex 1 or 2. The reaction mixture was incubated at 30 oC for 15 min. Reactions were stopped,
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processed, and then subjected to gel electrophoresis.
2.7. DNA strand passage assay
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The DNA strand passage activity of Topo I was determined by investigating the ability of Topo I to relax negatively supercoiled plasmid DNA [29] or to supercoil plasmid DNA [31]. Reactions contained 0.05 μg relaxed or supercoiled pBR322 plasmid DNA in buffer D (total volume = 10 μL). Assays were carried out in the absence of either complex 1 or 2 or in the presence of 25 μM Ru(II) complexes or EB. Following a 5-min incubation, 1 unit of Topo I was added, and reactions were incubated up to 15 min at 37 oC. Reactions were stopped, processed, and subjected to gel electrophoresis as described above. Regarding the DNA strand passage activity of Topo IIa, reactions contained relaxed or supercoiled 0.05 μg pBR322 plasmid DNA and 4 units of Topo IIa in buffer E (total volume = 10 μL). Following a 5-min incubation, 4 units of Topo IIa was added and 8
ACCEPTED MANUSCRIPT then reactions were incubated up to 15 min at 30 oC. The rest of the operations were the same as
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described above.
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3. Results and discussion
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3.1. DNA binding studies
The interaction of complexes 1 and 2 with CT-DNA was primarily examined by absorption
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spectra titration experiments. Binding of a complex to DNA through a π-π stacking interaction
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between an aromatic chromophore of a complex and the base pairs of DNA typically results in hypochromism and a red shift [32]. Fig. 2 provides the changes in UV-Vis absorption spectra when
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complexes 1 and 2 bound to CT-DNA, and the representing data are summarized in Table 1. Fig. 2
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indicated that the addition of CT-DNA to the solution of either complex 1 or 2 in a Tris buffer led to
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obvious hypochromism in the UV/Vis absorption spectrum with moderate bathochromic spectral shift. For complex 1, the hypochromicity at 459 nm reached about 23% with a red shift of 3 nm at a
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[DNA]/[Comp. 1] ratio of 5.1, whereas for complex 2, the hypochromicity at 457 nm showed about 17% at a [DNA]/[Comp. 2] ratio of 6.0 with a red shift of 3 nm. The spectral characteristics suggested that complexes 1 and 2 could interact with CT-DNA. To further elucidate the binding strength of both complexes, the DNA binding constants (Kb) were estimated from the changes of the absorbance at 459 and 457 nm for complexes 1 and 2, respectively. The Kb values of complexes 1 and 2 are (7.95 ± 0.28) ×106 M-1 (s = 1.24 0.13) and (4.50 0.15) × 106 M-1 (s = 1.01 0.12), respectively. The Kb values of both complexes are comparable to that determined for [Ru(phen)2dppz]2+ [33, [Ru(dtzp)(ptn)]2+ [34] and the known DNA intercalator ethidium bromide [35], while stronger than that of [Ru(bpy)3]2+ (4.7 103 M-1)
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ACCEPTED MANUSCRIPT and [Ru(phen)3]2+ (5.4 103 M-1) [36]. These data indicate that the size and the shape of the intercalated ligand of Ru(II) complexes have a significant effect on the binding strength, and the
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most suitable intercalating ligand leads to the highest affinity of complexes with DNA. In addition,
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the Kb value of complex 1 was slightly greater than that of complex 2, which suggested that the
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binding of complex 1 with DNA is tighter than that of complex 2. In general, the binding reaction of small molecule with nucleic acids is driven primarily by hydrophobic interactions [37]. For
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complexes 1 and 2, as the ancillary ligand progresses from bpy to phen studied, the bulkiness and
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hydrophobic character of the ligand is increased. Consequently, the hydrophobic transfer of the large aromatic complexes 1 from solution into the DNA binding site caused by the ancillary ligand
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phen is easier to achieve, resulting in the binding sites effectively overlapping with each other.
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Therefore, the effects of ancillary ligands (phen and bpy) on the binding characteristics of
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complexes 1 and 2 with DNA should not be ignored, which is likely the main factor affecting the interactions of the two metal complexes with CT-DNA.
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3.2. Determination of the binding mode by viscosity studies To earnestly clarify the binding modes of the complexes and CT-DNA, viscosity measurements were performed. Hydrodynamic measurements that are sensitive to changes in length (i.e. viscosity and sedimentation) are regarded as the least ambiguous and the most critical tests of a binding model in solution in the absence of crystallographic or NMR structural data [38]. In general, the viscosity of double-stranded DNA increases when a complex binds to DNA through a classical intercalating mode [38]. In contrast, the viscosity of DNA will decline when the binding mode is a partial or non-classical intercalation, because this mode could bend (or kink) the DNA helix and concomitantly reduce its effective length [39]. However, it remains unchanged when a complex 10
ACCEPTED MANUSCRIPT binds DNA in an electrostatic mode [38]. The effects of complexes 1 and 2 together with EB and [Ru(bpy)3]2+ on the viscosity of rod-like DNA are presented in Fig. 3. As expected, EB, which is a
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well-known DNA intercalator, strongly increases the relative viscosity by lengthening the DNA
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double helix through intercalation. While for complex [Ru(bpy)3]2+, which has been known to bind
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DNA through electrostatic interactions, exhibited essentially no effect on the viscosity of DNA [40]. It is evident that the binding of either complex 1 or 2 can increase in the relative viscosity of the
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DNA solution upon increasing the amounts of the complex. The increased degree of viscosity,
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which may depend on its affinity to DNA, follows the order of EB > 1 > 2 > [Ru(bpy)3]2+. The results further suggest that the binding modes of complexes 1 and 2 are intercalation, and complex 1
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intercalates into CT-DNA base pairs more deeply than complex 2 does, complementing the above
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results.
3.3. Thermal denaturation studies
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DNA melting experiments are useful to determine the extent of intercalation, because the intercalation of the complex into DNA base pairs causes stabilisation of the base stacking [41] and, therefore, raises the melting temperature of the double-stranded DNA [42]. It is well accepted that when the temperature of the solution increases, the double-stranded DNA gradually dissociates into single strands, which generates a hyperchromic effect on the absorption spectra of the DNA bases (λmax 260 nm). To identify this transition process, the melting temperature (Tm), which is defined as the temperature where half of the total base pairs are unbonded, is usually introduced. Generally, the intercalation of a complex into DNA generally results in a considerable increase of Tm [40]. The melting curve of CT-DNA in the absence and presence of each complex is presented in Fig. 4. The Tm of Ru(II) complex-free CT-DNA was determined to be 64.5 oC. In the presence of either 11
ACCEPTED MANUSCRIPT complex 1 or 2, the Tm increases successively and reaches 78.5 and 74.9 oC, respectively, at a [Ru]/[DNA] ratio of 1 : 10. The ΔTm values (14.0 and 10.4 oC) of 1/2–DNA adducts are larger than
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those of some Ru(II) intercalators [43], which reveals that the modes of both complexes binding to
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DNA are intercalation, complementing the results form viscosity measurements.
