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(This is a sample cover image for this issue. The actual cover is not yet available at this time.)
This article appeared in a journal published by Elsevier. The attachedcopy is furnished to the author for internal non-commercial researchand education use, including for instruction at the authors institution
and sharing with colleagues.
Other uses, including reproduction and distribution, or selling orlicensing copies, or posting to personal, institutional or third party
websites are prohibited.
In most cases authors are permitted to post their version of thearticle (e.g. in Word or Tex form) to their personal website orinstitutional repository. Authors requiring further information
regarding Elsevier’s archiving and manuscript policies areencouraged to visit:
Copper(II) complexes of terpyridine derivatives: A footstep towards development ofantiproliferative agent for breast cancer
Subramaniyam Rajalakshmi a, Thomas Weyhermüller b, Murugan Dinesh c, Balachandran Unni Nair a,⁎a Chemical Laboratory, Central Leather Research Institute, Council of Scientific and Industrial Research, Adyar, Chennai-600 020, Tamil Nadu, Indiab Max‐Planck Institut für Bioanorganische Chemie, D-45470, Mülheim an der Ruhr, Germanyc Life Teck Research Centre, Vadapalani, Chennai-600 026, Tamil Nadu, India
a b s t r a c ta r t i c l e i n f o
Article history:Received 16 April 2012Received in revised form 14 August 2012Accepted 14 August 2012Available online 30 August 2012
Two copper(II) complexeswith terpyridyl conjugates, [Cu(meotpy)(dmp)](NO3)2 (1) and [Cu(bitpy)(dmp)](NO3)2(2) where meotpy, bitpy and dmp stand for methoxybenzyl terpyridine, benzimidazolyl terpyridine and dimethylphenanthroline respectively have been synthesized and characterized. Complex 1 has also been characterized crys-tallographically. Both the complexes have been found to bind CT-DNA intercalatively. The ability of these complexesto bring about DNA cleavage has been analyzed using gel electrophoresis. Both complexes 1 and 2 have been foundto bring about hydrolytic cleavage of DNA. The cytotoxicity of both these complexes has been tested against cancer-ous as well as non-cancerous cell lines. Towards non-cancerous cell line complex 2 exhibited very low toxicity. Onthe other handboth the complexes have been found to exhibit cytotoxic effects against cancerous cell lines. Complex2which has lower IC50, was found to be a potent antiproliferative agent against MCF-7 cells and was able to inducemitochondrial-mediated and caspase-dependent apoptosis with increase in G0/G1 and subsequent arrest in the Sphase, in cell cycle progression. Based on this study, it is hypothesized that 2may be a suitable candidate for furtherevaluation as a chemopreventive and chemotherapeutic agent for human cancer.
DNA is the primary target molecule for most of the anticancer andantiviral agents [1–3]. The cleavage of nucleic acids is an important en-zymatic reaction involved in several biological processes as well as inbiotechnological manipulation of genetic material. Naturally, nucleasesare the enzymes that rapidly hydrolyze the DNA phosphodiester bondby a factor of 1012 compared to the noncatalyzed reactions [4]. Whilemany reagents have been successfully employed to bring about hydro-lytic cleavage of RNA [5–10], not much information is available on thehydrolytic cleavage of DNA because of its relatively high hydrolytic sta-bility [11–13]. The search for small molecules capable of cleaving DNAbond is still ongoing although a great number of natural nucleases areknown. A large number of oxidative cleavage reagents have been uti-lized with grand success for DNA footprinting [14,15], loop region rec-ognition [16,17], locating conformational variations in DNA [18–21],and as chemotherapeutic agents [22]. Despite high efficiency and versa-tility, the oxidative DNA cleaving agents are dependent on cofactorssuch as light or an oxidizing or a reducing agent to execute their actions[23], and this hampers their use in vivo, especially in the case of thera-peutic applications. Oxidative cleavage processes are radical-based [24]and deliver products lacking 3′ or 5′‐phosphate groups [25] and as a
result are not amenable to further enzymatic manipulation and thusthe use of these reagents has been limited in the field of molecular biol-ogy. Hydrolytic DNA cleaving agents are not associatedwith such draw-backs and hence the development of reagents which hydrolyticallycleave nucleic acids under mild conditions is currently attracting greatinterest in the field of artificial metallonucleases.
Hitherto, numerous transition metal complexes have been found tobe active in terms of restraining the reproduction of cancer cells, andsome of them have been widely used in clinical treatments [26]. Inorder to obtain complexes with such an efficiency, a variety of metalcomplexes, including those of Cu(II), Ni(II), Zn(II), Co(II), Fe(II) and lan-thanide have been synthesized and investigated with respect to theirability to hydrolyze small phosphodiesters [27–29]. Copper(II) beingthe second most bio-essential transition metal ion, and its complexeswith tunable coordination geometries in a redox active environmentcould find better application at the cellular level. The role of Cu(II) atthe cellular level has been provenwith respect to Cu(II)mediated activa-tion of certain critical proteins such as Cu–Zn superoxide dismutase, ty-rosinase, ceruloplasmin, and cytochrome oxidase, which are importantin fundamental biological pathways [30]. Therefore, the copper concen-tration in human body is tightly regulated [31]. However, the concentra-tion of copper in cancerous tissues such as breast, prostate, lung, andbrain, is higher than that of the normal tissues [32–34]. Recently it hasbeen reported that pyridyl-type ligand [35] and copper(II) terpyridylcomplexes show moderate anticancer activities [36,37]. Complexes ofthese ligands would be good candidates for examination of their effect
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⁎ Corresponding author at: Chemical Laboratory, Central Leather Research Institute,Adyar, Chennai-600 020, India. Fax: +91 44 2491 1589.
on cell proliferation. In this paper, an attempt has been made to synthe-size two mixed ligand copper(II) complexes, with terpyridyl derivativespossessing both extended planarity and H-bonding ability, as one of theligands. We have also scrutinized their ability to bind and cleave DNAalong with their effect on a cancerous and a non-cancerous cell line.
2. Experimental
2.1. Materials
2-Acetyl pyridine, 2,9-dimethyl phenanthroline, 2-benzimidazolecarbaldehyde, 4‐methoxybenzaldehyde, calf thymus DNA, agarose,ethidium bromide, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazoli-um bromide, propidium iodide, proteinase K, phenylmethanesulfonylfluoride (PMSF), and RNAse A were purchased from Sigma ChemicalCompany (St. Louis, MO, USA). The Minimum Essential Medium Eagle(MEM) was purchased from Hi Media Laboratories and fetal bovineserum (FBS) was purchased from Cistron Laboratories. Trypsin, DMSO,Tris and Tris–borate–EDTA (TBE) were purchased from Sisco ResearchLaboratory (Mumbai, India). The human breast cancer cell line, MCF-7(GDC055) and the African green monkey kidney fibroblast cell line,VERO were obtained from the National Centre for Cell Sciences (NCCS,Pune, India). Both cells were maintained in MEM supplemented with10% FBS, penicillin (100 U/mL), and streptomycin (100 μg/mL) in a hu-midified atmosphere of 50 μg/mL CO2 at 37 °C. All the reagents used inthe study were of analytical grade. Antibodies to β-actin, PARP,caspase-3 and caspase-9, BCl2, and Bax were purchased from SantaCruzBiotechnology, CA, USA andNeoMarkers, USA.Milli‐Q triply deion-ized water was employed for all the studies.
2.2. Instrumentation
Electronic spectra were recorded using a Perkin-Elmer Lambda 35double beam spectrophotometer. Electrospray ionization mass spec-tra (ESI-MS) were obtained from a Thermo Finnigan LCQ 6000 advan-tage max ion trap mass spectrometer using acetonitrile as the carriersolvent. A stock solution of DNA was prepared by stirring a DNA sam-ple dissolved in 10 mM TrisHCl buffer (pH 7.2) at 4 °C and used with-in 4 days of preparation. The solution was exhaustively dialyzedagainst Tris buffer for 48 h and filtered using a membrane filterobtained from Sartorius (0.45 μM). The filtered DNA solution in thebuffer gave a UV absorbance ratio of (A260/A280) about 1.9, indicatingthat the DNA was sufficiently free from protein [38]. The concentra-tion of DNA was determined using an extinction coefficient of6600 M−1 cm−1 at 260 nm [39]. All the experiments were carriedout employing the prepared DNA solution in Tris buffer at pH 7.2.
2.3. Synthesis of [Cu(meotpy)(dmp)](NO3)2·H2O (1)
The ligands, meotpy and bitpy were synthesized employing theprocedure reported in literature [40]. Briefly, the complex 1 wasprepared by stirring a methanolic solution of Cu(NO3)2·3H2O(0.12 g, 0.5 mmol) with meotpy (0.15 g, 0.5 mmol) at room tem-perature for 30 min. Subsequently, dmp (0.12 g, 0.5 mmol) wasadded to the above solution and stirred continuously for another30 min. A green solid that separated out upon slow evaporation ofthe solvent was filtered, and washed with diethyl ether and driedin vacuum. The complex was recrystallized from an acetonitrile so-lution. Yield: 79%. Found: C, 68.61; H, 5.18; N, 11.18. Anal Calcd forC36H33CuN5O2: C, 68.50; H, 5.27; N, 11.09. ESI-MS m/z: 305. Singlecrystals which were suitable for X-ray diffraction studies wereobtained upon slow evaporation of acetonitrile–water solution ofthe complex.
2.4. Synthesis of [Cu(bitpy)(dmp)](NO3)2·H2O (2)
The complex 2was synthesized by following the above mentionedprocedure and employing bitpy (0.31 g, 1 mmol), dmp (0.21 g,1 mmol), and Cu(NO3)2·3H2O (0.24 g, 1 mmol). Yield: 72%. Found:C, 67.54; H, 4.78; N, 15.25. Anal Calcd: for C36H31CuN7O2: C, 67.43;H, 4.87; N, 15.29. ESI-MS m/z: 310. The dark reddish brown precipi-tate obtained was dissolved in an acetonitrile–water mixture andkept for volatilizing the solvent at room temperature. Attempts tocrystallize complex 2 were unsuccessful.
2.5. X-ray crystallographic data collection and refinement of thestructures
Blue single crystals of [Cu(meotpy)(dmp)](NO3)2·H2O were coatedwith perfluoropolyether and mounted in the nitrogen cold stream(100 K) of the diffractometer equipped with a Mo-target rotating-anode X-ray source (Mo-Kα, λ=0.71073 Å). Final cell constants wereobtained from least squares fits of several thousand strong reflections.The Siemens ShelXTL [41] software package was used for the solutionand artwork of the structure, while ShelXL97 was used for the refine-ment [42]. The structure was readily solved by Patterson method andsubsequent difference Fourier techniques. All non-hydrogen atomswere anisotropically refined. Hydrogen atoms attached to carbonatoms were placed at calculated positions and refined as riding atomswith isotropic displacement parameters. Hydrogen atoms of solventwatermolecules and hydrogen atoms bound to nitrogen atomswere lo-calized from the difference map and were refined with restrained O–Hdistances and displacement parameters. The occupation factors wererefined to values of about 0.49, 0.39, and 0.12 for the split positionscontaining N(500), N(510), and N(520), respectively. Only two posi-tions of the water molecule forming hydrogen bonds to the nitrate ionand other water molecules could be identified. The occupation ratiowas refined to 0.65:0.35 (O(670) and O(675)). The CCDC depositionnumber of the crystal is 824244.
2.6. DNA binding and cleavage experiments
Concentrated stock solutions of metal complexes were preparedby dissolving the complexes in acetonitrile and suitably dilutingwith 10 mM Tris buffer at pH 7.2 (1:10 as acetonitrile:buffer) to therequired concentration for all the experiments. Absorption spectral ti-tration experiments were carried out for both complexes 1 and 2 bymaintaining a constant concentration of the complex (20 μM) andvarying the CT-DNA concentration (5–120 μM). An equal amount ofDNA was added to the cell in the reference compartment.
