Accepted Manuscript Enantiomeric fluoro–substituted benzothiazole Schiff base–valine Cu(II)/Zn(II) complexes as chemotherapeutic agents: DNA binding profile, cleavage activity, MTT assay and cell imaging studies Rahman Alizadeh, Imtiyaz Yousuf, Mohd Afzal, Saurabh Srivastav, Saripella Srikrishna, Farukh Arjmand PII: DOI: Reference:

S1011-1344(14)00392-3 http://dx.doi.org/10.1016/j.jphotobiol.2014.12.027 JPB 9917

To appear in:

Journal of Photochemistry and Photobiology B: Biology

Received Date: Revised Date: Accepted Date:

22 October 2014 24 December 2014 27 December 2014

Please cite this article as: R. Alizadeh, I. Yousuf, M. Afzal, S. Srivastav, S. Srikrishna, F. Arjmand, Enantiomeric fluoro–substituted benzothiazole Schiff base–valine Cu(II)/Zn(II) complexes as chemotherapeutic agents: DNA binding profile, cleavage activity, MTT assay and cell imaging studies, Journal of Photochemistry and Photobiology B: Biology (2015), doi: http://dx.doi.org/10.1016/j.jphotobiol.2014.12.027

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Enantiomeric fluoro–substituted benzothiazole Schiff base–valine Cu(II)/Zn(II) complexes as chemotherapeutic agents: DNA binding profile, cleavage activity, MTT assay and cell imaging studies Rahman Alizadeha, Imtiyaz Yousufa, Mohd Afzala, Saurabh Srivastavb, Saripella Srikrishnab, Farukh Arjmanda* a

Department of Chemistry, Aligarh Muslim University, Aligarh–202002, India. Department of Biochemistry, Faculty of Science, Banaras Hindu University, Varanasi 221005, India * Corresponding author: Tel.: +91 9897157511 E–mail address: [email protected] (Farukh Arjmand) b

ABSTRACT To evaluate the biological preference of chiral drugs towards DNA target, new metal– based chemotherapeutic agents of Cu(II) and Zn(II), L–/D–fluorobenzothiazole Schiff base–valine complexes 1 & 2 (a and b), respectively were synthesized and thoroughly characterized. Preliminary in vitro DNA binding studies of ligand L and complexes 1 & 2 (a and b) were carried out in Tris−HCl buffer at pH 7.2 to demonstrate the chiral preference of L–enantiomeric complexes over the D–analogues. The extent of DNA binding propensity was ascertained quantitatively by Kb, K and Ksv values which revealed greater binding propensity by L–enantiomeric Cu(II) complex 1a and its potency to act as a chemotherapeutic agent. The cleavage studies with pBR322 plasmid DNA revealed higher nuclease activity of 1a as compared to 2a via hydrolytic cleavage mechanism. The complexes 1 & 2 (a and b) were also screened for antimicrobial activity which demonstrated significantly good activity for L–enantiomeric complexes. Furthermore, cytotoxicity of the complexes 1a and 1b was evaluated by the MTT assay on human HeLa cancer cell line which implicated that more than 50 % cells were viable at 15 µM.

1

These results were further validated by cell imaging studies which demonstrated the nuclear blebbing. Keywords: L–/D–fluorobenzothiazole Schiff base; enantiomeric Cu(II)/Zn(II) complexes; in vitro DNA binding; pBR322 plasmid DNA cleavage activity; MTT assay. Abbreviations CT DNA

Calf thymus DNA

Tris

Tris(hydroxymethyl)aminomethane

EB

Ethidium bromide

MTT

3–(4,5–dimethylthiazol–2–yl)–2,5–diphenyltetrazolium bromide

1. Introduction Benzothiazole– a bicyclic heterocyclic compound is a unique synthon in synthetic medicinal chemistry owing to its significant pharmacological activity [1], it constitutes bioactive pharmacophore of many drugs and therefore has been extensively studied for biological activities viz., antimicrobial, anti–inflammatory, anti–HIV activity, analgesic, anticancer and as lipid peroxidation inhibitors [2–6]. These compounds serve as unique versatile scaffolds for the development of newer drugs which may exhibit different mechanism of action by targeting many potential active sites at the molecular level [7]. Fluorobenzothiazole derivatives possess exquisite potency and selectivity in cytotoxicity with broad spectrum of action than their parent nonfluorinated counterparts and have been therefore extensively explored for chemotherapeutic applications in drug design. For example, fluorinated 2–(4–aminophenyl) benzothiazole exhibited potent cytotoxicity (GI50 < 1 nM) in vitro in sensitive human breast MCF7 and MDA cell lines, but was inactive (GI50 > 10 µM) against PC 3 prostrate and HCT 116 colon cancer cell lines [8,9]. Because of its broad spectrum of action in vitro fluorobenzothiazole derivatives have

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emerged as most potent of new generation of antitumor benzothiazoles and are favored drug candidates for clinical trials [10–12]. The ability of many heterocycles to produce stable complexes with metal ions has great biochemical significance in metal–based drug design [13]. Despite the phenomenal success of cisplatin, cis–diamminedichloroplatinum(II) in the treatment of solid malignancies, the challenges and shortcoming of cisplatin viz., systemic toxicity and drug resistance still remains an issue [14,15]. Therefore, researchers are opting for new elegant modalities in metal–based drug design which involves the drug combination strategy viz., appending a bioactive drug pharmacophore (as a recognition element) to a suitable anchoring ligand framework such as peptide or L–amino acid moiety etc. and incorporating endogenously biocompatible metal ion in its metal binding domain. In this work, we have chosen a Schiff base derivative of fluoro−substituted benzothiazole component in synergy with Cu(II) and Zn(II) metal ions with L–/D–valine amino acid appendage to yield 1 & 2 (a and b) potential antitumor chemotherapeutics. The endogenous metal ions, Cu(II) and Zn(II) not only tune but synchronize the organic ligand scaffold to act in concord at the target site and simultaneously control the lipophilicity of the free drug. This strategy was chosen in order to have a relatively lower dosage of drug that could exhibit less systemic toxicity than the prototypical anticancer chemotherapeutic drugs. The interaction studies of synthesized complexes with CT– DNA, employing biophysical experiments were evaluated to demonstrate their enantiomeric disposition towards the molecular drug target DNA. All the synthesized complexes were further screened for their in vitro antibacterial activity (B. subtilis, S. aureus, E. coli, and P. aeruginosa) and antifungal activities (C. albicans) which revealed

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varying degree of inhibitory effects on the growth of bacterial and fungal strains. In addition, cell proliferation studies of the complexes 1a and 1b were evaluated using MTT assay on the human cervical cancer cell line (HeLa) and were also analyzed by confocal microscopy. Finally, computer–aided molecular docking studies were carried out to validate the spectroscopic studies which revealed the selective recognition of GC base pairs in the DNA minor groove region by the complexes. 2. Experimental section 2.1. Materials Copper chloride dihydrate, zinc chloride (E. Merck), 2–amino–6–fluorobenzothiazole, D–/L–valine, (Sigma), and salicylaldehyde (Alfa Aesar) were used as received. 6X loading dye (Fermental Life Science), Supercoiled plasmid DNA pBR322 (Genei), Calf thymus DNA (CT–DNA), was purchased from Sigma chemical Co. and Fluka, respectively. All reagents were of the best commercial grade and were used without further purification. 2.2. Methods and instrumentation Carbon, hydrogen and nitrogen contents were determined using Carlo Erba Analyzer Model 1106. Molar conductance was measured at room temperature on a Digsun Electronic conductivity Bridge. Fourier–transform IR (FT−IR) spectra were recorded on an Interspec 2020 FTIR spectrometer. Electronic spectra were recorded on UV–1700 PharmaSpec UV–vis spectrophotometer (Shimadzu). Fluorescence measurements were made on Schimadzu RF–5301 fluorescence spectrophotometer. Data was reported in λmax/nm. The EPR spectra of the copper complexes were acquired on a Varian E 112 spectrometer using X–band frequency (9.1 GHz) at liquid nitrogen temperature in solid

4

state. The 1H and 13C NMR were obtained on a Bruker DRX–400 spectrometer operating at room temperature. Electrospray mass spectra were recorded on Micromass Quattro II triple quadrupol mass spectrometer. Cleavage experiments were performed with the help of Axygen electrophoresis supported by Genei power supply with a potential range of 50–500 Volts, visualized and photographed by Vilber–INFINITY gel documentation system. 2.3. Syntheses 2.3.1. Synthesis of Ligand, L The ligand L was synthesized by adding salicyaldehyde (0.244g, 2mmol) to a stirred methanolic solution (20 ml) of 2–amino–6–fluorobenzothiazole (0.336g, 2mmol) in a 1:1 molar ratio. The reaction mixture was refluxed for 3h and the progress of reaction was monitored by TLC. The yellow precipitate which formed was filtered off under vacuum, washed thoroughly with methanol and dried in vacuo. Yield: 77%, M.p. 138 °C. Anal. Calc. for C14H9N2OFS (%): C, 61.75; H, 3.33; N, 10.29, Found: C, 61.58; H, 3.12; N, 10.17. IR (KBr, cm–1): 3059 ν(O–H), 1611 ν(HC=N), 1511 ν(C=N), 1448 ν(C–N), 821 ν(C–S), 758 (Ar). UV–vis (λmax, nm) in DMSO: 244 (π–π*) and 314 (n–π*). 1H NMR (400 MHz, DMSO–d6, δ, ppm): 11.62 (OH), 9.37 (HC=N), 7.93–6.92 (Ar–H). 13C NMR (100 MHz, DMSO–d6, δ, ppm): 169.46 (C=N), 166.52 (C–O), 160.79 (C–N), 147.80– 107.07 (Ar–C). ESI–MS (m/z): 273.1 [C14H9N2OFS]+. 2.3.2. Synthesis of complexes 1(a and b) The complexes were prepared by a general synthetic method in which a methanolic solution (15 ml) of CuCl2.2H2O (0.170 g, 1 mmol) was added to the methanolic solution (20 ml) of ligand (0.272 g, 1 mmol) in a 1:1 molar ratio which was refluxed for 2 h to

5

obtain a clear green color solution. An equimolar amount of L–/D–valine (0.117 g, 1 mmol) dissolved in MeOH, was added to the above reaction mixture and refluxed for 3 h. The green colored product was isolated, washed with methanol and dried in vacuo. [C19H18N3O3FSCu], 1a: Yield: 69%, M.p. 290 °C. Anal. Calc. for C19H18N3O3FSCu (%): C, 50.60; H, 4.01; N, 9.32. Found: C, 50.41; H, 4.09; N, 9.18. IR (KBr, cm–1): 3178 ν(NH2), 1638 νas(COO–), 1607 ν(HC=N), 1543 ν(C=N), 1448 ν(C–N), 1369 νs(COO–), 849 ν(C–S), 758 (Ar), 464 ν(Cu–O), 420 ν(Cu–N). UV–vis (DMSO, λmax, nm): 255 (π–π*), 310 (n–π*) and 567 (d–d). ESI–MS (m/z): 451.2 [C19H18N3O3FSCu]+. [C19H18N3O3FSCu], 1b: Yield: 66%, M.p. 286 °C. Anal. (%) Calc. for C19H18N3O3FSCu (%): C, 50.60; H, 4.01; N, 9.32. Found: C, 50.48; H, 4. 15; N, 9. 49. IR (KBr, cm–1): 3173 ν(NH2), 1635 νas(COO–), 1603 ν(HC=N), 1539 ν(C=N), 1444 ν(C– N), 1373 νs(COO–), 845 ν(C–S), 722 (Ar), 460 ν(Cu–O), 428 ν(Cu–N). UV–vis (DMSO, λmax, nm): 249 (π–π*), 309 (n–π*) and 577 (d–d).

ESI–MS (m/z): 451.1

[C19H18N3O3FSCu]+. 2.3.3. Synthesis of complexes 2 (a and b) The complexes were synthesized in an identical manner as described for 1 with ZnCl2 (0.136 g, 1 mmol). [C19H18N3O3FSZn], 2a: Yield: 61%, M.p. 263 °C. Anal. (%) Calc. for C19H18N3O3FSZn (%): C, 50.39; H, 4.01; N, 9.28. Found: C, 50.21; H, 4.16; N, 9.12. FT−IR (KBr, cm–1): 3297 ν(NH2), 1623 νas(COO–), 1606 ν(HC=N), 1527 ν(C=N), 1472 ν(C–N), 1396 νs(COO–), 853 ν(C–S), 762 (Ar), 543 ν(Zn–O), 428 ν(Zn–N). UV–vis (DMSO, λmax, nm): 253 (π–π*) and 303 (n–π*). 1H NMR (400 MHz, DMSO–d6, δ, ppm):

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8.96 (HC=N), 7.64–6.93 (Ar–H), 3.20 (–CH–NH2), 2.19 (–CH(CH3)2), 0.96 (–CH3). 13C NMR (100 MHz, DMSO–d6, δ, ppm): 191.99 (C=O), 167.32 (C=N), 158.42 (C–O), 156.88

(C–N),

148.29–107.60

(Ar–C),

19.09

(CH3).

ESI–MS

(m/z):

451.2

[C19H18N3O3FSZn]+. [C19H18N3O3FSZn], 2b: Yield: 57%, M.p. 266 °C. Anal. (%) Calc. for C19H18N3O3FSZn (%): C, 50.39; H, 4.01; N, 9.28. Found: C, 50.58; H, 4.45; N, 9.64. IR (KBr, cm–1): 1613 ν(HC=N), 3301 ν(NH2), 1623 νas(COO–), 1523 ν(C=N), 1468 ν(C–N), 1392 νs(COO–), 853 ν(C–S), 762 (Ar), 543 ν(Zn–O), 428 ν(Zn–N). UV–vis (DMSO, λmax, nm): 257 (π–π*) and 305 (n–π*). 1H NMR (400 MHz, DMSO–d6, δ, ppm): 8.37 (HC=N), 7.82–6.97 (Ar−H), 3.71 (–CH–NH2), 2.54 (–CH(CH3)2), 0.97 (–CH3). 13C NMR (100 MHz, DMSO–d6, δ, ppm): 192.73 (C=O), 167.32, (C=N), 158.42 (C–O), 156.13 (C–N), 147.91–107.35 (Ar–C), 19.08 (CH3). ESI–MS (m/z): 451.6 [C19H18N3O3FSZn]+ . 2.4. DNA binding and cleavage experiments DNA binding experiments which include absorption spectral titrations and emission spectroscopy conformed to the standard methods and practices previously adopted by our laboratory [16–19] whereas DNA cleavage experiment has been performed by the standard protocol as described in [20]. While measuring the absorption spectra an equal amount of DNA was added to both the compound solution and the reference solution to eliminate the absorbance of the CT–DNA itself, and Tris–HCl buffer was subtracted through base line correction. All the experiments involving interaction of the complexes with

CT–DNA

were

performed

in

twice

distilled

buffer

containing

tris(hydroxymethyl)aminomethane and adjusted to pH 7.2 with hydrochloric acid. Solution of CT–DNA in buffer gave a ratio of UV absorbance at 260 and 280 nm of ca.