3.4. Photocleavage of pBR322 DNA by complexes 1 and 2
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The photoactivated cleavage reaction on plasmid DNA can be monitored by agarose gel
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electrophoresis. As well-known, DNA photocleavage is controlled by relaxation of the supercoiled circular form of plasmid DNA into nicked circular and linear forms. When circular plasmid DNA is
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subject to electrophoresis, relatively fast migration will be observed for the intact supercoil form
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(Form I). If scission occurs on one strand (nicking), the supercoil will relax to generate a
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slower-moving open circular form (Form II). If both strands are cleaved, a linear form (Form III) which migrates between Form I and Form II will be generated [44]. The results of the gel
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electrophoresis separation of pBR322 DNA after incubation with various concentrations of either complex 1 or 2 under irradiation at 365 nm for 30 minutes is presented in Fig. 5. No obvious DNA cleavage was observed for control experiments with DNA alone (A and B, lane 0). In contrast, photoreactions of using the wavelength light of 365 nm (A and B, lanes 1-4) resulted in significant production of nicked DNA with either complex 1 or 2, which suggested that complexes 1 and 2 are efficient DNA-photocleavers under irradiation at 365 nm. For complex 1, the complete conversions of DNA from Form I to Form II could be observed upon increasing the concentration to 15 µM, whereas for complex 2, the same phenomena were occurred at the concentration of 20 µM, suggesting the cleavage reactions may be dependent on the concentrations of each complex. In addition, cleavage of pBR322 DNA with a low concentration (5 µM) of each complex irradiated at 12
ACCEPTED MANUSCRIPT 365 nm under different time is given in Fig. 6, which indicates that the effective DNA cleavage activity of each complex also depends on the irradiation time. These results indicate that: (1) the
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DNA cleavage activity of both complexes not only depends on their concentrations of the
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cleavage activity than complex 1 under the same conditions.
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complexes, but also depends on the irradiation time. (2) Complex 1 exhibits more effective DNA
To identify the nature of the reactive species that are responsible for the photoactivated cleavage
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of the plasmid DNA, the influences of different potentially inhibiting agents were carried out. Fig 7.
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shows typical results obtained with either complex 1 or 2, suggesting that the cleavage of the plasmid was obviously inhibited in the presence of the singlet oxygen (1O2) scavengers (histidine,
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lane 5). This indicates that, for complexes 1 and 2, singlet oxygen (1O2) plays a significant role in
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the photocleavage mechanism and the photoreduction of Ru(II) complexes with concomitant
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hydroxide oxidation is an important step in the DNA cleavage reaction. However, in the presence of the superoxide anion radical inhibitors (superoxide dismutase, lane 4) and hydroxyl-radical (OH•)
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scavengers (mannitol and dimethyl sulfoxide, lanes 2 and 3), respectively, the decease of the amount of DNA cleavage does not occur, compared to that without any inhibitors (lane 1). Therefore, the results indicated that superoxide anion radical and hydroxyl radical (OH ) were not indeed in the DNA cleavage of the Ru(II) complexes under irradiation, the mechanism of DNA cleavage is an oxidative process by generating singlet oxygen. Similar cases are found in other Ru(II) complexes [45].
3.5. Topoisomerases I and II inhibition by complexes 1 and 2 Agarose gel electrophoresis assays were performed to assess the Topo I and IIα inhibitory activities of complexes 1 and 2. The results of Topo I inhibition assay by different concentrations of 13
ACCEPTED MANUSCRIPT either complex 1 or 2 are given in Fig. 8. Fig. 8 (top) indicated that both complexes could inhibit the ability of Topo I to relax negatively supercoiled plasmid DNA, implying that both complexes may
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block the DNA strand passage event of the enzyme, and may serve as inhibitor of Topo I. For
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complexes 1 and 2, the values of IC50 are about 4.5 and 22 µM, respectively, which indicates that
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the Topo I inhibitory activities of the complexes correspond to the DNA-binding affinities of the complexes. In addition, for both complexes, the IC50 values are smaller than that of
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Δ-[Ru(bpy)2(uip)]2+ and Λ-[Ru (bpy)2(uip)]2+(~ 40 µM), whereas for complex 1, the value of IC50 is
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comparable to that of Δ-[Ru(bpy)2(ipad)]2+ (4 µM) and Λ-[Ru(bpy)2(ipad)]2+ (3 µM) [24]. Notably, as DNA intercalators, the complexes can induce constrained negative and unconstrained positive
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superhelical twists in plasmid DNA, resulting in directly altering the topological state of the
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negatively supercoiled DNA substrate. The reason for this is that Topo I could only remove the
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unconstrained positive supercoils. Therefore, the negatively supercoiled DNA product would be identical to the topological state of the original plasmid substrate. In this case, the inhibition of
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enzyme catalysis may also arise in the presence of the complexes as DNA intercalators. To determine whether complexes 1 and 2 interfere with the DNA relaxation reaction by inhibiting Topo I catalysis or by altering the apparent topological state of DNA, the DNA strand passage assay was further carried out. The effects of the complexes on enzyme-catalyzed DNA strand passage were assessed by comparing the rate of relaxation of negatively supercoiled plasmid in the presence of either complex 1 or 2 to the rate of supercoiling of relaxed plasmid in the presence of EB. As shown in Fig. 8 (bottom), the rate of Topo I-poisoned DNA supercoiling in the presence of the Ru(II) complexes were lower than the rate of EB, which was identical to the rate of Topo I-poisoned DNA relaxation in the absence of either complex 1 or 2. These results suggest that complexes 1 and 2 are TopoI poisons but not catalytic inhibitors, similar to that observed for Ru(II) complexes with 14
ACCEPTED MANUSCRIPT phenolic hydroxyl groups [46], while differ from the inhibition mechanisms of Ru(II) anthraquinone complexes [24].
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The Topo IIα inhibition by either complex 1 or 2 was determined by the Topo IIα-mediated
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pBR322 DNA relaxation assay [46]. As can be seen from Fig. 9 (top), both complexes
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dose-dependently inhibited Topo IIα-mediated DNA relaxation. The IC50 values are about 4 and 8 µM for complexes 1 and 2, respectively, which are comparable to that of Topostatin (~ 4 µM) [47],
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[Ru(L)2(fmp)]2+ (L = phen or bpy; ~ 3 µM) [48], whereas significantly lower than that of the widely
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used Topo IIα poison VP-16 (IC50 = 35 mM) [49]. As described for Topo I, the DNA strand passage assay was also used to expore the effects of the Ru(II) complexes on the Topo IIα function from
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their effects on DNA topology. Fig. 9 (bottom) indicates that the religation rates of the relaxed
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plasmid in the presence of either complex 1 or 2 were obvioyusly lower than that in the case of EB,
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suggesting that both complexes can prevent the religation rates of the relaxed plasmid. The results reflect that the inhibition mechanisms of the two Ru(II) complexes are TopoIIα poisons instead of
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catalytic inhibitors, similar to that observed for some Ru(II) complexes such as Ru(L)2(fmp)]2+ [48] and VP-16 [49].
4. Conclusions In
conclusion,
the
DNA binding,
photocleavage
and
topoisomerase
inhibition
of
[Ru(phen)2(pnip)]2+ (1) and [Ru(bpy)2(pnip)]2+ (2) are investigated. DNA-binding studies indicate that the two complexes interact with CT-DNA via an intercalative mode, while complex 1 intercalates into CT-DNA base pairs slightly deeply than complex 2 does. In addition, complex 1 exhibits more efficient plasmid pBR322 DNA photocleavage under irradiation at 365 nm than complex 2, which may be ascribed to the enhanced DNA binding affinity and more singlet oxygen 15
ACCEPTED MANUSCRIPT generation efficiency of complex 1 with respect to complex 2. Furthermore, a topoisomerase inhibition assay suggests that the two Ru(II) complexes are dual Topo I and Topo IIα poisons and
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inhibit Topo I/IIα activity more efficiently than those of many reported inhibitors.
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Acknowledgements
We acknowledge the support of the National Natural Science Foundation of China (21541008
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and 21371146).
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ACCEPTED MANUSCRIPT [25] J. Li, Y. M. Suna, L. J Xie, X. J. He, Lifeng Tan, J. Inorg. Chem. 143 (2015) 56–63. [26] (a) S. Banerjee, E.B. Veale, C.M. Phelan, S.A. Murphy, G.M. Tocci, L.J. Gillespie, D.O.
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ACCEPTED MANUSCRIPT N.A.P. Kane-Maguire, Inorg. Chem. 49 (2010) 839−848. [38] (a) S. Satyanarayana, J.C. Dabrowiak, J.B. Chaires, Biochemistry 31 (1992) 9319–9324; (b) S.
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ACCEPTED MANUSCRIPT Captions for Schemes and Figures Fig. 1. Chemical structures. of complexes 1 and 2. Fig. 2. Representative absorption spectral changes of complexes 1 (A) and 2 (B) in Tris–HCl buffer
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upon addition of CT-DNA. [Complex 1] = [Complex 2 ] = 20 uM; [DNA] = 0–186 μM. Arrows indicate the change inabsorbance upon increasing the DNA concentration.
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Fig. 3. Representative fluorescence emission spectra of complexes 1 (A) and 2 (B) treated with CT-DNA. [Complex 1] = [Complex 2 ] = 2.0 uM; [DNA] = 0–8.7 μM.. Fig. 4. Effect of increasing amounts of EB (●), [Ru(bpy)3]2+(▼), complexes 1 (■ ) and 2 (○) on the
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relative viscosity of CT-DNA. Total DNA concentration: 0.5 mM, temperature is (28 ± 0.1) oC. Fig. 5. Photoactivated cleavage of pBR322 DNA in the presence of different concentrations of
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either (A) complex 1 or (B) complex 2, after irradiation at 365 nm for 30 min. Lanes 0–5 are the different concentrations of complex 1 or 2: (0) 0; (1) 5; (2) 10; (3) 15; (4) 20 µM, respectively.
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Fig. 6. Photoactivated cleavage of pBR322 DNA in the presence of (A) complex 1 (5 µM) or (B)
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complex 2 (5 µM), light after irradiation at 365 nm for different time. Lane 0, DNA alone and irradiation for 60 min. Lanes 1–6 are the different irradiation time of pBR322 DNA with Ru(II)
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complexes: (1) 10; (2) 20; (3) 30; (4) 40; (5) 50; (6) 60 min, respectively. Fig. 7. Photoactivated cleavage of pBR322 DNA by either (A) complex 1 (25 µM) or (B) complex 2 (25 µM) in the presence of different inhibitors after irradiation at 365 nm for 30 min. Lane 0,
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DNA alone; lane 1, no inhibitor; lane 2, in the presence of mannitol (100 mM); lane 3, in the presence of dimethyl sulfoxide (200 mM); lane 4, in the presence of superoxide dismutase (1000 units/mL); lane 5, in the presence of histidine (20 mM). the superoxide anion radical inhibitors (superoxide dismutase, lane 4) and hydroxyl-radical (OH•) scavengers (mannitol and dimethyl sulfoxide, lanes 2 and 3), Fig. 8. Top: effects of different concentrations of either complex 1 or 2 on the activity of Topo I (top); bottom: the time dependence of Topo I DNA strand passage assays in the presence of EB (30 µM), complexes 1 (3 µM) and 2 (10 µM), respectively (bottom). Fig. 9. Top: Effects of different concentrations of either complex 1 or 2 on the activity of Topo IIα; bottom: the time dependence of Topo IIα DNA strand passage assays in the presence of EB (30 µM), complex 1 (2 µM) or 2 (4 µM), respectively. 21
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Table 1 Hypochromicity (H) , bathochromic shifts, binding constants (Kb) and average binding site size (s) of complex 1 and 2. λmax, free (nm)
Δλa (nm)
λmax, bound (nm)
Hb(%)
Kb (× 106 M-1)c
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Complex
s
462 460
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459 457
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7.95 ± 0.28 1.24 0.13 4.50 0.15 1.01 0.12 a Δλ represents the difference in MLCT band of the metal complex between free and completely bound RNA states. b H represents the hypochromism at MLCT band, as defined by H% = 100% (Afree − Abound)/Afree (A represents the absorbance). c Kb was determined by monitoring the changes of absorption at the MLCT band.
1 2
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Fig. 1.
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Absorbance
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0.8 0.6 0.4 0.2 0.0 0
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6 9 12 15 18 5 10 x [DNA]
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1.0 : DNA alone : [Complex 2]:[DNA]=1:10 : [Complex 1]:[DNA]=1:10
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Results suggest that complexes 1 and 2 can bind to DNA via an intercalative mode and efficiently photocleavage pBR322 DNA in vitro under irradiation. In addition, the results of topoisomerase inhibition and DNA strand passage assay reflect that both complexes are efficient dual dual poisons of topoisomerases I and IIα.
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ACCEPTED MANUSCRIPT Highlights ► DNA binding, photocleavage and topoisomerase inhibitory activity of complexes 1 and 2 are
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evaluated. ► Complexes 1 and 2 bind to DNA via intercalation. ► Complex 1 exhibits more
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2 are efficient dual poisons of topoisomerases I and IIα.
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effective DNA cleavage activity than complexes 2 under the same conditions. ► Complexes 1 and
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S0162-0134(16)30110-6 doi: 10.1016/j.jinorgbio.2016.04.028 JIB 9986
To appear in:
Journal of Inorganic Biochemistry
Received date: Revised date: Accepted date:
28 December 2015 12 April 2016 18 April 2016
Please cite this article as: Yanmei Sun, Jia Li, Hong Zhao, Lifeng Tan, Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I and IIα, Journal of Inorganic Biochemistry (2016), doi: 10.1016/j.jinorgbio.2016.04.028
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ACCEPTED MANUSCRIPT Ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group as DNA binder, photonuclease, and dual inhibitors of topoisomerases I
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and IIα
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Yanmei Sun a,1, Jia Li a,1, Hong Zhao a, Lifeng Tan*,b a College of Chemistry, Xiangtan University, Xiangtan, P. R. China
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b Key Lab of Environment-friendly Chemistry and Application in Ministry of Education, Xiangtan
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University, Xiangtan, P. R. China
* Corresponding author at: College of Chemistry, Xiangtan University, Xiangtan 411105, PR China. Tel.: +86 731 58293997; Fax: +86 731 58293477. E-mail address: [email protected] (L.-F. Tan). 1
These two authors are both first authors.
ABSTRACT Two ruthenium(II) polypyridyl complexes containing 1,8-naphthalimide group as DNA binders, 1
ACCEPTED MANUSCRIPT photonucleases, and inhibitors of topoisomerases I and IIα are evaluated. The binding properties of [Ru(phen)2(pnip)]2+ {1; phen =1,10-phenanthroline; pnip = 12-[N-(p-phenyl)-1,8-napthalimide]-
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imidazo[4',5'-f][1,10]phenanthroline} and [Ru(bpy)2(pnip)]2+ (2; bpy = 2,2′-bipyridine) with calf
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thymus DNA increases with increasing the bulkiness and hydrophobic character of ancillary ligands,
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although the two complexes possess high affinities for DNA via intercalation. Moreover, photoirradiation (λ = 365 nm) of the two complexes are found to induce strand cleavage of closed
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circular pBR322 plasmid DNA via singlet oxygen mechanism, while complex 1 displays more
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effective photocleavage activity than complex 2 under the same conditions. Topoisomerase inhibition and DNA strand passage assay reflect that complexes 1 and 2 are efficient dual poisons of
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topoisomerases I and IIα.
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Keywords: Ruthenium(II) complexes; DNA binding; Photocleavage; Topoisomerase inhibition
2
ACCEPTED MANUSCRIPT 1. Introduction DNA topoisomerases are essential for the processes of chromosomal segregation and relaxation
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of DNA during replication and transcription, and may also be required for recombination [1]. Two
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types of DNA topoisomerases have been isolated from prokaryotes and eukaryote. In general,
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topoisomerase I (Topo I) catalyzes the relaxation of superhelical DNA by generating a transient single strand nick in the duplex through cycles of cleavage and religation [2], while topoisomerase
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II (Topo II) mediates the ATP-dependent induction of coordinated nicks in both strands of the DNA
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duplex through which another DNA segment is passed [3]. Under normal conditions, the step of DNA relegation is much faster than that of DNA cleavage, which may be tolerated by the cell.
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However, conditions that significantly change either the physiological concentration or the lifetime
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of the breaks are responsible for DNA alterations, playing a crucial role in inhibiting cell cycle
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progression [4]. Therefore, Topoisomerases are the cellular targets of clinically important anticancer and antibacterial drugs, and inhibition of topoisomerases has been considered as an
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effective strategy for the design of many anticancer agents [5]. In recent years, numerous natural and synthetic compounds have been chosen to detect the inhibition of topoisomerase activity [6–10]. To date, some topoisomerase inhibitors, such as camptothecin (CPT) and its derivatives that target Topo I [11] and etoposide (VP-16) [12], doxorubicin [13] and mitoxantrone [14] which target Topo II, have been clinically used as potent anticancer drugs. These agents have achieved great success in clinical cancer chemotherapy, while single inhibitors of topoisomerases are limited by several important negative consequences [15]. Furhtermore, the emergence of resistance phenomena to Topo I inhibitors is often accompanied by a concomitant rise at the level of Topo II expression and viceversa, resulting in the failure of clinical therapies [15]. In this regard, a single compound able to inhibit both Topo I and II may 3
ACCEPTED MANUSCRIPT present the advantage of improving antitopoisomerase activity, with reduced toxic side effects, with respect to the combination of two inhibitors [16]. Several classes of dual topos I and II inhibitors
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have been identified and described over the last decades [17–21]. Some of these dual inhibitors,
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such as Tafluposide and Batracylin, have been evaluated in clinical trials. Most of such studies at
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present, however, have been mainly revolved around organic compounds and, to a far lesser extent, around metal complexes [22]. In comparison with organic molecules, the interest in the field of
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Ru(II) polypyridyl complexes−nucleic acid interactions has burgeoned due to Ru(II) polypyridyl
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complexes with rich photochemical properties and varied coordination forms [23]. However, surprisingly and in contrast to studies on Ru(II) polypyridyl complexes−nucleic acid interactions,
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investigations of the inhibition of topoisomerases activity by Ru(II) polypyridyl complexes and the
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relationship between the structures of Ru(II) complexes and the enzymatic inhibition activities are
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very scarce [24]. Therefore, studies on the interaction between polypyridyl-based Ru(II) complexes and topoisomerases are very important for developing novel antitumor drugs and elucidating the
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underlying molecular mechanism.
Recently, we reported the synthesis and RNA-binding properties of Ru(II) polypyridyl complexes with 1,8-naphthalimide group (Fig. 1), [Ru(phen)2(pnip)]2+ (1; phen = 1,10-phenanthroline, pnip = 2-[N-(p-phenyl)-1,8-napthalimide]imidazo[4',5'-f][1,10]phenanthroline) and [Ru(bpy)2(pnip)]2+ (2; bpy = 2,2′-bipyridine), finding that the two complexes displayed high binding affinities for the RNA triplex as well as increasing the stability of the Hoogsteen base-paired strand of the triplex obviously [25]. In addition, we note that the development of functional 1,8-naphthalimide and its derivatives as DNA binders, photonucleases and enzyme inhibitors is a fast growing area [26]. However, biological investigations of ruthenium(II) polypyridyl complexes with 1,8-naphthalimide group have attracted little attention relatively [26c]. 4
ACCEPTED MANUSCRIPT Considering all the above, this paper deals with the DNA binding, photo-induced DNA cleavage, and topoisomerase inhibitory activity of complexes 1 and 2. We hope that this work will aid in
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advancing our knowledge of the interaction between polypyridyl-based Ru(II) complexes and DNA,
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as well as laying the foundation for the rational design of dual topoisomerases I and II inhibitors.
2. Experimental sections
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2.1. Materials
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Complexes 1 and 2 were synthesized according to the literature methods [25]. Calf thymus DNA (CT-DNA) approximately 200 base pairs in average length was purchased from the Sino-American
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Biotechnology Company. DNA topoisomerase I from calf thymus together with human
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topoisomerase IIa were purchased from TopoGen Inc. Tris–HCl buffer A (5 mM Tris–HCl, 50 mM
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NaCl, pH 7.0) (Tris = tris(hydroxymethyl)aminomethane) solution was prepared using doubly distilled water. A solution of CT-DNA in the buffer gave a ratio of UV (UV = ultraviolet)
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absorbance at 260 and 280 nm of ca. 1.8–1.9:1, indicating that the DNA was sufficiently free of protein [27]. The DNA concentration per nucleotide was determined by absorption spectroscopy using the molar absorption coefficient (6600 M-1 cm-1) at 260 nm [28].
2.2. DNA-binding experiments UV–vis spectra were collected using a PerkinElmer Lambda 25 spectrophotometer at 20 oC. A typical titration of each Ru(II) complex in buffer A was performed by using a fixed concentration of either complex 1 or 2, to which the DNA stock solution was gradually added up to saturation. After each addition, the solution was mixed and allowed to reequilibrate for 5 min before the absorption spectra were recorded. The intrinsic binding constant Kb to DNA was determined the changes of 5
ACCEPTED MANUSCRIPT MLCT (MLCT = metal to ligand charge transfer) band by using the following equation [29]:
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(1a)
(1b)
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b = 1+ K b C t + K b [DNA]/(2s)
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b b2 2 Kb2Ct DNA / s a f b f 2 KCt
where [DNA] is the concentration of DNA in base pairs and the apparent absorption
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coefficients εa, εf, and εb correspond to A obsd /[Ru], the extinction coefficient for the free Ru(II) complex, and the extinction coefficient for a Ru(II) complex in the fully bound form,
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respectively. Kb is the equilibrium binding constant in M-1, Ct is the total metal complex
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2.3. Thermal denaturation studies
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concentration, and s is the binding site size.
PerkinElmer Lambda 25
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The thermal denaturation studies were performed on a
spectrophotometer equipped with a water-circulating bath. The temperature was ramped from 50 to
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90 °C with an increasing rate of 1.0 °C min−1. The melting curves were obtained by measuring the absorption of DNA solution at 260 nm in the absence and presence of the ruthenium complex (dissolved in buffer A) as a function of temperature. The data were presented as (A − A0)/(Af − A0) versus temperature, where A, A0, and Af are the observed, the initial, and the final absorbance at 260 nm, respectively.
2.4. Viscosity studies Viscosity measurements were carried out using an ubbelohde viscometer maintained at 25.0 ± 0.1 °C in buffer A. The flow time was measured using a digital stopwatch, and each sample was measured three times. Relative viscosities for the triplex RNA in either the absence or presence of 6
ACCEPTED MANUSCRIPT metal complexes were calculated according to literature procedures reported earlierand an average
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flow time was calculated. Data are presented as (η/η0)1/3 versus the binding ratio [30].
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2.5. DNA photocleavage experiments
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For the gel electrophoresis experiment, supercoiled pBR322DNA (0.05 μg) was treated with different concentration of each Ru(II) complex in buffer B (50 mM Tris-HCl, 18 mM NaCl, pH 7.2),
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and then the solution was irradiated with a UV lamp (365 nm, 10 W) at room temperature. The samples were analyzed by electrophoresis for 1 h at 80 V in buffer C (89 mM Tris, 89 mM boron
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hydroxide, 2 mM EDTA) (EDTA = ethylene diamine tetraacetie acid) containing 0.75% agarose gel. The gel was stained with EB (EB = ethidium bromide;1 mg/mL), photographed under UV light and
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D
then analysed on the FluorChem FC2. The percentage of photocleavage (C) was determined according to the following equation:
DII 2 DIII 100% DI DII 2 DIII
CE P
C
(2)
AC
Where DI, DII and DIII are the relative density of supercoiled pBR322 DNA existing in the form of supercoil form (Form I), nicking form (Form II) and linear form (Form III) in the absence and presence of either complex 1 or 2, respectively.
2.6. Topoisomerase inhibition assay The relaxation assay was performed in buffer D (35 mM Tris-HCl, 72 mM KCl, 5 mM MgCl2, 5 mM dithiothreitol, 2 mM spermidine, 0.1 mg/ml bovine serum albumin, pH 8.0) containing 0.05 μg pBR322 DNA and 1 unit of Topo I in the absence and presence of either complex 1 or 2. Control experiments contained either pBR322 DNA alone or pBR322 DNA treated with Topo I only. The reaction mixture (total volume = 10 μL) was incubated at 37 oC for 30 min and quenched by 7
ACCEPTED MANUSCRIPT addition of 4 μL of loading buffer containing 0.25% bromophenol blue, 4.5% SDS and 45% glycerol. The DNA was fractionated by agarose gel electrophoresis by 1% agarose in TBE at 80 V
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for 1 h. The gel was then stained with EB (1 µg/mL), visualized using a UV illuminator and
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eventually photographed under UV light. In addition, IC50 values were the concentrations of the
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inhibitor that prevented 50% of the supercoiled DNA from being converted into relaxed DNA, which were calculated by the midpoint concentration for either complex 1 or 2 induced DNA
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unwinding.
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Topo IIa functional activity was assayed in buffer E (10 mM Tris-HCl, 50 mM NaCl, 50 mM KCl, 5.0 mM MgCl2, 0.1 mM Na2H2EDTA, 15 μg/mL bovine serum albumin, 1.0 mM ATP, pH 7.9)
D
containing 0.05 μg pBR322DNA and 4 units of Topo IIa in the absence and presence of either
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complex 1 or 2. The reaction mixture was incubated at 30 oC for 15 min. Reactions were stopped,
CE P
processed, and then subjected to gel electrophoresis.
2.7. DNA strand passage assay
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The DNA strand passage activity of Topo I was determined by investigating the ability of Topo I to relax negatively supercoiled plasmid DNA [29] or to supercoil plasmid DNA [31]. Reactions contained 0.05 μg relaxed or supercoiled pBR322 plasmid DNA in buffer D (total volume = 10 μL). Assays were carried out in the absence of either complex 1 or 2 or in the presence of 25 μM Ru(II) complexes or EB. Following a 5-min incubation, 1 unit of Topo I was added, and reactions were incubated up to 15 min at 37 oC. Reactions were stopped, processed, and subjected to gel electrophoresis as described above. Regarding the DNA strand passage activity of Topo IIa, reactions contained relaxed or supercoiled 0.05 μg pBR322 plasmid DNA and 4 units of Topo IIa in buffer E (total volume = 10 μL). Following a 5-min incubation, 4 units of Topo IIa was added and 8
ACCEPTED MANUSCRIPT then reactions were incubated up to 15 min at 30 oC. The rest of the operations were the same as
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described above.
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3. Results and discussion
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3.1. DNA binding studies
The interaction of complexes 1 and 2 with CT-DNA was primarily examined by absorption
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spectra titration experiments. Binding of a complex to DNA through a π-π stacking interaction
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between an aromatic chromophore of a complex and the base pairs of DNA typically results in hypochromism and a red shift [32]. Fig. 2 provides the changes in UV-Vis absorption spectra when
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complexes 1 and 2 bound to CT-DNA, and the representing data are summarized in Table 1. Fig. 2
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indicated that the addition of CT-DNA to the solution of either complex 1 or 2 in a Tris buffer led to
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obvious hypochromism in the UV/Vis absorption spectrum with moderate bathochromic spectral shift. For complex 1, the hypochromicity at 459 nm reached about 23% with a red shift of 3 nm at a
AC
[DNA]/[Comp. 1] ratio of 5.1, whereas for complex 2, the hypochromicity at 457 nm showed about 17% at a [DNA]/[Comp. 2] ratio of 6.0 with a red shift of 3 nm. The spectral characteristics suggested that complexes 1 and 2 could interact with CT-DNA. To further elucidate the binding strength of both complexes, the DNA binding constants (Kb) were estimated from the changes of the absorbance at 459 and 457 nm for complexes 1 and 2, respectively. The Kb values of complexes 1 and 2 are (7.95 ± 0.28) ×106 M-1 (s = 1.24 0.13) and (4.50 0.15) × 106 M-1 (s = 1.01 0.12), respectively. The Kb values of both complexes are comparable to that determined for [Ru(phen)2dppz]2+ [33, [Ru(dtzp)(ptn)]2+ [34] and the known DNA intercalator ethidium bromide [35], while stronger than that of [Ru(bpy)3]2+ (4.7 103 M-1)
9
ACCEPTED MANUSCRIPT and [Ru(phen)3]2+ (5.4 103 M-1) [36]. These data indicate that the size and the shape of the intercalated ligand of Ru(II) complexes have a significant effect on the binding strength, and the
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most suitable intercalating ligand leads to the highest affinity of complexes with DNA. In addition,
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the Kb value of complex 1 was slightly greater than that of complex 2, which suggested that the
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binding of complex 1 with DNA is tighter than that of complex 2. In general, the binding reaction of small molecule with nucleic acids is driven primarily by hydrophobic interactions [37]. For
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complexes 1 and 2, as the ancillary ligand progresses from bpy to phen studied, the bulkiness and
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hydrophobic character of the ligand is increased. Consequently, the hydrophobic transfer of the large aromatic complexes 1 from solution into the DNA binding site caused by the ancillary ligand
D
phen is easier to achieve, resulting in the binding sites effectively overlapping with each other.
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Therefore, the effects of ancillary ligands (phen and bpy) on the binding characteristics of
CE P
complexes 1 and 2 with DNA should not be ignored, which is likely the main factor affecting the interactions of the two metal complexes with CT-DNA.
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3.2. Determination of the binding mode by viscosity studies To earnestly clarify the binding modes of the complexes and CT-DNA, viscosity measurements were performed. Hydrodynamic measurements that are sensitive to changes in length (i.e. viscosity and sedimentation) are regarded as the least ambiguous and the most critical tests of a binding model in solution in the absence of crystallographic or NMR structural data [38]. In general, the viscosity of double-stranded DNA increases when a complex binds to DNA through a classical intercalating mode [38]. In contrast, the viscosity of DNA will decline when the binding mode is a partial or non-classical intercalation, because this mode could bend (or kink) the DNA helix and concomitantly reduce its effective length [39]. However, it remains unchanged when a complex 10
ACCEPTED MANUSCRIPT binds DNA in an electrostatic mode [38]. The effects of complexes 1 and 2 together with EB and [Ru(bpy)3]2+ on the viscosity of rod-like DNA are presented in Fig. 3. As expected, EB, which is a
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well-known DNA intercalator, strongly increases the relative viscosity by lengthening the DNA
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double helix through intercalation. While for complex [Ru(bpy)3]2+, which has been known to bind
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DNA through electrostatic interactions, exhibited essentially no effect on the viscosity of DNA [40]. It is evident that the binding of either complex 1 or 2 can increase in the relative viscosity of the
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DNA solution upon increasing the amounts of the complex. The increased degree of viscosity,
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which may depend on its affinity to DNA, follows the order of EB > 1 > 2 > [Ru(bpy)3]2+. The results further suggest that the binding modes of complexes 1 and 2 are intercalation, and complex 1
D
intercalates into CT-DNA base pairs more deeply than complex 2 does, complementing the above
CE P
TE
results.
3.3. Thermal denaturation studies
AC
DNA melting experiments are useful to determine the extent of intercalation, because the intercalation of the complex into DNA base pairs causes stabilisation of the base stacking [41] and, therefore, raises the melting temperature of the double-stranded DNA [42]. It is well accepted that when the temperature of the solution increases, the double-stranded DNA gradually dissociates into single strands, which generates a hyperchromic effect on the absorption spectra of the DNA bases (λmax 260 nm). To identify this transition process, the melting temperature (Tm), which is defined as the temperature where half of the total base pairs are unbonded, is usually introduced. Generally, the intercalation of a complex into DNA generally results in a considerable increase of Tm [40]. The melting curve of CT-DNA in the absence and presence of each complex is presented in Fig. 4. The Tm of Ru(II) complex-free CT-DNA was determined to be 64.5 oC. In the presence of either 11
ACCEPTED MANUSCRIPT complex 1 or 2, the Tm increases successively and reaches 78.5 and 74.9 oC, respectively, at a [Ru]/[DNA] ratio of 1 : 10. The ΔTm values (14.0 and 10.4 oC) of 1/2–DNA adducts are larger than
T
those of some Ru(II) intercalators [43], which reveals that the modes of both complexes binding to
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DNA are intercalation, complementing the results form viscosity measurements.
3.4. Photocleavage of pBR322 DNA by complexes 1 and 2
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The photoactivated cleavage reaction on plasmid DNA can be monitored by agarose gel
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electrophoresis. As well-known, DNA photocleavage is controlled by relaxation of the supercoiled circular form of plasmid DNA into nicked circular and linear forms. When circular plasmid DNA is
D
subject to electrophoresis, relatively fast migration will be observed for the intact supercoil form
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(Form I). If scission occurs on one strand (nicking), the supercoil will relax to generate a
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slower-moving open circular form (Form II). If both strands are cleaved, a linear form (Form III) which migrates between Form I and Form II will be generated [44]. The results of the gel
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electrophoresis separation of pBR322 DNA after incubation with various concentrations of either complex 1 or 2 under irradiation at 365 nm for 30 minutes is presented in Fig. 5. No obvious DNA cleavage was observed for control experiments with DNA alone (A and B, lane 0). In contrast, photoreactions of using the wavelength light of 365 nm (A and B, lanes 1-4) resulted in significant production of nicked DNA with either complex 1 or 2, which suggested that complexes 1 and 2 are efficient DNA-photocleavers under irradiation at 365 nm. For complex 1, the complete conversions of DNA from Form I to Form II could be observed upon increasing the concentration to 15 µM, whereas for complex 2, the same phenomena were occurred at the concentration of 20 µM, suggesting the cleavage reactions may be dependent on the concentrations of each complex. In addition, cleavage of pBR322 DNA with a low concentration (5 µM) of each complex irradiated at 12
ACCEPTED MANUSCRIPT 365 nm under different time is given in Fig. 6, which indicates that the effective DNA cleavage activity of each complex also depends on the irradiation time. These results indicate that: (1) the
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DNA cleavage activity of both complexes not only depends on their concentrations of the
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cleavage activity than complex 1 under the same conditions.
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complexes, but also depends on the irradiation time. (2) Complex 1 exhibits more effective DNA
To identify the nature of the reactive species that are responsible for the photoactivated cleavage
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of the plasmid DNA, the influences of different potentially inhibiting agents were carried out. Fig 7.
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shows typical results obtained with either complex 1 or 2, suggesting that the cleavage of the plasmid was obviously inhibited in the presence of the singlet oxygen (1O2) scavengers (histidine,
D
lane 5). This indicates that, for complexes 1 and 2, singlet oxygen (1O2) plays a significant role in
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the photocleavage mechanism and the photoreduction of Ru(II) complexes with concomitant
CE P
hydroxide oxidation is an important step in the DNA cleavage reaction. However, in the presence of the superoxide anion radical inhibitors (superoxide dismutase, lane 4) and hydroxyl-radical (OH•)
AC
scavengers (mannitol and dimethyl sulfoxide, lanes 2 and 3), respectively, the decease of the amount of DNA cleavage does not occur, compared to that without any inhibitors (lane 1). Therefore, the results indicated that superoxide anion radical and hydroxyl radical (OH ) were not indeed in the DNA cleavage of the Ru(II) complexes under irradiation, the mechanism of DNA cleavage is an oxidative process by generating singlet oxygen. Similar cases are found in other Ru(II) complexes [45].
3.5. Topoisomerases I and II inhibition by complexes 1 and 2 Agarose gel electrophoresis assays were performed to assess the Topo I and IIα inhibitory activities of complexes 1 and 2. The results of Topo I inhibition assay by different concentrations of 13
ACCEPTED MANUSCRIPT either complex 1 or 2 are given in Fig. 8. Fig. 8 (top) indicated that both complexes could inhibit the ability of Topo I to relax negatively supercoiled plasmid DNA, implying that both complexes may
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block the DNA strand passage event of the enzyme, and may serve as inhibitor of Topo I. For
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complexes 1 and 2, the values of IC50 are about 4.5 and 22 µM, respectively, which indicates that
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the Topo I inhibitory activities of the complexes correspond to the DNA-binding affinities of the complexes. In addition, for both complexes, the IC50 values are smaller than that of
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Δ-[Ru(bpy)2(uip)]2+ and Λ-[Ru (bpy)2(uip)]2+(~ 40 µM), whereas for complex 1, the value of IC50 is
MA
comparable to that of Δ-[Ru(bpy)2(ipad)]2+ (4 µM) and Λ-[Ru(bpy)2(ipad)]2+ (3 µM) [24]. Notably, as DNA intercalators, the complexes can induce constrained negative and unconstrained positive
D
superhelical twists in plasmid DNA, resulting in directly altering the topological state of the
TE
negatively supercoiled DNA substrate. The reason for this is that Topo I could only remove the
CE P
unconstrained positive supercoils. Therefore, the negatively supercoiled DNA product would be identical to the topological state of the original plasmid substrate. In this case, the inhibition of
AC
enzyme catalysis may also arise in the presence of the complexes as DNA intercalators. To determine whether complexes 1 and 2 interfere with the DNA relaxation reaction by inhibiting Topo I catalysis or by altering the apparent topological state of DNA, the DNA strand passage assay was further carried out. The effects of the complexes on enzyme-catalyzed DNA strand passage were assessed by comparing the rate of relaxation of negatively supercoiled plasmid in the presence of either complex 1 or 2 to the rate of supercoiling of relaxed plasmid in the presence of EB. As shown in Fig. 8 (bottom), the rate of Topo I-poisoned DNA supercoiling in the presence of the Ru(II) complexes were lower than the rate of EB, which was identical to the rate of Topo I-poisoned DNA relaxation in the absence of either complex 1 or 2. These results suggest that complexes 1 and 2 are TopoI poisons but not catalytic inhibitors, similar to that observed for Ru(II) complexes with 14
ACCEPTED MANUSCRIPT phenolic hydroxyl groups [46], while differ from the inhibition mechanisms of Ru(II) anthraquinone complexes [24].
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The Topo IIα inhibition by either complex 1 or 2 was determined by the Topo IIα-mediated
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pBR322 DNA relaxation assay [46]. As can be seen from Fig. 9 (top), both complexes
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dose-dependently inhibited Topo IIα-mediated DNA relaxation. The IC50 values are about 4 and 8 µM for complexes 1 and 2, respectively, which are comparable to that of Topostatin (~ 4 µM) [47],
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[Ru(L)2(fmp)]2+ (L = phen or bpy; ~ 3 µM) [48], whereas significantly lower than that of the widely
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used Topo IIα poison VP-16 (IC50 = 35 mM) [49]. As described for Topo I, the DNA strand passage assay was also used to expore the effects of the Ru(II) complexes on the Topo IIα function from
D
their effects on DNA topology. Fig. 9 (bottom) indicates that the religation rates of the relaxed
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plasmid in the presence of either complex 1 or 2 were obvioyusly lower than that in the case of EB,
CE P
suggesting that both complexes can prevent the religation rates of the relaxed plasmid. The results reflect that the inhibition mechanisms of the two Ru(II) complexes are TopoIIα poisons instead of
AC
catalytic inhibitors, similar to that observed for some Ru(II) complexes such as Ru(L)2(fmp)]2+ [48] and VP-16 [49].
4. Conclusions In
conclusion,
the
DNA binding,
photocleavage
and
topoisomerase
inhibition
of
[Ru(phen)2(pnip)]2+ (1) and [Ru(bpy)2(pnip)]2+ (2) are investigated. DNA-binding studies indicate that the two complexes interact with CT-DNA via an intercalative mode, while complex 1 intercalates into CT-DNA base pairs slightly deeply than complex 2 does. In addition, complex 1 exhibits more efficient plasmid pBR322 DNA photocleavage under irradiation at 365 nm than complex 2, which may be ascribed to the enhanced DNA binding affinity and more singlet oxygen 15
ACCEPTED MANUSCRIPT generation efficiency of complex 1 with respect to complex 2. Furthermore, a topoisomerase inhibition assay suggests that the two Ru(II) complexes are dual Topo I and Topo IIα poisons and
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inhibit Topo I/IIα activity more efficiently than those of many reported inhibitors.
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Acknowledgements
We acknowledge the support of the National Natural Science Foundation of China (21541008
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and 21371146).
16
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ACCEPTED MANUSCRIPT Captions for Schemes and Figures Fig. 1. Chemical structures. of complexes 1 and 2. Fig. 2. Representative absorption spectral changes of complexes 1 (A) and 2 (B) in Tris–HCl buffer
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upon addition of CT-DNA. [Complex 1] = [Complex 2 ] = 20 uM; [DNA] = 0–186 μM. Arrows indicate the change inabsorbance upon increasing the DNA concentration.
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Fig. 3. Representative fluorescence emission spectra of complexes 1 (A) and 2 (B) treated with CT-DNA. [Complex 1] = [Complex 2 ] = 2.0 uM; [DNA] = 0–8.7 μM.. Fig. 4. Effect of increasing amounts of EB (●), [Ru(bpy)3]2+(▼), complexes 1 (■ ) and 2 (○) on the
NU
relative viscosity of CT-DNA. Total DNA concentration: 0.5 mM, temperature is (28 ± 0.1) oC. Fig. 5. Photoactivated cleavage of pBR322 DNA in the presence of different concentrations of
MA
either (A) complex 1 or (B) complex 2, after irradiation at 365 nm for 30 min. Lanes 0–5 are the different concentrations of complex 1 or 2: (0) 0; (1) 5; (2) 10; (3) 15; (4) 20 µM, respectively.