For viscositymeasurements, theUbberhold viscometer (1 mL capac-ity) was thermostated in a water bath maintained at 25 °C. The flowtime for each sample was measured three times using a digital stop-watch and the average flow time was calculated. The rate of flow forthe buffer (10 mM Tris), DNA (100 μM) and DNAwith the copper com-plexes at various concentrations (5–120 μM) was measured. The rela-tive specific viscosity was calculated using the equation, η=(t−to)/to, where to is the flow time for the buffer and t is the observed flowtime for DNA in the absence and presence of the complex. Data arepresented as (η/ηo)1/3 versus 1/R {R=[complex]/[DNA]}, where η isthe viscosity of DNA in the presence of the complex and ηo is the viscos-ity of DNA alone [43,44]. Thermal denaturation experiments were car-ried out with a Perkin-Elmer Lambda 35 spectrophotometer equippedwith a Peltier temperature-control programmer. The temperature wasgradually increased from 25 to 95 °C at a rate of 5 °C min−1. The absor-bance at 260 nmwas recorded for CT-DNA (100 μM) in the absence andpresence of both copper(II) complexes (2.5 μM) at an interval of 5 °C.The Tm values were determined from the plot of relative absorbanceversus temperature.
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Circular dichroic spectra were recorded with a Jasco J-715spectropolarimeter at 25 °C using a 0.1 cmpath quartz cell and scannedin the spectral region of 220–300 nm. The concentration of CT-DNA(100 μM) was kept constant and the concentration of copper(II) com-plexes 1 and 2 varied from 5 to 120 μM.
Cleavage of DNA by copper(II) complexes was monitored by usingthe agarose gel electrophoresis technique. The DNA cleavage efficien-cy of complexes 1 and 2 was monitored by determining their abilityto convert supercoiled DNA (form I) to open circular (form II) and lin-ear forms (form III). Both complexes with varying concentrationswere incubated for 1 h at 37 °C with pUC 18 plasmid DNA. A loadingbuffer containing 0.25% bromophenol blue, 40% (w/v) sucrose and0.5 M EDTA was added and the electrophoresis of the DNA cleavageproducts was performed on a 0.8% agarose gel containing 0.5 μg/mLethidium bromide. The gels were run at 50 V for 2 h in Tris–boricacid–ethylenediamine tetra acetic acid (TBE) buffer at pH 7.4. Thebands were viewed by placing the gel on a UV illuminator and werephotographed using a gel documentation system. A set of experi-ments was also conducted in the presence of DMSO (10 mM) aswell as sodium azide (20 mM). For relegation experiments, theform II obtained in the presence of the complexes was recoveredfrom agarose gel by phenol extraction method and the recoveredNC DNA was purified by ethanol precipitation. Subsequently the puri-fied NC DNA was incubated with T4 DNA ligase in a 10× ligation buff-er for 16 h at 25 °C and subjected to gel electrophoresis.
2.7. In vitro assay for cytotoxicity activity
The cytotoxic effect of complexes 1 and 2 on MCF-7 cells was de-termined by the MTT assay. Cells (1×105/well) were plated in 100 μLof medium/well in 96-well plates. Usually, after 48 h of incubation,cells reach the state of confluence. In the present experiment, afterthe confluence state was reached, the cells were incubated in thepresence of various concentrations of the samples in Tris buffer for48 h at 37 °C. After removal of the sample solution and washingwith phosphate-buffered saline (pH 7.4), 20 μL/well (5 mg/mL) of0.5% 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-tetrazolium bromidecell (MTT) phosphate-buffered saline solution was added. After 4 hof incubation, 0.04 M DMSO was added. Viable cells were determinedby taking the absorbance at 570 nm. Measurements were performedand the concentration required for a 50% inhibition of viability (IC50)was determined graphically. The effect of the samples on the prolifer-ation of human breast cancer cells was expressed as the % cell viabil-ity, using the following formula
% cell viability ¼ A570 nm of treated cells=A570 nm of control cells� 100% :
2.8. Western blot analysis
Typically, 1×105 exponentially growing MCF-7 cells of the experi-mental groupwere seeded onto 6-well plates for 24 h followed by treat-ment with IC50 concentration of the complex for 48 h. Cells werecollected by trypsinization and washed three times with PBS, lysed incell lysis buffer containing 0.05 M Tris–HCl (pH 7.2), 0.15 M NaCl,0.001 M PMSF, 0.001 M EDTA (pH 8.0), 1% Triton X-100, 0.1% SDS, and2 μg/mL leupeptin, and then centrifuged at 12,000 rpm for 5 min at4 °C. The extracted protein samples (10 μg total protein/lane) wereadded in 5 times the volume of the sample buffer and subjected to dena-turation at 100 °C for 10 min, then electrophorized on SDS-PAGE (8%PARP, 15%, caspases-3, and ‐9, β-actin) at 200 V for 45 min, and finallytransferred to a PVDF membrane. The PVDF membrane was treatedwith PBS containing 5% skimmed milk at room temperature for 1 hfollowed by incubation with the primary antibodies, anti PARP (dilution1:2000), and mouse monoclonal primary antibodies against anti human
caspase 3 (1:2000), anti-human caspase 9 (1:2000), and anti-humanBax(1:2000) at 37 °C for 1 h or at 4 °C overnight. After being washed withPBS 3 times for 15, 10, and 10 min, respectively, the corresponding sec-ondary antibody (dilution 1:1000) was added and incubated at roomtemperature for 1 h. The membrane was then washed thrice for 15, 10,and 10 min respectively. After reaction with horseradish peroxidase-conjugated goat anti-mouse antibody, the immune complexes were vi-sualized by using the chemiluminescence ECL PLUS detection reagentsfollowing the manufacturer's procedure (Amersham Bioscience).
2.9. Cell cycle analysis by flow cytometry
Typically, MCF cells (5×105) were seeded into 6 well plates andpre-cultured for 24 h. The cells were treated with the IC50 concentra-tion of the complex. The cells were trypsinized, washed twice withPBS, and suspended in 500 μL of PBS for 15 min on ice. The cell sus-pension was mixed with 5 mL of cold 70% ethanol and stored at4 °C until analysis. On the day of analysis, cells were washed twicewith PBS buffer and re-suspended in 1 mL PBS. After incubationwith RNase A (250 μg/mL) for 30 min and staining with propidiumiodide (PI, 10 μg/mL) for 10 min, cell cycle analysis was carried outusing the FACScan fluorescence-activated cell sorter(BD Biosciences,San Jose, CA, USA). Histograms were generated and the subsequentanalysis was carried by Cell Quest software.
3. Results
3.1. Synthesis and characterization of the complexes
Complexes [Cu(meotpy)(dmp)](NO3)2 (1) and [Cu(bitpy)(dmp)](NO3)2 (2) (Scheme 1) where meotpy is methoxybenzyl terpyridine,bitpy is benzimidazolyl terpyridine and dmp is dimethyl phenanthroline,have been synthesized from the reaction of meotpy, bitpy, and dmpwithCu(NO3)2·3H2O in methanol solution. The tridentate meotpy/bitpy andbidentate dmp ligands with copper(II) form Cu(II)N5 type complexes.The authenticities of the respective complexes were confirmed by singlecrystal X-ray diffraction study for complex 1 and ESI-MS spectroscopictechnique for both the complexes. ESI-MS spectrum of both complexesshowed signals for their respective molecular ions, at (m/z) 305.20 forcomplex1 and at (m/z) 311.87 for complex2 (Fig. 1a andb). The electron-ic absorption spectra of complexes 1 and 2 are shown in Fig. 2. The UV–
Scheme 1. Representation of complex 1 and complex 2.
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visible spectrum of complex 1 shows a strong CT band at 343 nm andintraligand transition at 277 nm and 218 nm. The corresponding transi-tions for complex 2 have been observed at 350, 274, and 212 nm respec-tively. Both the complexes also exhibit low intensity ligandfield transitionaround 550–650 nm.
3.2. Crystal structure
Complex 1 has been structurally characterized by single crystalX-ray diffraction. The crystallographic data for complex 1 is given inTable 1. The crystal structure shows a mononuclear dicationic natureof the complex having the chelating bidentate phenanthroline andthe tridentate terpyridine derivative. The ORTEP representation ofthe structure of complex 1 including the atom numbering scheme isshown in Fig. 3. Selected bond lengths and bond angles of complex
1 are listed in Table 2. The basal plane of complex 1 is formed byN(31), N(42) of the bidentate (dmp) ligand and N(12) of thetridentate (meotpy) ligand. The apical position is occupied by N(1)and N(18) of the tridentate ligand.
3.3. DNA binding studies
3.3.1. Absorption spectral studiesAssessment of the DNA binding ability of metal complexes is very
important in the development of effective metal based chemothera-peutic drugs. Monitoring the effect of DNA on the absorption spec-trum of a metal complex is one of the most widely used methods todetermine the binding constant of a complex with DNA. Binding ofa metal complex to DNA through intercalation usually results inhypochroism and red shift (bathochromism) in the absorption
Fig. 1. (a) Electrospray mass spectrum of complex 1 in an acetonitrile–water solution (1:2). (b) Electrospray mass spectrum of complex 2 in an acetonitrile–water solution (1:2).
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spectral bands of the complex due to strong stacking interaction be-tween the aromatic chromophore and the base pairs of DNA [45].
The absorption spectrum of complexes 1 and 2 in the absence andpresence of CT-DNA is shown in Fig. 4a and b respectively. A decreasein the intensity of the spectral bands of complex 1 (at 353 nm) wasobserved with an increase in the concentration of DNA (5–120 μM).A similar change was observed in the electronic spectrum of complex2 (at 354 nm) upon incremental addition of DNA. However, in thiscase no significant change was observed in the spectrum of complex2 beyond the 25 μM DNA. In order to quantitatively compare thebinding affinity of complexes 1 and 2 to CT-DNA, the intrinsic bindingconstants (Kb) of the complexes were determined by monitoring thechanges in absorbance of the intraligand bands of both complexeswith increasing concentration of DNA [46]. The Kb has been calculatedfrom Eq. (1)
where εa, εf, and εb correspond to Aobsd/[Cu], the extinction coefficientfor free copper(II) complex and the extinction coefficient for the
copper(II) complex in the fully bound form, respectively [47]. The Kb
of complexes 1 and 2 were determined to be (7.1±0.25)×104 and(1.8±0.31)×105 M−1, respectively.
3.3.2. Thermal denaturation studiesThermal denaturation studies can conveniently be used in
predicting the nature of binding of the complexes to DNA and theirrelative binding strength. Fig. 5 shows the melting curves of CT-DNAin the absence and presence of complexes 1 and 2. Tm, the tempera-ture at which 50% of the double stranded DNAmolecules have under-gone strand scission has been calculated. Incubation of these twocomplexes with DNA resulted in an increase in Tm of DNA by 7 and4 °C for complexes 1 and 2, respectively.
3.3.3. Viscosity measurementsTo further substantiate the binding nature of the complexes with
DNA, viscosity of DNA was measured in the presence of complexes1 and 2 as shown in Fig. 6a and b. The relative specific viscosity hasbeen calculated using the equation (t− to) /to, where to is the flowtime for the buffer and t is the observed flow time for DNA in the ab-sence and presence of the complex. Data are presented as (η/ηo)1/3 vs1/R {R=[complex]/[DNA]}, where η is the viscosity of DNA in thepresence of a complex and ηo is the viscosity of DNA alone. The valuesof relative specific viscosities were plotted against 1/R for both com-plexes. An exponential increase in the viscosity of DNA was observedwith an increase in the concentration of complex 1 (5–120 μM) andattained saturation around 100 μM of complex 1. Similarly, in thepresence of complex 2 an increase in the viscosity of DNA was ob-served. However, in this case saturation was observed around35 μM of complex 2.
3.3.4. Circular dichroic spectral studiesCircular dichroic (CD) spectroscopic studies were carried out to
gain an insight into the interaction of complexes 1 and 2 with DNA.The circular dichroic spectra of CT DNA were recorded in the wave-length range of 220–300 nm with increasing concentration of com-plexes 1 and 2 (5–120 μM). The spectra of CT DNA were typical ofthe B-form DNA, with a positive Cotton effect near 277 nm due tobase stacking and a negative Cotton effect near 245 nm due to poly-nucleotide helicity of B-DNA [48]. The CD spectrum of DNA in thepresence of complex 1 is shown in Fig. 7a. Complex 1 led to a gradualincrease in the intensity of the positive band of the DNA with a smallblue shift in the energy of the band. There were no appreciablechanges in the negative band. On the other hand, the CD spectrumof the DNA in the presence of complex 2 (Fig. 7b) was moreperturbed. In the presence of complex 2 there was an appreciable in-crease in the intensity of the positive band with a blue shift by almost7 nm in the wavelength. The magnitude of the negative band becameless negative with incremental addition of complex 2. Surprisingly, inthis case at higher concentrations of complex 2, the CD spectrum ofthe DNA showed an additional negative band around 285 nm.
