Oxidovanadium(v) complexes of aroylhydrazones incorporating heterocycles: synthesis, characterization and study of DNA binding, photo-induced DNA cleavage and cytotoxic activities

Article abstract of DOI:10.1039/c4ra14369h

Four neutral oxidovanadium(v) complexes VO₂L¹ (1), VO₂L² (2), VOL³(OEt) (3) and VOL⁴(OEt)EtOH (4) [where HL¹ = 2-thiophenoylhydrazone of 2-acetylpyridine, HL² = 2-amino benzoylhydrazone of 2-benzoyl pyridine, H₂L³ = isonicotinoylhydrazone of 2-hydroxy acetophenone, H₂L⁴ = 2-furoylhydrazone of 2-hydroxy-1-napthaldehyde] with a hydrazone scaffold containing either furan, thiophene and pyridine residues have been synthesised. All complexes were thoroughly characterized by various spectroscopic (IR, UV-Vis, NMR and ESI-MS) and single crystal X-ray diffraction techniques. Crystallography establishes five-coordinate geometries, distorted toward square pyramidal for each of 1 and 2, based on a tridentate-O,N,N coordinating anion and two oxido-O atoms. The dianion in 3 is tetradentate, coordinating one V atom as for 1 and 2, and bridging another via the pyridyl-N atom, and the N₂O₄ octahedral coordination geometry is completed by oxido- and ethanolate-O atoms. The result of the V-N bridging is a helical coordination polymer. An NO₅ octahedral geometry is found in 4 defined by a tridentate-O,N,O anion, as well as oxido-, ethanolate- and ethanol-O atoms. Biological studies reveal that 1-4 have DNA binding propensity and show these to interact with CT-DNA through the minor groove binding mode, with binding constants ranging from 10³ to 10⁵ M^-1. All complexes show good photo-induced cleavage of pUC19 supercoiled plasmid DNA with 3 showing the highest photo-induced DNA cleavage activity of ～65%. Additionally, 1-4 are cytotoxic against the human cervical cancer cell line (HeLa) with IC₅₀ values ranging from 10 to 20 μM.

Full text of DOI:10.1039/c4ra14369h

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DOI: 10.1039/C4RA14369H
Oxidovanadium(V) complexes of aroylhydrazones incorporating
heterocycles: Synthesis, characterization and study of DNA binding,
photo-induced DNA cleavage and cytotoxic activities
Subhashree P. Dash, Alok K. Panda, Sagarika Pasayat, Rupam Dinda, Ashis Biswas, Edward R. T.
Tiekink, Subhadip Mukhopadhyay, Sujit K. Bhutia, Werner Kaminsky and Ekkehard Sinn
Synthesis, characterization and study of DNA binding, photo-induced DNA cleavage and cytotoxic activities of four neutral
1
oxidovanadium(V) complexes VO L (1), VO L (2), VOL (OEt) (3) and VOL (OEt)EtOH (4) with hydrazone scaffold have
2
3
4
2
2
been reported.

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Oxidovanadium(V) complexes of aroylhydrazones incorporating
heterocycles: Synthesis, characterization and study of DNA binding,
photoꢀinduced DNA cleavage and cytotoxic activities
a
b
a
a
b
Subhashree P. Dash, Alok K. Panda, Sagarika Pasayat, Rupam Dinda,* