investigation of diorganotin(iv) complexes: synthesis, characterization, in vitro dna binding...

Inorganica Chimica Acta xxx (2014) xxx–xxx

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Inorganica Chimica Acta

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Investigation of diorganotin(IV) complexes: Synthesis, characterization,in vitro DNA binding studies and cytotoxicity assessmentof di-n-butyltin(IV) complex

http://dx.doi.org/10.1016/j.ica.2014.07.0560020-1693/� 2014 Elsevier B.V. All rights reserved.

⇑ Corresponding author. Tel.: +91 9358255791.E-mail address: [email protected] (S. Tabassum).

Please cite this article in press as: S. Tabassum, S. Yadav, Inorg. Chim. Acta (2014), http://dx.doi.org/10.1016/j.ica.2014.07.056

Sartaj Tabassum ⇑, Shipra YadavDepartment of Chemistry, Aligarh Muslim University, Aligarh 202002, India

a r t i c l e i n f o

Article history:Received 8 May 2014Received in revised form 24 July 2014Accepted 25 July 2014Available online xxxx

Keywords:Diorganotin(IV) complexesIn vitro DNA bindingMolecular dockingSRB assay

a b s t r a c t

Diorganotin(IV) complexes of general formula R2SnL (R = Me, 1; Bu, 2; Ph, 3) with tridentate ONO donorSchiff base ligand were synthesized and structurally characterized by adopting various spectroscopic(IR, 1H 13C and 119Sn NMR, UV–Vis, ESI MS, XRD) and analytical techniques. In vitro DNA binding profileof 1–3 were carried out by various biophysical methods viz., UV–Vis titrations, fluorescence, circulardichroic and viscosity measurements which revealed the electrostatic mode of interaction via phospho-diester backbone of DNA duplex. The intrinsic binding constant Kb values of L and 1–3 were found to be7.53 � 103, 2.98 � 104, 5.74 � 104 and 3.64 � 104 M�1, respectively suggesting the higher bindingpropensity of 2, di-n-butyltin(IV) complex as compared to 1 and 3. Further, the computer-aided molec-ular docking technique was carried out to validate and rationalize the observed binding affinities towardsthe molecular target DNA. The resulting binding energies of docked complexes were found to be �212.2,�317.8 and �286.0 KJ mol�1, respectively. In addition, complex 2 was found remarkably effective againstU373MG (CNS), PC3 (prostrate), Hop62 (lung), HL60 (leukemia), HCT15 (colon), SK-OV-3 (ovarian), HeLa(cervix) and MCF7 (breast) cancer cell lines with GI50 values <10 lg/ml.

� 2014 Elsevier B.V. All rights reserved.

1. Introduction

Cancer as a leading cause of death worldwide, accounting for7.6 million deaths in 2008 and anticipates �13 million moredeaths by 2030 [1]. Cancer chemotherapy is the primary treatmentmodality against cancer, which was initially fueled by the seren-dipitous discovery of antitumor drug- cisplatin, cis-diamminedi-chloroplatinum(II) in 1969 [2]. Potentially, cisplatin and itsanalogues exhibit a broad antineoplastic spectrum and in combi-nation with other chemotherapeutic agents, can be highly effectivein the treatment of many cancers [3,4]. Platinum drugs arebelieved to induce cytotoxicity by cross-linking DNA, causingchanges to the DNA structure that inhibit replication and proteinsynthesis which would ultimately lead to induction of apoptosis[5]. However, serious adverse effects, such as dose limiting, neph-rotoxicity, peripheral neuropathy and hearing loss, are well known[6]. These limitations have stimulated an extensive search forunconventional chemotherapeutic strategies. In this context, orga-nometallic compounds are the promising chemotherapeutics can-didates in cancer therapy, which offer ample opportunities in the

design of novel classes of medicinal compounds, potentially withnew metal-specific modes of action [7–10]. Interestingly, amongthe organometallics, organotin(IV) compounds are recognized asan effective alternative to platinum drugs, affording differentmechanisms of action than those of cisplatin and its analogues,thus leading to alternative therapeutic protocols showing advanta-ges both in terms of lower toxicity and platinum induced resis-tance [11–13]. In particular, a large number of organotinderivatives have been prepared and were tested in vitro andin vivo against different panels of human cell lines, which exhibitedremarkable antiproliferative properties [14]. In general, organo-tin(IV) moieties bind to glycoproteins or to cellular proteins, anddirectly interact with DNA, causing cell death by apoptotic mecha-nisms [15,16].

Ligands can significantly alter the biological properties by mod-ifying reactivity or substitution inertness. The role of tailoredligand framework introduced into the metal-based medicinalagents is of considerable importance in tuning the cytotoxiccharacteristics of the complex as it not only mutes the potentialtoxicity of metallodrugs but also significantly alters the reactivityof the metal ion [17,18]. In recent years, polydentate proligand2-amino-2-methyl-1,3-propanediol (ampdH2) have been a centerof attention and attraction for chemists and biologists because of

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their wide structural and pharmacological relevance [19–21]. Theligand with mixed N and O donor atoms like ampdH2 was selectedbecause its Schiff base derivatives are versatile ligands in terms ofdonor properties and so are able to form higher nuclearity com-pounds (Scheme 1).