4. DNA cleavage properties
The DNA strand scission chemistry of the synthesized complexeshas been investigated by quantification of the supercoiled (SC)form. Generally, when the original SC form (form I) of plasmid DNAis nicked, an open circular relaxed form (form II) exists in the systemand the linear form (form III) can be found upon further cleavage. Thedistribution of SC, NC (nicked circular) and linear forms of DNA in theagarose gel electrophoresis provides a measure of the extent of cleav-age of DNA [49]. The results of the gel electrophoresis experimentsperformed on DNA incubated with complexes 1 and 2 are shown inFig. 8a and b, respectively. The control experiments performed inthe absence of complexes 1 and 2 (lane 1 of Fig. 8a and b) did notshow any DNA cleavage. Complexes 1 and 2 induced conversion of
Fig. 2. UV–visible spectrum of complex 1 and complex 2 in an acetonitrile–water solu-tion (1:2).
Table 1Crystal data and structure refinements of complex 1.
Parameters Complex 1
Chem formula C36H31CuN7O8
Fw 753.22Space group P212121, no. 19a, Å 15.896(3)b, Å 18.028(3)c, Å 22.749(2)β, deg 90V, Å 6519.3(17)Z 8T, K 100(2)ρ calcd, g cm−3 1.535Refl. collected/2Θmax 97,982/65.00Unique refl./I>2σ(I) 23,542/19,052No. of params/restr. 955/5μ, cm−1/λ, Å 7.38/0.71073R1a/goodness of fit b 0.0428/1.076wR2c (I>2σ(I)) 0.0812Residual density, eÅ−3 +0.49/−0.59
a Observation criterion: I>2σ(I). R1=Σ||Fo|− |Fc|| /Σ|Fo|.b GooF=[Σ[w(Fo2−Fc2)2]/(n−p)]1/2.c wR2=[Σ[w(Fo2−Fc2)2] /Σ[w(Fo2)2]]1/2 where w=1/σ2(Fo2)+(aP)2+bP, P=
(Fo2+2Fc2) /3.
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DNA from the SC form to the NC form upon 1 h of incubation at 37 °C.Since no other coreagent was added to the reaction mixture, it is ev-ident that the copper(II) complexes are able to hydrolytically cleaveDNA. Ligase experiment was simultaneously carried out to ascertainthe mechanism of the DNA cleavage reaction of the two complexes[50]. Fig. 8c shows the relegation of nicked circular (obtained in thepresence of complex 1) to supercoiled form. Fig. 8c also shows the re-sults of DNA cleaving experiments performed in the presence of thehydroxyl radical quencher DMSO and the singlet oxygen quenchersodium azide.
5. Cytotoxic effects on MCF-7 and VERO cell lines
The cytotoxicity of the two copper(II) complexes were studiedagainst a human cancerous breast adenocarcinoma cell line (MCF-7)and a noncancerous monkey kidney fibroblast cell line (VERO) usingthe MTT assay. The MTT assay serves as an index of cell viability bydetecting the reduction of tetrazolium salt to blue formazan by themitochondrial enzymatic activity of succinate dehydrogenase inliving cells [51]. In the present investigation, a significant variation
was observed on the survival of two specific cell lines in the presenceof complexes 1 and 2. Fig. 9 represents the comparative study of com-plexes 1 and 2 with the two different cell lines. Both the complexesexhibited a significant inhibitory potency towards proliferation ofthe MCF-7 cell line as can be seen from the figure. Half-maximal in-hibitory concentration (IC50) values were determined to be 6.25 and3.125 μM for complexes 1 and 2, respectively. Both the complexeswhen tested against the VERO cell line exhibited a cell viability of92.34 and 95.67%, respectively at a low concentration of 0.78 μM.Whereas, in the case of the MCF-7 cell line, the % cell viability was ob-served to be 87.77 for complex 1 and 76.89% for complex 2 at a similarconcentration. At the higher concentration of 100 μM, the cell viabil-ity for the MCF-7 cell line was 16.77 and 5.67% for the respective com-plexes 1 and 2. In the VERO cell line, at the higher concentration of100 μM the cell viability was found to be 32.34% for complex 1 and43.33% for complex 2. It is also evident from Fig. 9 that complex 2 iscomparably less toxic towards the VERO cell line than complex 1.Since among the two complexes studied, complex 2 showed betteractivities in terms of inhibiting cell proliferation of a cancer cell line(MCF-7) and non-toxic effect against a noncancerous (VERO) cellline, further investigations were carried out only with complex 2.DNA fragmentation assay was performed to get an insight to themechanism of cell death brought about by complex 2. The results(Fig. 10) show fragmented DNA, indicating that complex 2 inducesapoptosis.
5.1. Western blot analysis
To investigate further the mechanistic pathway of cell death bycomplex 2 on MCF‐7 cell lines, a study on the protein levels of a fewapoptotic markers such as Caspase-3, Bax, Bcl-2, caspases-9 and p53in the presence of complex 2 was performed by employing Westernblot analysis (Fig. 11). A reduction in the expression of Bcl-2 was ob-served when cancerous cells were exposed to complex 2 with in-creasing concentrations (1.56, 3.15, and 6.25 μM). On the otherhand, an increased expression of Bax, caspase-3, and p53 was ob-served upon exposure of cancerous cells to the similar dosage of com-plex 2. The level of cleaved PARP product (85 kD) was higher in thetreated cells than in the untreated cells. The expression of β-actin,which served as a loading control remained unaltered.
Table 2Selected bond length and bond angles of complex 1.
Fig. 3. ORTEP representation of complex 1 together with the atom numbering scheme.
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5.2. Cell cycle study
Further, to scrutinize the mechanistic pathway and the growth in-hibitory effect of complex 2 on the MCF-7 cell line, the fluorescence-activated cell sorting (FACS) analysis employing propidium iodide (PI)staining was performed. To investigate the molecular mechanism un-derlying the inhibitory effect of complex 2 on the cell growth of a can-cerous cell line, the different phases of the cell cycle were examinedby flow cytometry, which involved quantitative measurement of theDNA content of cells at each phase of the cell cycle. The MCF-7 cellswere treated with a 3.125 μM solution (the IC50 value) of complex 2,as well as with 1.56 μM (lower than the IC50 value of the complex)and 6.25 μM (higher than the IC50 value) of complex 2 and subsequent-ly harvested after 24 h of incubation. The PI-stained cells were assessedby flow cytometry with the FL2-A channel. In vitro DNA-flowcytometric analysis was carried out to determine whether the complex2 induced cell growth inhibition was the result of apoptosis or cell cyclearrest or a combination of both. The cell cycle progression in thepresence of complex 2 is presented in Fig. 11. Control experimentsshowed the distribution of cells as 45.32% in the G0/G1 phase, 38.90%in the G2/M phase and 17.61% in the S phase. Cells on treatment with
Fig. 4. (a) Absorption spectral titration of complex 1 (10 μM) in a Tris buffer solutionwith increasing concentration of CT-DNA (5–120 μM). (b) Absorption spectral titrationof complex 2 (10 μM) in a Tris buffer solution with increasing concentration of CT-DNA(5–100 μM).
Fig. 5. Melting curves of CT-DNA (100 μM) upon addition of complexes 1 (20 μM) and2 (20 μM).
Fig. 6. (a) Effect of complex 1 (5–120 μM) on the relative viscosity of CT-DNA in Tris buff-er. (b) Effect of complex 2 (5–100 μM) on the relative viscosity of CT-DNA in Tris buffer.
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1.56, 3.12 and 6.25 μM of complex 2 for 24 h resulted in an increase inthe proportion of cells in the G0/G1 phase by 52.45, 68.91, and 81.26%respectively. The proportion of cells in the G2/M and S phases on theother hand decreased in the presence of various concentrations of com-plex 2.
6. Discussion
Two copper(II) mixed ligand complexes with terpyridyl conjugateswere synthesized and characterized using ESI-MS and UV–visible
spectroscopic techniques. The ESI being a soft ionization technique inmass spectrometry has become a very useful tool in confirming the mo-lecular identity of a compound. ESI mass spectral measurements of theacetonitrile–water solution containing complex 1 showed a peak atm/z305.20, which could be attributed to the complex cation [Cu(meotpy)(dmp)]2+. Similarly, the ESI-MS spectrumof the acetonitrile–water solu-tion of complex 2 showed a peak atm/z 311.87 attributable to the com-plex cation [Cu(bitpy)(dmp)]2+. Hence, the ESI-MS clearly establishesthe authenticity of the two synthesized complexes. The mass spectrumof the two complexes also establishes that the complex cation remainsintact, without any ligand loss in the solution. Complex 1was also char-acterized using single crystal X-ray diffraction study which showed thegeometry of the complex to be distorted trigonal bipyramidal. The crystalstructure of complex 1, shown in Fig. 3 revealed (3+2), the five coordi-nate geometry of the complex. The two Cu\N bond distances,Cu\N(31) and Cu\N(42) in the basal plane, arising from the coordina-tion of copper(II) ion to the bipyridyl ligand,were found to be 2.2216 and1.9942 Å. The third Cu\N(12) bond distance in the basal plane, arisingfrom the coordination of the terpyridyl ligand to the copper(II) ion was1.9352 Å. The apical Cu\N(1) and Cu\N(18) bond lengths (2.0357–2.0483 Å)were almost the same, and comparablewith otherfive coordi-nate complexes [36]. Since both complexes 1 and 2 possess similar li-gands coordinated to the Cu(II) ion, both these complexes showedsimilar electronic spectrum in the spectral region 200–450 nm. Basically,this specific region of the spectrum is dominated by the intraligand tran-sition and CT transition. Complex 1 showed a low intensity broad bandcentered at 628 nm due to ligand field transition of the complex. Thesame transition was observed at 623 nm for complex 2.
Fig. 7. (a) CD spectra of CT-DNA in the presence of 5–120 μM of complex 1 in Tris buff-er. (b) CD spectra of CT-DNA in the presence of 5–100 μM of complex 2 in Tris buffer.
Fig. 8. (a) Cleavage of SC pUC 18 DNA upon addition of complex 1. DNA (200 ng) was incubated with the complex for 60 min in Tris buffer (pH 7.2). Lane 1, DNA control; lane 2,DNA+1 (40 μM); lane 3, DNA+1 (60 μM), lane 4, DNA+1 (80 μM). (b) Cleavage of SC pUC 18 DNA upon addition of complex 2. DNA (400 ng) was incubated with the complex for60 min in Tris buffer (pH 7.2). Lane 1, DNA control; lane 2, DNA+2 (40 μM); lane 3, DNA+2 (60 μM), lane 4, DNA+2 (80 μM). (c) Analysis of the capacity of T4 DNA ligase torelegate DNA cleaved by complex 1. Lane 1, DNA+1 (40 μM); lane 2, NC obtained using 1+T4 ligase; lane 3, DNA+1 (40 μM)+DMSO (10 mM); lane 4, DNA+1(40 μM)+NaN3 (20 mM).
Fig. 9. Effects of complexes 1 and 2 on the survival of MCF-7 (dotted line) and VEROcells (solid line).
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Interaction of the synthesized complexes with CT‐DNA was car-ried out to understand the DNA binding mode of the two complexes.Electronic spectra of the two complexes were monitored with incre-mental amount of DNA (5–120 μM) (Fig. 4a and b). The intensity ofboth spectral bands in the UV region of the two complexes steadilydecreased with incremental addition of DNA, subsequently reachinga saturation level. The saturation level in the spectral change of com-plex 2 in the presence of DNA was observed at a much lower concen-tration of DNA (5–35 μM), than that which was observed in the caseof complex 1. This clearly indicates that complex 2 has higher affinityfor the DNA. Clear isosbestic points, at 365 nm for complex 1 and at360 nm for complex 2, were observed in the spectral changes duringthe absorption titration. This observation clearly illustrates that onlytwo species, free complex and DNA bound complex exist in the solu-tion. The spectral changes observed in the two cases are clearly indic-ative of intercalative binding of the two complexes to DNA. Since boththe complexes have a planar aromatic ring (terpyridyl derivatives),which can stack between the DNA base pairs, such an intercalativebinding of two complexes to DNA is not surprising. In the presenceof an incremental amount of DNA the apparent binding constant, Kb
was calculated as (7.1±0.25)×104 and (1.8±0.31)×105 M−1 forcomplexes 1 and 2, respectively. The benzimidazolyl head group ofthe terpyridine ligand of complex 2 possesses planar aromatic struc-tures as well as the N and NH moieties. Hence, it is possible thatthis head (benzimidazolyl) group in complex 2 could possibly be in-volved in hydrogen bonding with the DNA base pair. To study this fur-ther, the melting temperature of DNA in the presence of the twocomplexes was examined (Fig. 5). In the presence of complex 1,DNA showed a 7 °C increase in its melting temperature, whereas in
the presence of complex 2, only an increase of 4 °C was observed.The lower ΔTm for DNA observed in the presence of complex 2 clearlyindicates involvement of the benzimidazolyl head group of complex 2with DNA, where the possibility of H-bonding of N and NH of the li-gand to DNA base pairs exists. As a result, the base pairs might be dis-turbed and so the stabilization of the DNA helix in this case will belesser than in the case where no such hydrogen bonding is involved.In order to confirm further the intercalative binding of the two com-plexes to DNA, the viscosity of DNA was measured in the presence ofvarious concentrations of the complexes (5–120 μM). As can be seenin Fig. 6a and b, the viscosity of DNA increased gradually with the ad-dition of complexes 1 and 2, and reached a saturation point. In thepresence of complex 2 the saturation level was observed at a muchlower concentration of the complex, than that observed in thepresence of complex 1. The observed increase in the viscosity ofDNA in the presence of complexes 1 and 2 confirms intercalativebinding of the two complexes to DNA. In the case of complex 2, theviscosity of DNA even decreased marginally after attaining the satura-tion level. The fact that at higher concentrations of complex 2, therewas even a marginal decrease in the viscosity of DNA indicates thatin this case, intercalation of the ligand is accompanied by partial un-winding of the DNA helix. This destabilization could occur due tobreakage of hydrogen bonds among the base pairs. The circular di-chroism spectrum of DNA in the presence of two Cu(II) complexeswas investigated to understand if binding of these complexes leadsto any change in the conformation of DNA (Fig. 7a and b). The CDspectrum of B-DNA exhibits two conservative bands, a positive banddue to base stacking at 273 nm and a negative band at 242 nm dueto DNA helicity. As can be seen from Fig. 7a, the intensity of the pos-itive band was found to increase with the increase in the concentra-tion of complex 1. The energy of the band was also found toincrease by 2–3 nm in the presence of complex 1. On the otherhand, not much change was observed in the energy and magnitudeof the negative band of DNA. The observed changes in the CD spec-trum of DNA in the presence of complex 1 support the intercalativebinding of the complex to DNA. Simple groove binding and
Fig. 10. DNA band analysis by UV gel doc. Complex 2 treatment induced DNA fragmen-tation in the MCF-7 cell line.
Fig. 11. Western blot analysis of the levels of the Bcl-2 protein family, p53, Bcl-2, Bax,caspase3, and caspase 9 observed with complex 2. Control consisted of samples ofuntreated cells. Actin was used as an internal loading control.
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electrostatic interaction of the complexes with DNA generally lead tono perturbations on the base stacking and helicity bands. In thepresence of complex 2 (Fig. 7b), there was an appreciable increasein the intensity of the positive band of DNA with a blue shift in theband maxima by almost 7 nm. The intensity of the negative band atthe short wavelength decreased with increase in the concentrationof complex 2. The decrease in the intensity of the negative bandmay be due to partial unwinding of the DNA helix upon interactionof complex 2, because of the H-bonding of the N and NH of thebenzimidazolyl group of the terpyridyl ligand with DNA bases. Athigher concentrations of complex 2, a new negative band appearedin the CD spectrum of DNA. It is hypothesized that this negativeband could be due to the induced CD effect of complex 2. These re-sults are in agreement with absorption titration, DNA melting andviscosity experiments. The ability of complexes 1 and 2 to induceDNA cleavage was monitored using gel electrophoretic mobilityassay. Both the complexes were able to convert the supercoiledform to the nicked circular form of DNA without any additives. Evenin the presence of DMSO and sodium azide, which are quenchers ofhydroxyl radicals and singlet oxygen respectively, complexes 1 and2 were able to convert the supercoiled form of DNA to the nicked cir-cular form of DNA. This observation clearly indicates that both thesecomplexes are capable of promoting hydrolytic cleavage of DNA.The ability of the two complexes to bring about hydrolytic cleavageof DNA has been conclusively proved through T4 DNA ligase experi-ment, the results of which are shown in Fig. 8c. Complex 1 wasfound to be more efficient in bringing about hydrolytic cleavage of
DNA compared to complex 2. Complex 1 was able to convert almost90% of supercoiled DNA to the nicked circular form of DNA. On theother hand under identical conditions complex 2 brought aboutonly 10% conversion.