7

1.9:1 indicating that DNA was sufficiently free of protein. The DNA concentration per nucleotide was determined by absorption spectroscopy with the molar absorption coefficient 6600 M–1cm–1 at 260 nm. 2.5. Antimicrobial studies 2.5.1. Antibacterial activity All the synthesized complexes 1 & 2 (a and b) were screened for their in vitro antibacterial activity against two Gram–negative Escherichia coli (ATCC 25922), Pseudomonas aeruginosa (ATCC 27853)] and two Gram-positive [Staphylococcus aureus (ATCC 25923) and Bacillus subtilis (MTCC 121)] bacterial strains. The agar well diffusion method of Perez et al., also described earlier by Ahmad et al. was adopted for measuring the antibacterial assays. Briefly, all cultures were routinely maintained on NA (nutrient agar) and incubated at 37 °C for overnight. The culture was centrifuged at 1000 rpm and pellets were re-suspended and diluted in sterile Normal Saline Solution to obtain viable count of 105 cfu/ml. Volume of 0.1 ml diluted bacterial culture suspension was spread uniformly with the help of spreader on NA plates. Wells of 8 mm diameter were punched into the agar medium and loaded with different concentrations of the metal complexes. Antibiotic disc, Doxycycline (100 mg/disc) and solvent were used as positive and negative control, respectively. The plates were then incubated for 24 h at 37 °C and the resulting zones of inhibition (mm) were measured. 2.5.2. Antifungal activity All cultures were routinely maintained on SDA and incubated at 28 °C. The inoculums of non–sporing fungi, Candida albicans were performed by growing the culture in SD broth at 37°C for overnight. Volume of 0.1 ml of diluted fungal culture suspension was spread

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with the help of spreader on SDA plates uniformly. Sterile 8 mm discs were impregnated with the test complexes. Wells of 8 mm size were cut and loaded with different concentrations of the complexes. Antibiotic disc, nystatin (100 mg/disc) was used as positive control. C. albicans plates were incubated at 37 °C for 18−48 h and antifungal activity was determined by measuring the diameters of the inhibition zone (mm). 2.6. Cell Proliferation and viability assay To measure cell viability and growth MTT assay was done. In this assay, cells seeded in 96–well plates (5x103 cells/well) in duplicate were cultured in complete DMEM medium. Cells were treated with different concentrations (5, 10, 15, 20, 25 µM) of 1a and 1b separately for 24 hours. The cells, after treatment were incubated with MTT for 2 hours. The yellow tetrazolium salt (MTT) is reduced in metabolically active cells to form insoluble purple formazan crystals, which are solubilized by the addition of DMSO (Sigma). The colour was then quantified by spectrophotometric measurement at 570 nm wavelength on a microtiter plate reader (Bio–Rad model 680 microplate reader). 2.7. Maintenance of cell lines HeLa cell line was maintained in DMEM (Dulbeco’s Modified Eagle Medium) medium supplemented with 10% FBS (Fetal Bovine Serum) and 1% antibiotics and antimycotic (Himedia) as per manufacturer′s protocol. Cells were incubated in CO2 incubator at 5% CO2 and 37 °C temperature. 2.8. Cell imaging studies with HeLa cells using confocal microscopy In order to analyze the efficacy and utilization of 1a and 1b as an anti–cancerous probe through confocal fluorescence imaging of HeLa cell line treated with 1a and 1b separately. HeLa cells were incubated separately with 15 µM of 1a and 1b for 60 min in

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dark. Cells were washed for 5 min with 1X PBS (Phosphate buffered saline, pH 7.4) twice. Cells collections after each step was done by centrifugation at 2000 rpm for 2 min. Cells were mounted with DABCO on Poly–lysine coated slide. Control for complex 1b and complex 1b were also taken. Slides were observed under 20X oil objective lens of Zeiss LSM 510 META confocal microscope. 2.9. Molecular docking Studies The rigid molecular docking studies were performed by using HEX 8.0 software [21] which is an interactive molecular graphics program to calculate and display feasible docking modes of a pairs of protein, enzymes and DNA. Structures of the complexes were sketched by ChemSketch (http://www.acdlabs.com), energy minimized and converted into pdb format from mol format by the Mercury software. The crystal structure of the B–DNA dodecamer d(CGCGAATTCGCG)2 (PDB ID: 1BNA) and was downloaded from the protein data bank (http://www.rcsb.org./pdb). All calculations were carried out on an Intel pentium4, 2.4 GHz based machine running MS Windows XP SP2 as operating system. Visualization of the docked pose has been done by using CHIMERA (www.cgl.ucsf.edu/chimera), PyMol (http://pymol.sourceforge.net/) and Discovery Studio molecular graphics program. 3. Results and discussion 3.1. Synthesis and characterization The complexes 1 & 2 (a and b) were synthesized by reacting 1:1:1 stoichiometric amounts of Schiff base ligand L, Cu(II)/Zn(II) chloride and L–/D–valine as depicted in Scheme I (the energy minimized three dimensional ball and stick model of complexes 1a and 2a are shown in Fig. S1). The complexes were thoroughly characterized by analytical

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and spectroscopic data. All the complexes, 1 & 2 (a and b) are air stable for extended periods and soluble in DMF and DMSO solvents. Molar conductance values of both the complexes in DMSO (1x10–3 M) at 25 ºC suggest their non–electrolytic nature (12–20 Ω– 1

cm2 mol–1). On the basis of spectral studies, the coordination geometry of copper(II) and

zinc(II) was proposed to be square planar and tetrahedral, respectively. The IR spectrum of a free ligand L revealed a characteristic band at 3439 cm–1 corresponding to phenolic –OH group which was found absent in the IR spectra of complexes, 1 & 2 (a and b) implicating its coordination to central Cu(II)/Zn(II) metal ions via deprotonation. Another important band in the IR spectrum of L appeared at 1611 cm–1 due to azomethine group stretching vibration ν(C=N) which was shifted to 1613−1603 cm–1 upon complexation, implying its coordination via azomethine nitrogen to the metal center [22]. Moreover, in complexes 1 & 2 (a and b) the presence of IR frequency bands in the region ca. 1638–1623 cm−1 and 1392–1369 cm−1 were attributed to antisymmetric and symmetric COO stretching vibrations, respectively which validated the presence of D/L−valine ligand scaffold. However, these bands were found in the range 1660 and 1430 cm−1, respectively in the free amino acid, indicating the coordination of carboxylate group (COOH) to metal ion via deprotonation. Additionally, the formation of complexes 1 & 2 (a and b) was also ascertained by the presence of medium intensity bands in the region 543–460 and 428–420 cm–1 assigned to ν(Cu/Zn– O) and ν(Cu/Zn–N) respectively. The NMR spectra of the Schiff base ligand, L and its diamagnetic complexes 2 (a and b) were recorded in DMSO–d6 solvent. In the 1H NMR spectrum the signal of the phenolic (OH) proton in the ligand observed at 11.62 ppm was absent in the spectra of the