DNA is the main intracellular target for many anticancer drugs[22] and metal complexes which can bind with specificity to DNAare of importance in the development of new anticancer agents.DNA binding is a major criterion for the designing of novel antican-cer agents as many molecules exert their anticancer activities bybinding with DNA, causing alteration in DNA replication andthereby inhibiting the tumor cells growth [23,24]. In fact, the cyto-toxic activity of metallodrugs has often been correlated to theirDNA-binding properties [25]. The phosphate group of the DNAsugar backbone acts as an anchoring site and the binding of thenitrogen of the DNA base are extremely effective; this often resultsin the stabilization of the tin center as an octahedral species [12].

In lieu of above and in continuation of our previous work [26],herein, we report the synthesis and structural characterization ofnew diorganotin(IV) complexes by using an amino polyalcohol asa ligand backbone. Furthermore, the interaction studies of thecomplexes 1–3 with CT DNA were explored by employing UVabsorption, emission and circular dichoric spectroscopic tech-niques. Specifically, the cytotoxicity of di-n-butyltin(IV) complex,2 were preliminarily investigated by SRB assay against the panelof human tumor cells, demonstrating that the complex could bea potent therapeutic agent for the treatment of cancer.

2. Experimental

2.1. Materials and instrumentation

Reagent grade chemicals were used without further purificationfor all syntheses and experiments. Acetylacetone (acac) (Merck),2-amino-2-methylpropane-1,3-diol (ampdH2), dimethyltin(IV)dichloride, di-n-butyltin(IV) dichloride, diphenyltin(IV) dichloride,triethylamine (Et3N), tris (hydroxymethyl)aminomethane or Tris(Sigma–Aldrich) and supercoiled plasmid pBR322 DNA (Genei)were utilized. Disodium salt of calf thymus DNA (CT DNA)purchased from Sigma Chem. Co. and was stored at 4 �C.

The 1H, 13C and 119Sn NMR spectra were obtained on a BrukerDRX-400 spectrometer operating at 400, 100 and 150 MHz, respec-tively. Molar conductance was measured at room temperature onEutech con 510 electronic conductivity bridge. Infrared spectrawere recorded on Interspec 2020 FTIR spectrometer in KBr pelletsfrom 400–4000 cm�1. Electrospray mass spectra were recorded onMicromass Quattro II triple quadrupol mass spectrometer. Microa-nalyses (C, H and N) were performed on a Elementar Vario EL III.XRD were recorded on Rikagu mini Flex II instrument. UV�Vis spec-tra were recorded at room temperature on a Perkin–Elmer Lambda25 spectrometer. Fluorescence measurements were made onShimadzu RF–5301PC Spectrofluorophotometer. The SEM imageswere recorded with JEOL JSM-6510LV Scanning Electron Micro-scope. CD spectra were measured by a Jasco-J-815 spectrometer

NH2

OH

OH

ampdH2

CH3

HO

OH

CH3

H3C N

OH

Imine nitrogen

Schiff base ligand

Scheme 1. Ligand used in the synthesis of diorganotins.

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using a quartz cell (0.1 cm) at 0.1 nm intervals, adjusting the bandwidth to 1.0 nm and the scan speed to 20 nm min�1. Viscositymeasurements were carried out from observed flow time of CTDNA containing solution (t > 100 s) corrected for the flow time ofbuffer alone (t0), using Ostwald’s viscometer at 25 ± 0.01 �C. Flowtime was measured with a digital stopwatch.

2.2. Synthesis

2.2.1. Synthesis of ligand (L)To a methanolic solution of ampdH2 (1.051 g, 10 mmol) the

stoichiometric amounts of acetylacetone (1.03 ml, 10 mmol) wasadded drop wise. The reaction mixture was stirred and refluxedfor about 6–7 h during which the color of the solution turned topale-yellow, the reaction was monitored by TLC till its completion.The solvent was removed by using a rotavapor which afforded theproduct in the form of an oily yellow viscous liquid followed byrepeated washings with petroleum ether.

2.2.2. Synthesis of organotin(IV) complexes R2SnL with R = Me (1), n-Bu (2), Ph (3)

Diorganotin(IV) complexes were synthesized by refluxingequivalent molar ratio of R2SnCl2 (2 mmol; R = Me, 0.438 g for 1;R = n-Bu, 0.606 g for 2; R = Ph, 0.686 g for 3) and ligand L(0.374 g, 2 mmol) in the presence of Et3N (0.276 ml, 2 mmol) indry methanol. The resulting solution was refluxed for 4–5 h andsubsequently concentrated to dryness under reduced pressure toget the final product. The product obtained was washed withhexane in order to remove impurities and dried in vacuo.

[Me2SnL], 1: Yield: 68%. M.p. = 220 �C. % Anal. Calc. forC11H21NO3Sn (334.0): C, 39.56; H, 6.34; N, 4.19. Found: C, 39.53;H, 6.26; N, 4.11%. Selected IR data (cm�1/m): 3617 m(O–H), 1640m(C@N), 687 m(Sn–C); 532 m(Sn–O); 442 m(Sn–N). Molar Conduc-tance, KM (10�3 M, DMSO): 21 X�1 cm2 mol�1 (non electrolyte).UV–Vis (DMSO, kmax, nm): 284. 1H NMR (DMSO–d6, 400 MHz) d(ppm); 5.08 (C�CH@C, s, 1H), 3.20 and 3.13 (�CH2, s, 4H), 2.9(CH2–OH, s, 1H), 2.25 (O–C�CH3, s, 3H), 1.82 (N@C–CH3, s, 3H),0.84 (Sn–CH3, s, 6H, 2J[119Sn–1H] = 76 Hz). 13C NMR (DMSO-d6,100 MHz) d (ppm): 190.2 (C@N), 170.0 (O–C@), 99.8 (@CH–),63.8 (CH2�OH), 58.4 (CH2�O), 49.4 (Cter), 27.8 (CH3�C@N), 18.5(Sn–CH3, 1J[119Sn–13C] = 652 Hz). 119Sn NMR (DMSO-d6, 146 MHz)d (ppm): –162.2. ESI–MS (m/z+): [C11H21NO3Sn]+ 334.0.