The cytotoxic effects of complexes 1 and 2 were tested against twodifferent cell lines, human breast cancer cell line (MCF-7) and Africangreen monkey kidney cell line (VERO). As can be seen from Fig. 9 boththe complexes exhibited cytotoxic effects on the cancerous MCF-7 cellline. Complex 2 exhibited an IC50 of 3.125 μM whereas for complex 1the IC50 was observed to be 6.25 μM. The effect of complexes 1 and 2on the cell viability of the non-cancerous VERO cell line was also investi-gated. The % cell viability of the cells was found to be 92.34 and 95.67%,respectively at 0.78 μM of complexes 1 and 2. Under similar experimen-tal conditions, the viability of the MCF-7 cell line in the presence of com-plexes 1 and 2was found to be 87.77 and 76.89%, respectively indicatingthat complexes 1 and 2were relativelymore cytotoxic against theMCF-7cell line than against the VERO cell line. Among the two complexes, com-plex 2 exhibited a much lower IC50 than complex 1 when studied withthe MCF-7 cell lines and also demonstrated greater cell viability withthe VERO cell line. Therefore, the results suggest that complex 2 is rela-tively amore potent inhibitor for cancer cell line proliferationwhen com-pared to complex 1. Hence, the subsequent cellular studies were carriedout only with complex 2. DNA fragmentation assay was performed withcomplex 2 to confirm the cell death pathway. During apoptosis, endonu-clease is known to cleave the nucleosomes,which leads to the generationof DNA fragments. In control (MCF-7) cells, genomic DNA showedsmear; however, cells treated with complex 2 showed fragmented
Fig. 12. Effect of complex 2 on the distribution of MCF-7 cells in the cell cycle progression. The cell cycle phase was determined by the cellular DNA content measured by flow cy-tometry of the PI‐stained cells. G0/G1, G2/M and S denote the corresponding phases of the cell cycle. Cell cycle distribution is shown as a histogram. (a) Below IC50 (1.56 μM). (b) IC50
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DNA, indicating that complex 2 induced cell death via apoptosis (Fig. 10).Western blot analysis of complex 2 (Fig. 11) was performed to identifythe expression level of various cellular proteins. Caspase activation isconsidered to be the hallmark of apoptosis, and it is mediated by theBcl-2 family of proteins which includes both the anti-apoptotic member,Bcl-2 and the pro-apoptotic member, Bax [52]. Basically, Bcl-2 familyproteins have been described as key regulators for mitochondrial medi-ated apoptosis [53,54]. The Bcl-2 family of proteins regulates apoptosisby controllingmitochondrial permeability and the release of cytochromec. Conversely, the anti-apoptosis protein Bcl-2 resides in the outer mito-chondrial wall and inhibits cytochrome c release. A reduction in the ex-pression of Bcl-2 protein and an increase in the expression of Bax wereobserved when the cancerous cells were exposed to increasing concen-trations (1.56, 3.15, and 6.25 μM) of complex 2. These results supportcomplex 2 induced apoptosis associated with a reduced Bcl-2 proteinlevel and with the proapoptotic protein, upregulation of Bax was evi-dence. The relation between the tumor suppressor protein p53 and apo-ptosis comes from the ability of p53 to control the transcription of thepro-apoptotic members of the Bcl-2 family. The p53 protein expressionwas concentration-dependent and it was significantly enhancedwith in-creasing concentration of complex 2. The expression of caspase-3 and ‐9levels was activated when treated with complex 2. Activation ofcaspase-3 leads to the cleavage of a number of proteins, one of whichis poly-(ADP-ribose) polymerase (PARP). PARP (112 kDa) is cleaved tothe 85 kDa fragment during apoptosis. Also, the level of cleaved PARPproduct (85 kD) was higher in the treated cells than in the untreatedcells [55]. It shows that the main mechanism of apoptosis appears to bea mitochondrial mediated pathway accompanied by cytochrome c re-lease from the mitochondria into the cytosol, resulting in activation ofcaspase cascades which leads to apoptosis [56]. Furthermore, these re-sults also demonstrate that complex 2 caused a dose-dependent increasein the protein levels of p53, and cleaved caspase-3, cleavage ofpoly(ADP-ribose)polymerase (cPARP), and a reduction in theBcl-2 levels(Fig. 11). Taken together, the results provide evidence that if apoptosisneeds to be caspase-dependant therewould be a corresponding increasein cleavage of PARP, a known consequence of caspase-3 activation. Thiswas subsequently confirmed by Western blotting analysis and showedthat complex 2 induces apoptosis in MCF‐7 cells by the intrinsic mito-chondrial pathway. It seemed useful to further investigate as to whichphase of the cell cycle was directly implicated in the biological behaviorof complex 2. FACS analysis (Fig. 12) was performed to check the regula-tion of cell cycle profile and to understand themechanismof cell divisionand cell death induced by complex 2 onMCF-7 cells. Increase in the per-centage of cells in the G0/G1 phase, accompanied by a reduction in thecell population in the G2/M phase was observed. It was noticed that, inS phase the DNA population decreased (less than ~10%) in comparisonto untreated cells. Based on these observations, it is appropriate tostate that complex 2 inhibits the proliferation of cultured human breastcancer MCF-7 cells and the growth inhibition is associated with the cellcycle arrest and induction of apoptosis. Generally, the S phase is associat-ed with DNA synthesis and plays a crucial role in cell cycle progression,and a series of copper complexes such as CuHQDMTS and CuHQTShave been found to induce S phase arrest and apoptosis of cancer cells[57]. Earlier reports have indicated that cisplatin and other platinumagents inhibit cell cycle progression at the G2/M and S phases [58,59].Complex 2 inhibited MCF-7 cell proliferations as a result of the accumu-lation of cells in the G0/G1 and subsequent reduction in the G2/M and Sphases of the cell cycle, which suggests that complex 2 could inhibitDNA synthesis of MCF‐7 cells.
7. Conclusion
Two copper(II) complexes with terpyridyl derivatives, [Cu(meotpy)(dmp)](NO3)2 (1) and [Cu(bitpy)(dmp)](NO3)2 (2) have been synthe-sized and characterized. Both the complexes were found to exhibit anintercalative mode of binding with CT-DNA. Complex 2 exhibited
hydrogen bonding with DNA bases due to the presence of the N andNH groups on the substituent of terpyridine. Complexes 1 and 2 werealso demonstrated to bring about hydrolytic cleavage of DNA andexhibited modest to high cytotoxicity levels against a cancerous cellline. Complex 2 which exhibited higher DNA binding affinity whencompared to complex 1, showed higher cytotoxicity towards a cancer-ous cell line than complex 1. Complex 2, whichwas found to be a potentantiproliferative agent against a cancer cell line, was also able to inducemitochondrial-mediated and caspase-dependent apoptosis in humancancer cells with increase in G0/G1 and arrest in the S phase, in cellcycle progression. On the basis of the present investigation, we proposethat [Cu(bitpy)(dmp)]2+ may be a better candidate for further evalua-tion as a chemopreventive and chemotherapeutic agent for humancancer.
Abbreviations
meotpy methoxybenzylterpyridinebitpy benzimidazolylterpyridinedmp 2,9-dimethyl phenanthrolineFBS fetal bovine serumMEM minimum essential mediumPMSF phenylmethanesulfonyl fluoridePVDF polyvinylidene difluoridePARP poly-(ADP-ribose) polymeraseMCF human breast adenocarcinoma cell linePBS phosphate buffered salineCT-DNA calf-thymus DNA
Acknowledgment
The authors thankYamini Asthana for valuable discussion.Oneof theauthors (SR) wishes to thank CSIR for the Senior Research Fellowship.
Appendix A. Supplementary data
Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.jinorgbio.2012.08.010.
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