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complexes 2 (a and b) confirming the coordination of phenolic group to metal center via deprotonation. Similarly, absence of signal in the region 12.0–10.0 ppm in complexes 2 (a and b) due to the –OH proton of carboxylic group supported the unidentate coordination of the –COOH of L–/D–valine through deprotonation [23]. The characteristic azomethine proton signal at 9.37 ppm in ligand scaffold was shifted to 8.38 and 8.37 ppm in complexes, 2a and b, respectively, indicating the coordination through azomethine nitrogen [24]. A broad multiplet observed in the range of 7.83–6.80 ppm was attributed to the merging of aromatic ring and NH2 protons in the same region [25]. The NMR spectra revealed characteristic signals of the –CH–NH2, –CH(CH3)2 and –CH3 protons in the range of 3.71–3.20, 2.54–2.19 and 0.97–0.96 ppm, respectively. The 13C NMR spectrum of the ligand exhibited signals at 169.46 (HC=N), 166.52 (C–O), 160.79 (C–N), 158.38 (C–S), and 147.80–107.07 (Ar–C). However, the 13C NMR spectra of complexes 2 (a and b) exhibited various resonance peaks centered at 167.32, 158.42, 107.60–148.29, 191.99 and 167.49, 158.49, 107.35–147.91, 192.73 which were assigned to HC=N, C–O–Zn, Ar–C, O–C=O, respectively. In addition, the methyl carbon revealed its resonance signature at 19.09–18.18 ppm. The solid state X–band EPR spectra of complexes 1 (a and b) was recorded at room temperature under the magnetic field strength 3000 ± 1000 G using tetracyanoethylene as a field marker as shown in Fig. 1 (a and b). The EPR spectra of 1 (a and b) displayed an isotropic signal and exhibits axial symmetrical line shape with g|| = 2.11 and g⊥ = 2.06 and gav =2.07 and g|| = 2.31 and g⊥ = 2.05 and gav = 2.13, respectively computed from the formula gav2 = g||2+2g⊥2/3. These parameters were consistent with the square planar geometry of the copper(II) metal ion [26,27].

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The electronic absorption spectra of ligand L displayed high–energy bands at 244 and 316 nm corresponds to π→π* transition of the aromatic ring and n–π* transitions of the C=N groups, respectively. However, these bands were shifted to ca. 249–257 and 314– 301 nm in the complexes attributed to intraligand and LMCT transitions of coordinated ligand. Additionally, the electronic spectra of complexes 1a and b displayed low intensity broad bands at ca. 567 and 572 nm, respectively in the visible region attributable to d–d [2B1g → 2E1g (υ1)(dx2–y2 →dyz) transitions, typical for distorted square–planar geometry around the Cu(II) metal ion [28]. 4. DNA Binding Studies 4.1. Electronic absorption titration Electronic absorption spectroscopy is one of the most useful techniques to understand the drug–DNA binding studies. Upon concomitant addition of CT−DNA (0.00–5.5 x 10–5 M), to Schiff base ligand, L and its L–/D–enantiomeric complexes 1 & 2 (a and b) of fixed concentration (6.67 x 10–5 M), an increase in the absorbance (hyperchromic effect) of the intraligand absorption band was observed with a minor red shift in the absorption maxima (Fig. 2a–e). The observed spectral behavior clearly rule out intercalative binding of the complexes to DNA, since intercalation leads to hypochromism in the spectral bands. The spectral ‘hyperchromic effect’ was attributed to contraction and overall damage caused to the secondary structure of DNA double helix [29,30], while the red shift has been associated with the decrease in the energy gap between the highest and lowest molecular orbitals (HUMO and LUMO) after binding of the complexes to DNA [31]. Hyperchromism with less or no shift in absorbance is consistent with groove

13

binding, therefore in these complexes it can be attributed to external contact (surface binding) with the DNA duplex. To quantify the enantioselective approach of the complexes, the DNA binding affinities of 1 & 2 (a and b) with CT DNA, the intrinsic binding constants Kb were determined by using Wolfe–Shimer Eq. (1) [32] by monitoring the change in the absorbance of the π–π* bands with increasing concentration of CT–DNA. [DNA]/εa–εf = [DNA]/εb–εf + 1/Kb│εb–εf │

(1)

The binding constant (Kb) were given in Table 1, which followed the order 1a > 2a > 1b > 2b, clearly indicating the marked enantioselective approach of the complexes; emphasizing the stronger binding affinity of L–complexes towards DNA target in comparison to D–complexes. 4.2. Fluorescence spectroscopic studies In the emission spectra, the Schiff base ligand, L and its enantiomeric complexes 1 & 2 (a and b) emit luminescence in Tris–HCl buffer at ambient temperatures with a maxima appearing at ca. 290–310 nm. Upon addition of increasing concentration of CT–DNA (0– 33.3 x 10–6 M) to the fixed amount of L/complex concentration (6.67 x 10–6 M), there was a gradual enhancement in the fluorescence intensity of the complexes (Fig. 3a–e). The observed enhancement could be due to relatively non–polar environment of the bound metal complex in the presence of DNA, such that the complexes were less deeply inserted inside the hydrophobic pockets or grooves of CT DNA [33]. In general cationic complexes bind to DNA non–covalently through electrostatic attraction with the oxygen phosphate backbone of DNA and thus precluding substantial overlap with the base pairs leading to higher emission intensity, and the mobility of the complex is restricted at the

14

binding site ultimately leading to decrease in vibrational mode of relaxation [34]. The binding constant determined for L and the complexes 1a, 1b, 2a and 2b by Scatchard equation [35] were 6.50 x 103, 2.77 x 105, 5.01 x 104, 7.04 x 104 M–1 and 3.56 x 104 M–1, respectively, consistent with the results obtained from UV–vis spectral titration studies. 4.3. Ethidium Bromide Displacement Assay In order to further investigate the interaction mode of Schiff base ligand, L and enantiomeric complexes 1 & 2 (a and b) with DNA, a competitive binding experiment using EB as a probe was carried out. EB (3,8–diamino–5–ethyl–6–phenylphenanthrium bromide) is a conjugate planar molecule with very weak fluorescence intensity due to fluorescence quenching of the free EB by solvent molecules but it is greatly enhanced when EB is specifically intercalated into the adjacent base pairs of double stranded DNA. The enhanced fluorescence can be quenched upon the addition of the second molecule which could replace the bound EB or break the secondary structure of the DNA. On addition of L/complexes 1 & 2 (a and b) to CT DNA pretreated with EB ([DNA]/[EB] = 1) a significant reduction in the emission intensity (Fig. 4a–e) was observed , indicating that the replacement of the EB fluorophore by these complexes, which results in a decrease of the binding constant of ethidium bromide to DNA. As there was incomplete quenching of the EthBr–induced emission intensity, the intercalative binding mode was ruled out. The extent of quenching of the emission intensity gives a measure of the binding propensity of the interacting molecule to CT DNA. The Stern–Volmer quenching constant value, Ksv, obtained as a slope of I0/I vs r ([complex]/[DNA]) were evaluated for L/complexes 1a, 1b, 2a and 2b, and were found to be 0.85, 2.32, 1.45, 2.01 and 1.25, respectively. The high Ksv values of L–form of complexes indicated their superior binding

15

with CT–DNA relative to D–form of complexes. The enantiomeric preference of L–form towards DNA has also been demonstrated by similar studies in previously reported experiments from our laboratory [36]. 4.4. DNA cleavage activity 4.4.1. Concentration dependent DNA cleavage It is known that plasmid DNA cleavage produces relaxation of the supercoiled circular conformation to the nicked circular and/or linear conformations. When circular pBR322 DNA is subjected to gel electrophoreses, relatively fast migration is observed for the intact supercoiled form (Form I). If scission occurs on one strand (nicking), the supercoiled form will relax to generate a slower–moving open circular form (Form II). If both strands are cleaved, a linear form (Form III) is generated that migrates in between Forms I and II. The DNA cleavage ability of complexes 1a and 2a was studied by agarose gel electrophoresis using supercoiled pBR322 plasmid DNA as a substrate due to their high binding ability (Fig. 5a and b). In a concentration dependent cleavage experiment of complex 1a, significant conversion of supercoiled form (Form I) of pBR322 DNA to nicked form (Form II) was observed (Lanes 2–4). The most impressive cleavage feature of complex 1a was the appearance of linear form (Form III) before the disappearance of supercoiled form (Form I) of pBR322 plasmid (Lanes 5 and 6). This phenomenon indicates that complex 1a was capable of performing direct double–strand scission. However, complex 2a was able to convert supercoiled form (Form I) to nicked open circular form (Form II) with the increasing concentration (Fig. 5b, Lanes 2–6) without

16

appearance of linear form (Form III) of pBR322 plasmid DNA which indicated that no double strand DNA cleavage was observed. Thus, it was clear that cleavage of pBR322 DNA was directly correlated to higher DNA binding ability of L–enantiomeric Cu(II) complex 1a. The gel pattern of 1a with the appearance of linear form (Form III) further validated the direct double–strand scission of DNA; as a consequence, this complex could be better suited for therapeutic applications particularly, in cancer chemotherapeutics. 4.4.2. DNA cleavage in presence of activators and reactive oxygen species The interaction between metal complexes and dioxygen or redox reagents are believed to be a major cause of DNA damage [37]. Therefore, DNA cleavage activity of complex 1a was evaluated in presence of H2O2, 3–mercaptopropionic acid (MPA), ascorbate (Asc) and glutathione (GSH) (Fig. 6; Lane 2–5). Many copper (II) complexes can cleave DNA more efficiently in presence of exogenous agents. The cleavage activity was significantly enhanced in presence of these activators and follow the order MPA > H2O2 > Asc ≈ GSH. Thus complex 1a in presence of MPA exhibited significant DNA cleavage activity, followed by complete degradation of DNA was observed, suggesting that these activators play an important role to aid the copper(II) complex in DNA cleavge. To postulate a possible mechanism responsible for DNA cleavage mediated by complexes 1a and 2a, gel electrophoretic study with standard radical scavengers, viz., DMSO and ethanol as hydroxyl radical scavengers, sodium azide and SOD as singlet oxygen scavenger and superoxide radical scavenger, respectively was carried out. The addition of DMSO and EtOH to complex 1a inhibited the DNA cleavage activity (Lanes 6 and 7) which was indicative of the involvement of diffusible (HO· ) hydroxyl radicals as

17

one of the ROS responsible for DNA breakage. Similarly, addition of sodium azide decreased the cleavage efficiencies (Lane 8) which revealed that 1O2 was the activated oxygen intermediate responsible for the cleavage. However, there was enhancement of DNA cleavage in case of SOD (superoxide dismutase) (Lane 9) followed by the appearance of linear form (Form III) which demonstrated the non–involvement of superoxide radical in DNA scission suggestive. Groove binding preference of 1a in presence of minor groove binding agent DAPI and major groove binding agent methyl green (MG) were performed to probe the potential interacting site with plasmid pBR322 DNA (Lanes 10 and 11). When supercoiled pBR322 was treated with DAPI or methyl green prior to the addition of complex, the cleavage reaction mediated by 1a was quenched in presence of DAPI while it enhances in the presence of MG indicating the minor groove–binding preference of complex 1a. In case of complex 2a, addition of DMSO and ethyl alcohol (Fig. 7, Lanes 2 and 3), DNA cleavage was inhibited suggesting the possibility of hydroxyl radical as one of the active species. Thus, free radicals participate in the oxidation of the deoxyribose moiety, followed by hydrolytic cleavage of the sugar phosphate back bone in the absence of scavengers. The cleavage ability was also inhibited in presence of NaN3 and SOD (Lanes 4 and 5), indicating the involvement of the singlet oxygen radical and superoxide anion radical in the mechanistic pathway of DNA cleavage. The cleavage reaction mediated by 2a was inhibited in presence of DAPI and methyl green indicating that complex 2a has ability to interact with minor and major groove of pBR322 DNA (Lane 6 and 7).

18

5. Antimicrobial activity The complexes 1 & 2 (a and b) were screened for the in vitro antibacterial activity against Gram–positive B. subtilis [MTCC 121], S. aureus [NRRLB 767] and Gram– negative E. coli [K 12], P. aeruginosa, and in vitro antifungal activity against C. albicans and the relevant data are presented in Table 2. The diameter of the zone of inhibition (mm) was used to compare the antimicrobial activity of the all complexes with the commercial drug (Doxycycline and Nystatin were used as a reference drugs for antibacterial and anti–fungal activity, respectively). The results revealed that complexes exhibited varying degree of inhibitory effects on the growth of bacterial and fungal strains may be due to the effect of the metal ion on the cell metabolism. The complex 1a showed remarkably good anti−bacterial activity against B. subtilis and E. coli with zone diameter 19 and 22 mm, respectively, while moderate activity against S. aureus and P. aeruginosa with zone diameter 17 and 16 mm, respectively. While complex 2b exhibited pronounced activity against bacteria S. aureus, E. coli and Gram–negative bacteria P. aeruginosa with zone diameter 20, 18 and 17 mm, respectively. The synthesized complexes 1 & 2 (a and b) were also screened for their antifungal activity against C. albicans. Interestingly the L–enantiomeric complexes, 2a and 2b exhibited prominent antifungal activity against the C. albicans with zone diameter 23 and 21 mm, respectively. The significant antimicrobial activity of the complexes can be attributed to the fact that the lipid membranes that surrounds the cell favors the passage of only the lipid–soluble materials which makes liposolubility as an important factor that controls the antibacterial activity. On chelation with metal ions, the polarity of metal ion will be reduced to a

19

greater extent due to the overlap of the ligand orbital and partial sharing of the positive charge of the metal ion with donor groups. Furthermore, the chelation increases the delocalization of π–electrons over the whole chelate ring and also enhances the lipophilicity of the metal complexes [38,39]. 6. MTT assay and nuclear blebbing in HeLa live cells The in vitro cytotoxicity of complexes 1a and 1b on the human cervical cancer cell line (HeLa) was examined using the MTT assay. In this assay, cell count of 5x103 cells/well were seeded in 96–well plates in duplicate were cultured in Dulbecco’s modified Eagle’s medium. Cells were treated with different concentrations of (5, 10, 15, 20, 25 µM) of 1a and 1b separately for 24 hours which implicated that more than 50 % cells were viable in 15 µM concentration while at ≥ 20 µM cell viability was compromised at varying degree (Fig. 8a and b). The treatment of 15 µM of complexes 1a and 1b with HeLa cells resulted nuclear blebbing as depicted from PI stained HeLa cells (Fig. 9). No such blebbing was observed in control. Complexes 1a and 1b have potential to initiate nuclear blebbing in HeLa cancer cell line suggesting that cytotoxicity induced by the metal complexes resulted in apoptotic cell death. Similar studies on human HeLa cells in the copper (II) complexes of quinoline derivatives appended with benzothiazole substituents revealed cell death with IC50 values in the micromolar range of 6–53 µM [40] 7. Molecular Docking Molecular docking studies were undertaken in order to provide profound insight into complex–DNA interaction and conclusively complement the spectroscopic studies. Energetically favorable docked poses were obtained from the rigid molecular docking

20

experiments carried out between the optimized energy–minimized structures of complexes 1 & 2 (a and b) with the DNA duplex of sequence d(CGCGAATTCGCG)2 dodecamer (PDB ID : 1BNA). The results revealed that the complexes fitted snugly into the curved contour of the DNA target into the minor groove in close proximity of the G–C rich region. Minor groove is a preferential binding site for small molecules as the convex surface of minor groove floor complements the concave shape of typical minorgroove binding molecules [41] In general, the deeper and narrower minor groove is optimal for accommodating the shape of the complexes and also maximizes the stabilized van der Waal’s contacts, hydrogen bonding and electrostatic interactions [42]. As depicted in Fig. 10 (a and b) and Fig. S2, it is clear that the complexes slightly bend the DNA in such a way that the part of the planar aromatic rings of benzothiazole Schiff base pharmacophore make a favorable stacking interactions between DNA base pairs that bring about van der Waals and hydrophobic interactions with DNA functional groups of the minor groove. The resulting relative binding energy of docked chiral metal complexes 1a, 1b, 2a and 2b with DNA was found to be −286.3, −242.4, −259.2 and −233.2 KJ mol−1, respectively. The more negative relative binding, the more is the binding affinity which correlated well with the experimental DNA binding results. 8. Conclusion New enantiomeric Cu(II) and Zn(II) L–/D–fluorobenzothiazole Schiff base–valine complexes, 1 & 2 (a and b) have been synthesized and thoroughly characterized by various spectroscopic methods. Preliminary in vitro DNA binding studies of Schiff base ligand, L and its enantiomeric Cu(II)/Zn(II) complexes were carried out by absorption

21

and fluorescence spectroscopic titration to establish whether they demonstrated any enantioselectivity in DNA binding profile. Since enantiomeric effects are quite subtle and sensitive to the environment, therefore, it was desirable to carry out the comparative DNA binding studies with enantiomeric L–and D–complexes of Cu(II) and Zn(II). The results revealed that L–enantiomer of Cu(II) complex 1a exhibits highest DNA binding propensity and cleavage activity, possessing distinct chiral preference over its D– enantiomer. All the complexes were screened for in vitro antimicrobial activity and exhibited varying degree of inhibitory effects on the growth of bacterial and fungal strains. Furthermore, cytotoxicity of the complexes 1a and 1b was evaluated on the human cervical cancer cell line (HeLa) which implicated that more than 50 % cells were viable at 15 µM and these results were further validated by cell imaging studies that clearly showed the nuclear blebbing characteristic of apoptotic cell death mechanism. Molecular docking studies were carried out to authenticate the spectroscopic studies which revealed that the complexes recognize the narrow minor groove region situated within the GC region of the DNA duplex. Acknowledgements The authors are thankful to SAIF, Panjab University, Chandigarh, for NMR and ESI– Mass facility; STIC, Cochin University, Cochin, for providing elemental analysis and IIT Bombay for EPR studies. We are grateful to Dr. Iqbal Ahmad, Department of Agricultural Microbiology, Faculty of Agricultural Sciences, Aligarh Muslim University for carrying out antimicrobial studies. The authors also acknolwledge the generous financial support from DST−PURSE programme and DRS−1 (SAP) from UGC, New Delhi.

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Appendix A. Supplementary Data Additional Supplementary Data may be found with the online version of this article. References [1] R. Dua, S. Shrivastava, S.K. Sonwane, S.K. Srivastava, Pharmacological Significance of Synthetic Heterocycles Scaffold, Adv. Biol. Res. 5 (2011)120–144. [2]

B. Soni, M.S. Ranawat, R. Sharma, A. Bhandari, S. Sharma, Synthesis and evaluation of some new benzothiazole derivatives as potential antimicrobial agents, Eur. J. Med. Chem. 45 (2010) 2938–2942.

[3] S. Shafi, M.M. Alam, N. Mulakayala, C. Mulakayala, G. Vanaja, A.M. Kalle, R. Pallu, M.S. Alam, Synthesis of novel 2–mercapto benzothiazole and 1,2,3–triazole based bis–heterocycles: Their anti–inflammatory and anti–nociceptive activities, Eur. J. Med. Chem. 49 (2012) 324–333. [4] D. Bhavsar, J. Trivedi, S. Parekh, M. Savant, S. Thakrar, A. Bavishi, A. Radadiya, H. Vala, J. Lunagariya, M. Parmar, L. Paresh, R. Loddo, A. Shah, Synthesis and in vitro anti–HIV

activity

of

N–1,3–benzo[d]thiazol–2–yl–2–(2–oxo–2H–chromen–4–

yl)acetamide derivatives using MTT method, Bioorg. Med. Chem. Lett. 21 (2011) 3443–3446 [5]

F. Delmas, A. Avellaneda, C. Di Giorgio, M. Robin, E.D. Clercq, P. Timon– David, J.P. Galy, Synthesis and antileishmanial activity of (1,3–benzothiazol–2– yl)amino–9–(10H)–acridinone derivatives, Eur. J. Med. Chem. 39 (2004) 685–690.

[6] G.A. Pereira, A.C. Massabni, E.E. Castellano, L.A.S. Costa, C.Q.F. Leite, F.R. Pavan, A. Cuin, A broad study of two new promising antimycobacterial drugs: Ag(I) and Au(I) complexes with 2–(2–thienyl)benzothiazole, Polyhedron 38 (2012) 291–296. [7] A. Burger, S.N. Sawhey, Antimalarials. III. Benzothiazole amino alcohols, J. Med. Chem. 11 (1968) 270–273. [8] I. Hutchinson, M.–S. Chua, H.L. Browne, V. Trapani, T.D. Bradshaw, A.D. Westwell, M.F.G Stevens, Antitumor benzothiazoles. 14. Synthesis and in vitro biological properties of fluorinated 2–(4–aminophenyl)benzothiazoles. J. Med. Chem. 44 (2001) 1446–1455.

23

[9] C.–O. Leong, M. Gaskell, E.A. Martin, R.T. Heydon, P.B. Farmer, M.C. Bibby, P.A. Cooper, J.A. Double, T.D. Bradshaw, M.F.G. Stevens, Antitumour 2–(4– aminophenyl)benzothiazoles generate DNA adducts in sensitive tumour cells in vitro and in vivo, Br. J. Cancer 88 (2003) 470–477. [10] T.D. Bradshaw, M.–S. Chua, S. Orr, C.S. Matthews, M.F.G Stevens, Mechanisms of acquired resistance to 2–(4–aminophenyl)benzothiazole, Br. J. Cancer 83 (2000) 270–277. [11] M.–S. Chua, E. Kashiyama, T.D. Bradshaw, S.F. Stinson, E. Brantley, E.A, Sausville, M.F.G, Stevens, Role of CYP1A1 in modulation of antitumour properties of the novel agent 2–(4–amino–3–methylphenyl)benzothiazole in human breast cancer cells, Cancer Res. 60 (2000) 5196–5203. [12] D.–F. Shi, T.D. Bradshaw, S. Wrigley, C.J. McCall, P. Lelieveld, I. Fichtner, M.F.G Stevens, Antitumour benzothiazoles 3 Synthesis of 2–(4–aminophenyl)– benzothiazoles and evaluation of their activities against breast cancer cell lines in vitro and in vivo, J. Med. Chem. 39 (1996) 3375–3384. [13] (a) M.A. Jakupec, M. Galanski, V.B. Arion, C.G. Hartinger, B.K. Keppler, Antitumour metal compounds: more than theme and variations, Dalton Trans. (2008) 183–194; (b) A.T. Balaban, Aromaticity as a cornerstone of heterocyclic chemistry, Chem. Rev. 104 (2004) 2777−2812; (c) F. Arjmand, I. Yousuf, T.B Hadda, L. Toupet, Synthesis, crystal structure and antiproliferative activity of Cu(II) nalidixic acid–DACH conjugate: Comparative in vitro DNA/RNA binding profile, cleavage activity and molecular docking studies, Eur. J. Med. Chem. 81 (2014) 76– 88. [14] (a) B. Rosenberg, L. VanCamp, J.E. Trosko, V.H. Mansour, Platinum compounds: a new class of potent antitumour agents, Nature 222 (1969) 385–386; (b) Y. Jung, S.J. Lippard, Direct cellular responses to platinum–induced DNA damage, Chem. Rev. 107 (2007) 1387–1407. [15]