[n-Bu2SnL], 2: Yield: 70%. M.p. = 191 �C. % Anal. Calc. forC17H33NO3Sn (418.16): C, 48.83; H, 7.95; N, 3.35. Found: C,48.78; H, 7.87; N, 3.29%. Selected IR data (cm�1/m): 3640 m(O–H),1647 m(C@N), 679 m(Sn–C); 548 m(Sn–O); 439 m(Sn–N). MolarConductance, KM (10�3 M, DMSO): 30 X�1 cm2 mol�1 (non electro-lyte). UV–Vis (DMSO, kmax, nm): 283. 1H NMR (DMSO-d6, 400 MHz)d (ppm); 5.31 (C�CH@C, s, 1H), 3.43 and 3.34 (�CH2, s, 4H), 3.09(CH2–OH, s, 1H), 2.33 (O–C�CH3, s, 3H), 2.11 (N@C–CH3, s, 3H),2.06 (aCH2, t, 4H, 2J[119Sn–1H] = 75 Hz), 1.99–1.77 (bCH2, m, 4H),1.14 (cCH2, m, 4H), 1.06 (dCH3, t, 5J[119Sn–1H] = 12 Hz). 13C NMR(DMSO-d6, 100 MHz) d (ppm): 192.3 (C@N), 171.2 (O–C@), 97.4(@CH–), 64.5 (CH2�OH), 58.2 (CH2�O), 48.5 (Cter), 31.4–24.5(CH3�C@N + CH3�C@CH + aCH2), 23.3 (bCH2), 20.5 (cCH2), 17.4(dCH3). 119Sn NMR (DMSO–d6, 146 MHz) d (ppm): �171.9.ESI–MS (m/z+): [C17H33NO3Sn+H]+ 419.1, [C17H33NO3Sn–C4H9]+

361.4, [C17H33NO3Sn–2C4H9]+ 303.2.[Ph2SnL], 3: Yield: 76%. M.p. = 263 �C. % Anal. Calc. for C21H25

NO3Sn (458.14): C, 55.05; H, 5.50; N, 3.06. Found: C, 55.00; H,5.43; N, 3.01%. Selected IR data (cm�1/m): 3595 m(O–H), 1643m(C@N), 681 m(Sn–C); 550 m(Sn–O); 447 m(Sn–N). Molar Conduc-tance, KM (10�3 M, DMSO): 28 X�1cm2 mol�1 (non electrolyte).UV–Vis (DMSO, kmax, nm): 284. 1H NMR (DMSO-d6, 400 MHz) d(ppm); 7.77�7.01 (m, C6H5, 10H), 5.23 (C�CH@C, s, 1H), 3.24–3.01 (�CH2, s, 4H), 2.32 (CH2–OH, s, 1H), 2.19 (O–C�CH3, s, 3H),

14), http://dx.doi.org/10.1016/j.ica.2014.07.056


H3C OH

HO CH3NH2

CH3

O

HO+ CH3

HO

OHCH3

H3C NOH

CH3

HO

OHCH3

H3C NOH

CH3

O

OH

CH3

H3CN

OEt3N, SnR2Cl2

MeOH, 60 0C

Sn

R R

R = Me, 1= Bu, 2= Ph, 3

MeOH, 60 0C

L

L

2-Amino-2-methylpropane-1,3-diol

Acetylacetone

Scheme 2. Schematic representation of the formation of ligand and complexes 1–3.

CH3

HO

OH

CH3

H3C N

OH

Scheme 3. Transamination of acetylacetone.

S. Tabassum, S. Yadav / Inorganica Chimica Acta xxx (2014) xxx–xxx 3

1.86 (N@C–CH3, s, 3H). 13C NMR (DMSO-d6, 100 MHz) d (ppm):192.1 (C@N), 166.9 (O–C@), 126.6–136.1 (Sn–C6H5, 3J[119Sn–13C] =59 Hz), 101.2 (@CH–), 63.3 (CH2�OH), 48.6 (Cter), 27.6 (CH3�C@N).119Sn NMR (DMSO–d6, 146 MHz) d (ppm): �317.5. ESI–MS (m/z+):[C21H25NO3Sn+H]+ 459.1.

2.3. DNA binding

DNA binding experiments which include absorption spectraltitrations, emission spectroscopy, and circular dichroism spectralanalysis conformed to the standard methods and practices previ-ously adopted by our laboratory standard protocol [27–29].

2.4. Morphological and structural analysis

The samples for SEM analysis of the complex and its condensatewith CT DNA were prepared from the solution of the complex 2(prepared in DMSO) and equimolar mixture of 2 and pBR322(dissolved in 0.01 M Tris–HCl buffer pH 7.2). The samples wereair dried at room temperature for 24 h and the images wererecorded with JEOL JSM-6510LV Scanning Electron Microscope atan accelerating voltage between 10 and 15 kV. Before observationof SEM images, the samples were coated with platinum by JEOLJFC-1600 auto fine coater for 2 min at 20 mA.