D
Fig. 6. Photoactivated cleavage of pBR322 DNA in the presence of (A) complex 1 (5 µM) or (B)
TE
complex 2 (5 µM), light after irradiation at 365 nm for different time. Lane 0, DNA alone and irradiation for 60 min. Lanes 1–6 are the different irradiation time of pBR322 DNA with Ru(II)
CE P
complexes: (1) 10; (2) 20; (3) 30; (4) 40; (5) 50; (6) 60 min, respectively. Fig. 7. Photoactivated cleavage of pBR322 DNA by either (A) complex 1 (25 µM) or (B) complex 2 (25 µM) in the presence of different inhibitors after irradiation at 365 nm for 30 min. Lane 0,
AC
DNA alone; lane 1, no inhibitor; lane 2, in the presence of mannitol (100 mM); lane 3, in the presence of dimethyl sulfoxide (200 mM); lane 4, in the presence of superoxide dismutase (1000 units/mL); lane 5, in the presence of histidine (20 mM). the superoxide anion radical inhibitors (superoxide dismutase, lane 4) and hydroxyl-radical (OH•) scavengers (mannitol and dimethyl sulfoxide, lanes 2 and 3), Fig. 8. Top: effects of different concentrations of either complex 1 or 2 on the activity of Topo I (top); bottom: the time dependence of Topo I DNA strand passage assays in the presence of EB (30 µM), complexes 1 (3 µM) and 2 (10 µM), respectively (bottom). Fig. 9. Top: Effects of different concentrations of either complex 1 or 2 on the activity of Topo IIα; bottom: the time dependence of Topo IIα DNA strand passage assays in the presence of EB (30 µM), complex 1 (2 µM) or 2 (4 µM), respectively. 21
ACCEPTED MANUSCRIPT
Table 1 Hypochromicity (H) , bathochromic shifts, binding constants (Kb) and average binding site size (s) of complex 1 and 2. λmax, free (nm)
Δλa (nm)
λmax, bound (nm)
Hb(%)
Kb (× 106 M-1)c
T
Complex
s
462 460
3 3
23 17
AC
CE P
TE
D
MA
NU
SC R
459 457
IP
7.95 ± 0.28 1.24 0.13 4.50 0.15 1.01 0.12 a Δλ represents the difference in MLCT band of the metal complex between free and completely bound RNA states. b H represents the hypochromism at MLCT band, as defined by H% = 100% (Afree − Abound)/Afree (A represents the absorbance). c Kb was determined by monitoring the changes of absorption at the MLCT band.
1 2
22
ACCEPTED MANUSCRIPT 2+
2+ N N
N
H N
N
N
Ru N
N
O
N
H N
N
N
Ru
N
N
O
O N O
N
IP
T
N
N
Complex 2
AC
CE P
TE
D
MA
NU
SC R
Complex 1
23
Fig. 1.
ACCEPTED MANUSCRIPT
A
1.0 0.6 0.4
T
1.6
0.8
0.2 0.0
1.2
0
5
0.8
20
NU
0.4
250 300 350 400 450 500 550 600 Wavelength (nm)
AC
1.0
1.0 (a-f)/(b-f)
CE P
Absorbance
1.5
B
TE
2.0
D
MA
0.0
10 15 5 10 x [DNA]
SC R
Absorbance
2.0
IP
(a-f)/(b-f)
2.4
0.8 0.6 0.4 0.2 0.0 0
3
6 9 12 15 18 5 10 x [DNA]
0.5 0.0
250 300 350 400 450 500 550 600 Wavelength (nm)
Fig. 2.
24
T
ACCEPTED MANUSCRIPT
SC R
IP
1.28
(/)
1/3
1.20
NU
1.12
0.02
0.04 0.06 [Ru]/[DNA]
0.08
TE
D
0.00
MA
1.04
AC
CE P
Fig. 3.
25
ACCEPTED MANUSCRIPT
1.0 : DNA alone : [Complex 2]:[DNA]=1:10 : [Complex 1]:[DNA]=1:10
T IP
0.6 0.4
SC R
(A-A0)/(Af-A0)
0.8
0.2 0.0
NU
50 55 60 65 70 75 80 85 90 o
AC
CE P
TE
D
MA
T ( C)
Fig. 4.
26
IP
T
ACCEPTED MANUSCRIPT
Form II
(A)
Lane
0
1
2
SC R
Form I
3
4
Form II
0
1
2
3
Form I
4
CE P
TE
D
MA
Lane
NU
(B)
AC
Fig. 5.
27
ACCEPTED MANUSCRIPT
T
Form II (A)
0
1
2
3
4
5
6
SC R
Lane
IP
Form I
Form II
(B)
Form I
0
1
2
3
4
5
6
D
MA
NU
Lane
AC
CE P
TE
Fig. 6.
28
ACCEPTED MANUSCRIPT
Form II
T
(A)
0
1
2
3
4
1
2
3
4
Form II Form I
5
MA
0
NU
(B)
Lane
5
SC R
Lane
IP
Form I
AC
CE P
TE
D
Fig. 7.
29
ACCEPTED MANUSCRIPT Complex 1
Topo I
4
8
12
Complex 2
16
20
5
10
15
20
25 Form II
CE P
TE
D
MA
NU
SC R
IP
T
Form I
Fig. 8.
AC
DNA
30
ACCEPTED MANUSCRIPT Complex 1 DNA Topo II 5
10
15
Complex 2 20
25
DNA Topo II
5
10
15
20
25
Form I
TE
D
MA
NU
SC R
IP
T
Form II
AC
CE P
Fig. 9.
31
ACCEPTED MANUSCRIPT Graphical abstract 2+
N
H N
N
N
Ru N
N
O
N
H N
N
N
Ru
N
N
O
O N
N
O
SC R
IP
N
N
T
N N
2+
Complex 1
Complex 2
AC
CE P
TE
D
MA
NU
Results suggest that complexes 1 and 2 can bind to DNA via an intercalative mode and efficiently photocleavage pBR322 DNA in vitro under irradiation. In addition, the results of topoisomerase inhibition and DNA strand passage assay reflect that both complexes are efficient dual dual poisons of topoisomerases I and IIα.
32
ACCEPTED MANUSCRIPT Highlights ► DNA binding, photocleavage and topoisomerase inhibitory activity of complexes 1 and 2 are
T
evaluated. ► Complexes 1 and 2 bind to DNA via intercalation. ► Complex 1 exhibits more
AC
CE P
TE
D
MA
NU
SC R
2 are efficient dual poisons of topoisomerases I and IIα.
IP
effective DNA cleavage activity than complexes 2 under the same conditions. ► Complexes 1 and
33