J. Reedijk, New clues for platinum antitumor chemistry: kinetically controlled metal binding to DNA, Proc. Natl. Acad. Sci. USA 100 (2003) 3611–3616.

[16]

J. Marmur, A procedure for the isolation of deoxyribonucleic acid from micro organisms, J. Mol. Biol. 3 (1961) 208–218.

24

[17]

M.E. Reicmann, S.A. Rice, C.A. Thomas, P. Doty, A further examination of the molecular weight and size of de–oxypentose nucleic acid, J. Am. Chem. Soc. 76 (1954) 3047–3053.

[18]

A. Wolfe, G.H. Shimer, T. Meehan, Polycyclic aromatic hydrocarbons physically intercalate into duplex regions of denatured DNA, Biochemistry 26 (1987) 6392– 6396.

[19]

J.R. Lakowiez, G. Webber, Quenching of fluorescence by oxygen. Probe for structural fluctuations in macromolecules, Biochemistry 12 (1973) 4161–4170.

[20]

F. Arjmand, I. Yousuf, M. Afzal, L. Toupet, Design and synthesis of new Zn(II) nalidixic

acid–DACH

based

Topo–II

inhibiting

molecular

entity:

Chemotherapeutic potential validated by its in vitro binding profile, pBR322 cleavage activity and molecular docking studies with DNA and RNA molecular targets, Inorg. Chim. Acta 421 (2014) 26– 37. [21]

(a) D. Mustard, D.W. Ritchie, Docking essential dynamics eigen structures, Proteins Struct. Funct. Bioinf. 60 (2005) 269–274. (b) S. J. Raharjo, C. Mahdi, N. Nurdiana, T. Kikuchi, F. Fatchiyah, Binding Energy Calculation of Patchouli Alcohol Isomer Cyclooxygenase Complexes Suggested as COX-1/COX-2 Selective Inhibitor, Adv. Bioinform. 2014 (2014) 1–12.

[22]

K. Nakamoto, Infrared and Raman Spectra of Inorganic and Coordination Compounds, fifth ed., John Wiley and Sons, New York, 1997. pp. 15–20.

[23]

F. Arjmand, A. Jamsheera, Synthesis, characterization and in vitro DNA binding studies of tin(IV) complexes of tert–butyl 1–(2–hydroxy–1–phenylethylamino)– 3–methyl–1–oxobutan–2–yl carbamate, J. Organomet. Chem. 696 (2011) 3572– 3579.

[24]

H.D. Yin, Q.B. Wang, S.C. Xue, Synthesis and structural characterization of diorganotin(IV) esters of salicylidene–amino acids, J. Organmet. Chem. 689 (2004) 2480–2485.

[25]

A. Lalehzari, J. Desper, C.J. Levy, Double–stranded monohelical complexes from an unsymmetrical chiral Schiff–base ligand, Inorg. Chem. 47 (2008) 1120–1126.

25

[26]

R. Alizadeh, M. Afzal, F. Arjmand, In vitro DNA binding, pBR322 plasmid cleavage and molecular modeling study of chiral benzothiazole Schiff–base– valine Cu(II) and Zn(II) complexes to evaluate their enantiomeric biological disposition for molecular target DNA, Spectrochim. Acta, Part A, 131 (2014) 625–635.

[27]

A.M. Herrera, R.J. Staples, S.V. Kryatov, A.Y. Nazarenko, E.V. Rybak– Akimova, Nickel(II) and copper(II) complexes with pyridine–containing macrocycles bearing an aminopropyl pendant arm: synthesis, characterization, and modifications of pendant amino group, Dalton Trans. (2003) 846–856.

[28]

F. Arjmand, M. Muddassir, Design and synthesis of heterobimetallic topoisomerase I and II inhibitor complexes: In vitro DNA binding, interaction with 5′–GMP and 5′–TMP and cleavage studies, J. Photochem. Photobiol. B: Biol. 101 (2010) 37–46.

[29]

Y. An, M.–L. Tong, L.–N. Ji, Z.–W. Mao, Double–strand DNA cleavage by copper complexes of 2,2′–dipyridyl with electropositive pendants. Dalton Trans. (2006) 2066–2071.

[30]

J. Liu, H. Zhang, C. Chen, H. Deng, T. Lu, L. Ji, Interaction of macrocyclic copper(II) complexes with calf thymus DNA: effects of the side chains of the ligands on the DNA–binding behaviors, Dalton Trans. (2003) 114–119.

[31]

F. Arjmand, M. Aziz, Synthesis and characterization of dinuclear macrocyclic cobalt(II), copper(II) and zinc(II) complexes derived from 2,2,2′,2′–S,S[bis(bis– N,N–2–thiobenzimidazolyloxalato–1,2–ethane)]: DNA binding and cleavage studies, Eur. J. Med. Chem. 44 (2009) 834–844.

[32]

A. Wolfe, G.H. Shimmer, T. Meehan, Polycyclic aromatic hydrocarbons physically intercalate into duplex regions of denatured DNA, Biochemistry 26 (1987) 6392–6396.

[33]

C.V. Kumar, J.K. Barton, N.J. Turro, Photophysics of ruthenium complexes bound to double helical DNA, J. Am. Chem. Soc. 107 (1985) 5518–5523.

[34]

B.L. Hauenstein, W.J. Dressick, S.L. Buell, J.N. Demas and B.A. De–Graff, A new probe of solvent accessibility of bound photosensitizers. Ruthenium(II) and

26

osmium(II) photosensitizers in sodium lauryl sulfate micelles, J. Am. Chem. Soc. 105 (1983) 4251–4255. [35]

G. Scatchard, The attractions of proteins for small molecules and ions, Ann. NY Acad. Sci. 51 (1949) 660–672.

[36] F. Arjmand, M. Muddassir, R.H. Khan, Chiral preference of L–tryptophan derived metal–based antitumor agent of late 3d–metal ions (Co(II), Cu(II) and Zn(II)) in comparison to D– and DL–tryptophan analogues: their in vitro reactivity towards CT DNA, 5′–GMP and 5′–TMP, Eur. J. Med. Chem. 45 (2010) 3549–3557. [37]

C.A. Detmer III, F.V. Pamatong, J.R. Bocarsly, Molecular Recognition Effects in Metal Complex Mediated Double–Strand Cleavage of DNA: Reactivity and Binding Studies with Model Substrates, Inorg. Chem., 36 (1997) 3676–3682.