2.5. Molecular docking

The molecular docking studies were performed by using HEX6.3 software [30], which is an interactive molecular graphics pro-gram for calculating and displaying feasible docking modes ofenzymes and DNA molecule. The structure of the complex wassketched by CHEMSKETCH (http://www.acdlabs.com) and con-verted to pdb format from mol format by AVOGADRO (http://www.vcclab.org/lab/babel/). The structure of the B-DNA dodecam-er d(CGCGAATTCGCG)2 (PDB ID: 1BNA) was downloaded from theprotein data bank (http://www.rcsb.org./pdb). Visualization of thedocked pose has been done by using CHIMERA and DiscoveryStudio molecular graphic programs.

2.6. In vitro antitumor studies

The cytotoxicity assessment was screened in terms of GI50

(concentration of drug that produces 50% inhibition of the cells),TGI (concentration of the drug that produces total inhibition of thecells) and LC50 (concentration of the drug that kills 50% of the cells)


values by the semi-automated sulforhodamineB (SRB) assay. Theresults in terms of GI50 values were compared to adriamycin. Thesehuman malignant cell lines were procured and grown in RPMI-1640medium supplemented with 10% fetal bovine serum and antibioticsto study growth pattern of these cells. The proliferation of the cellsupon treatment with chemotherapy was determined using the sul-forhodamine B (SRB) assay. Cells were seeded in 96 well plates at anappropriate cell density to give optical density in the linear range(from 0.5 to 1.8) and were incubated at 37 �C in CO2 incubator for24 h. Stock solutions of the complexes were prepared as 100 mg/ml in DMSO, and four dilutions, that is 10, 20, 40, and 80 ml, in trip-licates were tested; each well receiving 90 ml of cell suspension and10 ml of the drug solution. Appropriate positive control (Adriamy-cin) and vehicle controls were also run. The plates with cells wereincubated in CO2 incubator with 5% CO2 for 24 h followed by drugaddition. The plates were incubated further for 48 h. Terminationof experiment was carried out by gently layering the cells with50 ml of chilled 30% TCA (in case of adherent Q18 cells) and 50%TCA (in case of suspension cell lines) for cell fixation and kept at4 �C for 1 h. Plates were stained with 50 ml of 0.4 % SRB for 20 min.All experiments were made in triplicate.

3. Results and discussion

3.1. Synthesis and spectroscopic study

The Schiff base ligand L was synthesized by the condensationreaction of acetylacetone with 2-amino-2-methylpropane-1,3-diol(ampdH2) in 1:1 ratio. Subsequently, diorganotin(IV) analogues,1–3 were prepared using the methodology depicted in Scheme 2under anhydrous condition. The Schiff base was used in situ forthe synthesis of the diorganotin(IV) complexes. The acetylacetoneSchiff base ligand was selected due to its ability to exist inketo–enol tautomeric form, which prevents the formation of theligand in 1:2 stoichiometric ratio by restricting the transaminationreaction of the carbonyl group (Scheme 3). The formation of


http://www.acdlabs.com

http://www.vcclab.org/lab/babel/

http://www.vcclab.org/lab/babel/

http://www.rcsb.org./pdb



diorganotin(IV) analogues, 1–3 was confirmed by elemental analy-sis, molecular ion peak in ESI-MS spectra, characteristic absorptionbands in FT-IR, resonance signals in the 1H, 13C and 119Sn NMRspectra, which revealed trigonal bipyramidal geometry for the SnIV

metal ion. The molar conductance values of synthesized complexes1–3 in DMSO (10–3 M) at 25 �C were too low to account for anydissociative ions in the complexes, consistent with their non–electrolytic nature.

In complexes 1–3 the perturbation of m(C@N) absorption bandfrom 1655 cm�1 to lower frequencies indicated the involvementof N atom in coordination. Moreover, the absence of ketonic groupvibrations m(–C@O) at 1700 cm�1 confirmed the ligand coordina-tion to the SnIV atom in the enolic form rather than ketonic form[31]. Meanwhile, the strong characteristic envelope at 3640–3595 cm�1 was assigned to the vibrations of the unreacted OH moi-ety in the complexes. Additional stretching frequencies of interestwere those associated with m(Sn–N), m(Sn–O) and m(Sn–C) groupsconfirmed the coordination of ligand to the tin metal center [32].

The multinuclear (1H, 13C and 119Sn) NMR acquired in DMSO-d6

showed functional group signatures as expected and thus con-firmed the postulated structure of 1–3. In the 1H NMR spectrumof complex 1 the characteristic signals for the methyl protons oftin metal center appeared at 0.91–0.57 ppm [33]. Similarly, 2exhibited a multiple resonance due to –CH2–CH2–CH2– skeletonfor butyl protons in the range of 2.06–1.14 ppm and a triplet at1.06 ppm for terminal methyl groups [34]. While, the diphenyl-tin(IV) complex, 3 exhibited a characteristic aromatic signaturein the region of 7.77–7.01 ppm [35].

Further distinctive signals were found in the 13C NMR spectra.The resonance signals for the C@N and O–C@carbon atoms in thecomplexes were observed at ca. 192 and 170 ppm, respectively.Additionally, the characteristic chemical shifts of the Sn–CH3 andC(n–bu) carbon atoms observed at 20.6–18.5 and 24.5–17.4 ppm inthe spectra of complexes 1 and 2, indicative of the nature of sub-stituents at the Sn atom [33,34]. Moreover, a broad envelope at136.1�126.6 ppm confirmed the presence of aromatic skeletoncarbons in case of 3. The dihedral C–Sn–C angle of the diorgano-tin(IV) complexes 1–3 were calculated by applying Lockhart andManders equation, h (CSnC) = (0.0161) (2J)2 – (1.32) (2J) + 133.4,and were found to be 126.03�, 124.96� and 111.56� for , respec-tively which corresponds to the pentacoordinate geometry of Snatom.

Furthermore, the 119Sn NMR spectra of 1–3 displayed chemicalshifts at �162.2, �171.9 and �317.5 ppm, which further confirmedthe pentacoordinate geometry of Sn(IV) metal atoms.

The ESI-MS analyses allow the assignment of characteristicfragmentation peaks. The fragmentation scheme of 1 exhibitedpeak at m/z 334.0 assigned to [C11H21NO3Sn] fragment. The massspectrum of 2 revealed peaks at m/z 419.1, 361.4 and 303.2 for[C17H33NO3Sn+H]+ [C17H33NO3Sn–C4H9] and [C17H33NO3Sn–2C4H9] fragments, respectively. Similarly, 3 exhibited a fragmenta-tion peak at m/z 459.1 which could be ascribed to [C15H21

NO2Sn+H]+. Moreover, the presence of the tin atom is easilydetected due to the isotopic abundances of tin.

The electronic absorption spectra of the ligand, L and complexes1–3 (in DMSO at room temperature) displayed broad band at 271and ca. 284 nm, respectively ascribed to n ? p⁄ intraligand transi-tions of the azomethine chromophore. Indeed, the evident fre-quency shifts of n ? p⁄ transitions in the complexes suggestedthe coordination of azomethine nitrogen atom to the tin metalcenter.

To obtain further evidence about the structure of the organotin(IV) compounds, X-ray powder diffraction studies of the complexes1–3 were performed as it was difficult to isolate single crystalssuitable for single crystal X-ray crystallography. The XRPD patternobtained for the organotin(IV) compounds revealed well defined


crystalline peaks indicating the microcrystalline nature. The X-ray diffractogram of the complex 1 exhibited reflecting peaks atscattering angles (2h) of 10.86, 14.91, 18.96, 24.27, 27.91 and35.64 assigned to (021), (411), (202), (022), (443), (113) and(551) crystal planes, respectively. While, complex 2 revealedreflecting peaks at 2h of 16.49, 22.19, 24.42, 25.07, 37.66 and44.13 corresponding to reflection from (401), (131), (432), (402),(110) and (541) crystal planes and similarly in case of 3, 2h scatter-ing angles 7.81, 14.19, 16.42, 18.27, 22.32, 24.61, 37.13 and 43.86were assigned to (400), (202), (110), (331), (432), (402), (111)and (544) crystal planes, respectively. All complexes exhibitedcharacteristic of a well ordered hexagonal crystal system withspace group P.

3.2. DNA binding studies

The interactions of metal complexes with DNA have been thesubject of interest for the development of effective chemothera-peutic agents. DNA offers several potential binding sites for metalcomplexes, including the anionic phosphate backbone, electron-rich bases, and the major or minor grooves [36]. Therefore, thebinding mode and affinity involved between the complexes 1–3and CT DNA was investigated by UV–Vis absorption, fluorescence,CD spectroscopic studies and viscosity in Tris–HCl buffer.

3.2.1. Absorption spectral titrationsThe binding propensity of the ligand, L and diorganotins, 1–3 for

CT DNA were evaluated by electronic absorption spectroscopy bystudying DNA-complex interactions. The ligand L exhibited charac-teristic absorption peak at 271 nm while the complexes 1–3revealed absorption bands at ca. 284 nm ascribed to intra-ligandn ? p⁄ transitions. The electronic absorption spectral changes ofthe ligand, L and its complexes 1–3 in absence and presence ofCT DNA are illustrated in Fig. 1. Addition of increasing concentra-tions of CT DNA (0–3.80 � 10�4 M) to a fixed concentration of Land 1–3 (0.20 � 10�4 M) exhibited hyperchromism of 27.8%,51.3%, 64.7% and 57.4%, respectively with a blue shift of 3 nm incase of L and ca. 8 nm shifts in complexes. The observationssuggested that the complexes interact electrostatically with thephosphodiester backbone of DNA duplex. Lewis acidic Sn(IV)cations in the trigonal bipyramidal geometry exhibit preferentialselectivity toward the polyanionic phosphate backbone of DNA,and thus cause the contraction and conformational changes inDNA helix [37].

In order to compare quantitatively the binding strength of thecomplexes with CT DNA quantitatively, the intrinsic bindingconstants (Kb) were calculated by monitoring the changes in theintraligand band at corresponding kmax with increasing concentra-tion of CT DNA and is given by the ratio of slope to the intercept inplots from Wolfe–Shimer Eq. (1), through a plot of [DNA]/ea – ef

versus [DNA]:

½DNA�ea � ef

¼ ½DNA�eb � ef

þ 1Kbðea � efÞ

ð1Þ

where ea, ef and eb correspond to Aobsd/[M], the extinction coeffi-cient for free metal complex and the extinction coefficient for themetal complex in the fully bound form, respectively. In a plot of[DNA]/(ea – ef) versus [DNA], Kb is given by the ratio of the slopeto the intercept. The intrinsic binding constant Kb values obtainedfor L and complexes 1–3 were found to be 7.53 � 103, 2.98 � 104,5.74 � 104 and 3.64 � 104 M�1, respectively with standard devia-tion of ±0.02. The complexes 1–3 exhibited greater binding affinityfor DNA as compared to free ligand due to the presence of SnIV

metal center which could be attributed to its strong tendency forphosphate binding. Interestingly, di-n-butyltin, 2 exhibited the



Fig. 1. Absorption spectra of (a) L and complex (b) 1, (c) 2 and (d) 3 in the presence of increasing amount of CT DNA. Inset: Plots of [DNA]/(ea–ef) vs [DNA] for the titration ofCT DNA with complexes. [Complex] = 0.20 � 10�4 M, [CT DNA] = 0–3.80 � 10�4 M.


higher DNA binding affinity than their analogues which could beattributed to the additional hydrophobic interaction of the butylgroups with DNA bases which further provided the evidence thatthe type of group bound to central tin scaffold can modulate orenhance the binding affinity of the metal complex towards DNA.

3.2.2. Fluorescence spectroscopic studiesOn addition of increasing concentration of CT DNA (0–

1.20 � 10�5 M) to the fixed amounts of complexes 1–3(0.20 � 10�5 M), the significant enhancement in the emissionintensity centered at �365 nm were observed with no apparentchange in the shape and position of the emission bands (Fig. 2).The enhancement of the emission intensity indicated that the com-plexes bind with DNA and gets into a hydrophobic environmentinside the DNA which restrict the complex mobility at the bindingsite due to the inaccessibility of the solvent water molecules toreach into the hydrophobic environment inside the DNA helixand thus avoiding the quenching of the solvent molecules [38].The Scatchard binding constant, K of 1–3 were calculated fromthe plots of r/cf versus r, and were found to be 3.18 � 104,6.70 � 104 and 4.25 � 104 M�1, respectively.

3.2.3. Effect of K4[Fe(CN)6]Fluorescence quenching refers to any process, which is a

decrease of the fluorescence intensity from a fluorophore due toa variety of molecular interaction. These include excited-state


reactions, molecular rearrangements, energy transfer ground-statecomplex formation and collisional quenching. The steady-stateemission quenching experiment of diorganotin(IV) analogues,1–3 using [Fe(CN)6]4� as anionic quencher was performed to affordfurther binding information. [Fe(CN)6]4� is a dynamic fluorescencequencher and provides a sensitive tool to examine nature of theinteraction of probe with DNA [39]. The quenching efficiency for1–3 was evaluated by the classical Stern–Volmer constant Ksv,which varies with the experimental conditions. According to theStern–Volmer equation [40]:

I0=I ¼ 1þ Ksv:r ð2Þ

where I0 and I represent the fluorescence intensities in the absenceand presence of the compound, respectively; r is the concentrationratio of the complex to DNA and Ksv is the quenching constantobtained as the slope of I0/I versus r. The emission intensity of com-plexes in the absence of DNA was lower than that in the presence ofCT DNA under the same concentration of K4[Fe(CN)6] thereby,reflecting that complexes were protected by the DNA helix. Stern–Volmer plots of diorganotins, 1–3 in the absence of CT DNA gavethe value of Ksv of 1.6 � 104, 3.76 � 104 and 2.58 � 104 M�1, respec-tively while, in the presence of DNA, however, the slope wasremarkably decreased to 1.06 � 104, 1.34 � 104 and 1.2 � 104 M�1,respectively (Fig. 3). The greater decrease of the Ksv value for 2was attributed to the strong binding of the complex with DNAdouble helix.



Fig. 2. Emission spectra of complex (a) 1, (b) 2 and (c) 3 in Tris–HCl buffer (pH 7.2) in the absence and presence of CT DNA. [Complex] = 0.20 � 10�5 M,[DNA] = 0–1.20 � 10�5 M. Arrows indicate the change in emission intensity upon increasing the DNA concentration.


3.2.4. Effect of ionic strengthThe effect of ionic strength of strong electrolyte NaCl has been

studied on complex-DNA system to evaluate the electrostatic con-tribution into the DNA-binding event of the diorganotin(IV) ana-logues, 1–3. Increased ionic strength of the medium screens theelectrostatic repulsion between consecutive phosphate groups(negatively charged), prompting the helix to shrink due to reduc-tion in the unwinding tendency caused by the aforementionedelectrostatic repulsion. The fluorescence intensity of DNA boundcomplexes was significantly quenched with increasing ionicstrength through added NaCl (0–0.8 � 10�4 M). Due to the com-petitive interaction for phosphate anions, the addition of NaClweakens the surface binding interactions as well as hydrogenbonding between the CT DNA and the interacting molecules[41]. Therefore, the results implicate that the complexes 1–3 pre-dominantly binds to DNA phosphate backbone by electrostaticinteractions.

3.2.5. Circular dichroismCircular dichroism is a spectroscopic technique widely used to

study the binding mode and interaction affinity of small molecules,like metal compounds, with biomolecules, particularly with DNA[42]. Modifications of the CD signals in the spectral range of200–300 nm have shown to be useful to detect and follow DNA


conformational changes, damage and/or cleavage upon interactionwith metal complexes. The CD spectrum of CT DNA exhibits a posi-tive band at 275 nm due to base stacking and a negative band at245 nm due to right-handed helicity of B–DNA. Simple groovebinding and electrostatic interaction of the complexes with DNAshowed less or no perturbations on the base stacking and helicitybands, while intercalation can stabilize the helix conformation ofB–DNA, and enhances the intensities of both the CD bands [43].The change in conformation of DNA from B ? A or B ? Z showsincrease in intensities of the bands along with remarkable shiftsin the wavelength. Upon increasing the concentration of the com-plexes, the intensities of both the negative and positive bands of CTDNA decreased without any shift in the band position (Fig. 4),which is a clear indication of the interactions between the com-plexes and CT DNA. Such results suggested that the complexes1–3 induce distortion of the secondary structure of B-DNA withoutthe alteration in its conformation, probably via electrostatic modeof interactions. Furthermore, the changing intensity follows thetendency of 2 > 3 > 1, in line with the results obtained by UV andfluorescence spectroscopy.

3.2.6. Viscometric measurementsWith the aim of clarifying the binding mode of the diorganotins,

1–3 to DNA, viscosity measurements were carried out on CT DNA



0.0 0.2 0.4 0.6 0.80.0

0.5

1.0

1.5

2.0

2.5

3.0

[Fe(CN)6]-4 X 10-4 M-1

Ι/Ι0

0.0 0.2 0.4 0.6 0.80.0

0.5

1.0

1.5

2.0

2.5

3.0

[Fe(CN)6]-4 X 10-4 M-1

Ι/Ι0

(a) (b)

0.0 0.2 0.4 0.6 0.80.0

0.5

1.0

1.5

2.0

2.5

3.0

[Fe(CN)6]-4 X 10-4 M-1

Ι/Ι0

(c)Fig. 3. Emission quenching curves of complex (a) 1, (b) 2 and (c) 3 in the absence of CT DNA (w) and in the presence of CT DNA (d).

Fig. 4. CD spectra of CT DNA alone (a); in the presence of complex 1, (b); 2, (c) and3, (d) in Tris–HCl buffer at 25 �C. [Complex] = 10�4 M, [DNA] = 10�4 M.

0.0 0.2 0.4 0.6 0.8 1.00.0

0.2

0.4

0.6

0.8

1.0

1/R = [Complex]/[DNA]

η/η 0

Complex 2 Complex 3 Complex 1

Fig. 5. Effect of increasing amount of complexes 1–3 on the relative viscosity of CTDNA. [DNA] = 1.20 � 10�4 M, [Complex] = 0–1.0 � 10�4 M.


by varying the concentration of the complexes added. A classicalintercalative molecular interaction causes a significant increasein viscosity of the DNA solution due to the increase in separationof the base pairs at the intercalation sites and hence an increasein the overall DNA length. Whereas, complexes bound to DNA


through groove binding do not alter the relative viscosity of DNA,and complexes electrostatically bound to DNA will bend or kinkthe DNA helix, reducing its effective length and its viscosity,concomitantly [44]. The values of relative viscosity (g/go), whereg and go are the specific viscosities of DNA in the presence andabsence of the complexes, were determined and plotted againstvalues of 1/R (R = [DNA]/[complex]) (Fig. 5). For complexes 1–3,



Fig. 6. SEM images of (a and b) complex 2 and (c and d) condensation of CT DNA incubated in the presence of complex 2 after 24 h.


viscosity of CT DNA decreases with increase in the ratio of com-plexes to CT DNA, due to the interaction of complexes with DNAproducing kinks or bends in the DNA strand, thereby diminishingits effective length suggesting non-intercalative electrostaticbinding mode of complexes with DNA [45].

3.2.7. Scanning electron microscopy (SEM) analysisThe morphological changes in supercoiled pBR322 DNA upon

condensation with complex 2 was observed visually by using byusing scanning electron microscopy (SEM). The representativeSEM image of free complex 2 exhibited rectangular crystallinemorphology of di-n-butyltin(IV) complex (Fig. 6a and b). However,the complex-DNA condensate revealed the morphological transi-tion and formation of different structural features indicating thecondensation of DNA molecules into compact and massivestructures (Fig. 6c and d). Therefore, the differences observed inthe morphology of complex upon interaction with DNA areattributed to the conformational changes in secondary structureof DNA via electrostatic mode of interaction [46]. These morphol-ogy evidences are very coincident with the results obtained frombinding studies.