[38]

M.I. Rehman, W.J. Choudhary, Thomsen Bioassay Techniques for Drug Development, Harwood Academic Publishers, Amsterdam, The Netherlands, 2001. pp. 9.

[39]

P.K. Panchal, P.B. Pansuriya, M.N. Patel , In-vitro biological evaluation of some ONS

and

NS

donor

Schiff's

bases

and

their

metal

complexes

J. Enzyme Inhib. Med. Chem. 21 (2006) 453−458. [40]

R.A. Steiner, D. Foreman, H.X. Lin, B.K. Carney, K.M. Fox, L. Cassimeris, J.M. Tanski, L.A. Tyler, Synthesis, characterization, crystal structures and biological activity of set of Cu(II) benzothiazole complexes: Artificial nucleases with cytotoxic activities, J. Inorg. Biochem. 137 (2014) 1–11.

[41] S. Neidle, DNA minor-groove recognition by small molecules, Nat. Prod. Rep. 18 (2001) 291–309. [42] I.J. Simpson, M. Lee, A. Kumar, D.W. Boykin, S. Neidle, DNA Minor Groove Interactions and the Biological Activity of 2,5–Bis–[4–(N–alkylamidino)phenyl] Furans, Bioorg. Med. Chem. Lett. 10 (2000) 2593−2597.

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Tables Table 1: The binding constant (Kb) values of ligand L and complexes 1 & 2 (a and b) on interaction with CT–DNA (mean standard deviation of ±0.12). Kb (M–1) 8.49 x 103 2.05 x 105 5.91 x 104 7.12 x 104 4.58 x 104

L/complexes L 1a 1b 2a 2b

Table 2: The in vitro antimicrobial activity (antibacterial and antifungal ) of complexes 1 & 2 (a and b). Compounda

B. subtilis 19 17 17 16 24 –

1a 1b 2a 2b Doxycyclineb Nystatinc a Compound 800 µg/ml b Doxycycline100 µg/disc c Nystatin 100 µg/disc – Lack of growth inhibition area

Zone of inhibition (mm) S. aureus E. coli P. aeruginosa 17 22 16 15 18 14 20 17 17 17 16 13 21 20 19 – – –

C. albicans 23 15 21 13 – 20

FIGURE CAPTIONS Fig. 1. X–band polycrystalline powder EPR spectrum of complex 1 (a and b) at room temperature. Fig. 2. Absorption spectral traces of (a) Ligand L and complex (b) 1a (c) 1b (d) 2a (e) 2b in 5 mM Tris–HCl/50 mM NaCl buffer at pH 7.2 upon progressive addition of CT−DNA. [Complex] =6.67 x 10–5 M, [DNA]= 0–5.55 x 10–5 M. Fig. 3. Emission spectra of (a) Ligand L and complex (b) 1a (c) 1b (d) 2a (e) 2b in the presence of DNA in 5 mM Tris–HCl/50 mM NaCl at pH 7.2 buffer. Arrows show the intensity changes upon increasing concentration of the DNA.

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Fig 4. Emission quenching spectra of (a) Ligand, L and complex (b) 1a (c) 1b (d) 2a (e) 2b, with increasing concentration of quencher ethidium bromide, in the absence and presence of CT DNA in buffer 5mM Tris–HCl/50 mM NaCl, pH = 7.2 at room temperature. Fig. 5. Agarose gel electrophoresis diagram showing cleavage of pBR322 supercoiled DNA (300 ng) by complex (a) 1a and (b) 2a at 310 K after 45 min of incubation; Lane 1, DNA control; Lane 2, 10 µM of complex + DNA; Lane 3: 20 µM of complex + DNA; Lane 4: 30 µM of complex + DNA; Lane 5: 40 µM of complex + DNA; Lane 6: 50 µM of complex + DNA. Fig. 6. Agarose gel electrophoresis diagram showing cleavage of pBR322 supercoiled DNA (300 ng) by complex 1a (40 µM ) in presence of activators and reactive oxygen species at 310 K after incubation for 45 min. Lane 1: DNA control; Lane 2: DNA + 1a + H2O2 (0.4 M); Lane 3: DNA+ 1a + MPA (0.4 M); Lane 4: DNA+ 1a + Asc (0.4 M); Lane 5 : DNA + 1a + GSH (0.4 M); Lane 6 : DNA + 1a + DMSO (0.4 M); Lane 7 : DNA+ 1a + ethyl alcohol (0.4 M); Lane 8 : DNA+ 1a + sodium azide (0.4 M); Lane 9 : DNA+ 1a + SOD (15 units); Lane 10 : DNA+ 1a + DAPI (8 µM); Lane 11 : DNA+ 1a + methyl green (2.5 µL of a 0.01mg/ml solution). Fig. 7. Agarose gel electrophoresis diagram showing cleavage of pBR322 supercoiled DNA (300 ng) by complex 2a (40 µM ) in presence of activators and reactive oxygen species at 310 K after incubation for 45 min. Lane 1: DNA control; Lane 2: DNA + 2a + DMSO (0.4 M); Lane 3 : DNA+ 2a + ethyl alcohol (0.4 M); Lane 4 : DNA+ 2a + sodium azide (0.4 M); Lane 5 : DNA+ 2a + SOD (15 units); Lane 6 : DNA+ 2a + DAPI (8 µM); Lane 7 : DNA+ 2a + methyl green (2.5 µL of a 0.01mg/ml solution). Fig. 8. MTT assay of complex (a) 1a (b) 1b. Graph showing effect of concentration on viability of HeLa cells. More than 50% cells were viable at 15 µM, at concentration ≥ 20 µM viability was progressively compromised. * represent control set without complexes 1a and 1b, ** absorbance corresponds to number of cells.

29

Fig. 9. Confocal microscopy images of complexes (a) 1a and (a′) 1b localization in HeLa cells. Nucleus was stained with propidium iodide (b, b'). c and c' represent the Differential Interference Contrast (DIC) image of 1a and 1b treated HeLa cells respectively. Both complexes 1a and 1b were localized to nucleus as depicted from similar localization with propidium iodide (d and d' respectively). Fig. 10. Molecular docked structures of L−enantiomeric complexes (a) 1a and (b) 2a fitted into GC base pairs of minor groove inside the DNA dodecamer duplex of sequence d(CGCGAATTCGCG)2 (PDB ID: 1BNA).

Figures

Fig. 1

30

Fig. 2

31

Fig. 3

32

Fig. 4

Fig. 5

Fig. 6

33

Fig. 7

Fig. 8

34

Fig. 9

Fig. 10 (a)

35

Fig. 10 (b)

36

Scheme I: Synthetic route to complexes 1& 2 (a and b).

37

Graphical Abstract (Pictogram) MTT assay and molecular docked model of enantiomeric L–fluoro benzothiazole Schiff base–valine Cu(II) complex, 1 (a).

38

Research Highlights
New enantiomeric Cu(II)/Zn(II) complexes, 1 & 2 (a and b) derieved from flurobenzothiazole Schiff base ligand were synthesized.
Preliminary in vitro DNA binding studies with CT DNA revealed highest DNA binding propensity of L-enantiomeric Cu(II) complex 1a.
Complex 1a and b cleaved pBR322 DNA via hydrolytic pathway with significantly good activity shown by L–enantiomeric complex.
Cytotoxic results of complexes 1a and b against HeLa cancer line implicated that more than 50 % cells were viable at 15 µM.

39