4. Molecular docking with DNA

The molecular docking technique is an attractive scaffold tounderstand the drug-DNA interactions for rational drug designand discovery, as well as to establish the mode of action of thedrug entity by placing it into the binding site of the target specificregion of the DNA. Computer-aided molecular docking studies ofdiorganotin(IV) analogues, 1–3 with DNA duplex of sequenced(CGCGAATTCGCG)2 dodecamer (PDB ID: 1BNA) were carriedout in order to give insight into the binding modes and interac-tion pattern of the synthesized complexes. The results exhibitedthat complexes, 1–3 interact with DNA via an electrostatic modeof interactions with the phosphate backbone of DNA or with theDNA bases. Moreover, –OH group of the complexes acts as strongH-bond donor or acceptor and was found to be engaged in


hydrogen-bonding interactions with DNA nucleobases availablein the minor grooves. The energetically most favorable conforma-tion of the docked pose revealed that 1–3 approaches toward thecurved contour of the targeted DNA in the minor groove withinA–T rich regions of the dodecamer, thus making favorable hydro-gen bonding and electrostatic interactions with DNA functionalgroups that define the stability of groove (Figs. 7 and 8) [47].AT rich regions are narrower and slightly deeper than CG richregions allowing for better ligand fit. Thus, many minor groovebinders were shown to prefer AT rich sequences. The resultingrelative binding energy of docked metal complexes 1–3 withDNA was found to be �212.2, �317.8 and �286.0 KJ mol�1,respectively. The results provided convincible proofs for electro-static interactions between DNA and diorganotin entities andexhibited more potent binding between the DNA and complex 2in comparison to other two complexes. This correlates well withthe experimental DNA binding studies. Thus, the spectroscopicexperimental results are harmonized with the molecular dockingstudy as well.

5. In vitro cytotoxicity assay

Di-n-butyltin(IV) complex, 2 was selected for in vitro antitu-mor activity because it exhibited higher binding propensity ascompared to 1 and 3 in preliminary DNA binding studies. Thein vitro cytotoxicity of 2 was analyzed by SRB assay on a panelof human cancer cell lines of different histological origin viz.,U373MG (CNS), PC3 (prostrate), Hop62 (lung), HL60 (leukemia),HCT15 (colon), SK-OV-3 (ovarian), HeLa (cervix) and MCF7(human breast) [48]. From the anticancer screening data pre-sented in Table 1 it is obvious that the complex showedremarkable cytotoxicity against all cancer cell lines with pro-nounced GI50 values <10 lg/ml and thereby, also validates itspotential to act as promising chemotherapeutic antitumor drugcandidate (Fig. 9). According to Huber et al. the high cytotoxicactivity of complexes are characterized by (i) the availabilityof coordination positions at Sn and (ii) the occurrence of



Fig. 8. Molecular docked model of complex 2 with DNA dodecamer duplex of sequence d(CGCGAATTCGCG)2 (PDB ID:1BNA).

Fig. 7. Molecular docked model of complex (a) 1, and (b) 3 with DNA dodecamer duplex of sequence d(CGCGAATTCGCG)2 (PDB ID:1BNA).


relatively stable ligand-Sn bonds, e.g. Sn–N and Sn–S, and whichresult in slow hydrolytic decomposition [49]. In agreementwith that, the significant anticancer activity evaluated for 2could be attributed to the availability of coordination position


around Sn(IV) ion of the five coordinated tin atom. Moreover,the lipophilicity due to the presence of number of carbonatoms in the organotin moiety of 2 warranted its impressivecytotoxicity.



Fig. 9. Growth curve showing % control growth verses drug concentration (lg/ml) against different human carcinoma cell lines: U373MG (CNS), PC3 (prostrate), Hop62(lung), HL60 (leukemia), HCT15 (colon), SK-OV-3 (ovarian), HeLa (cervix) and MCF7 (human breast).

Table 1Summary of the screening data of complex 2 for the in vitro cytotoxicity (lM).

Cell line U373MG PC3 Hop62 HL60 HCT15 SK–OV–3 HeLa MCF7

GI50 complex 2 16.5 <10 <10 <10 <10 <10 <10 <10ADR 53.0 <10 <10 <10 <10 <10 <10 <10

TGI complex 2 >80 <10 <10 19.5 14.0 <10 10.2 <10ADR >80 12.7 21.9 31.9 40.0 44.4 12.8 22.6

LC50 complex 2 >80 44.5 31.4 59.9 48.7 36.7 46.6 45.7ADR >80 52.8 53.3 75.5 >80 >80 53.4 56.4

Where, GI50 = concentration of drug that produces 50% inhibition of the cells, TGI = concentration of the drug that produces total inhibition of the cells, LC50 = concentration ofthe drug that kills 50% of the cells values and ADR = adriamycin.


6. Conclusions

Aiming at the development of promising antitumor agents, thediorganotin(IV) analogues with tridentate ONO donor Schiff baseligand were synthesized and structurally characterized. The com-parative in vitro DNA binding studies of 1–3 was investigated byabsorption, fluorescence, viscosity and CD measurements and theresults revealed efficient binding of the complexes via electrostaticbinding mode to the DNA double helix. The binding affinity ofthese complexes with CT DNA followed the order 2 > 3 > 1, display-ing a higher binding propensity of complex 2 as compared to othercomplexes. Additionally, computer-aided molecular docking stud-ies were substantiated well with the experimental DNA bindingresults and provided valuable information about the mode of inter-action between complex and DNA. Interestingly, di-n-butyltin(IV)


complex, 2 displayed good cytotoxic study against different tumorcell lines with a potency significantly higher than the adriamycin.The present study highlights the importance of diorganotin(IV)complexes, which proven its worth to serve as potent antitumoragent by interacting with the biological target DNA at themolecular level.

Acknowledgments

The authors are grateful to SAIF, Panjab University, Chandigarh;and STIC, Cochin for providing ESI–MS, NMR and elemental analy-sis, respectively. The authors are grateful to ACTREC, Mumbai forcarrying out the cytotoxic studies. They also acknowledge financialsupport to the Department of Chemistry, AMU through UGCassisted DRS–SAP and DST PURSE Programme.




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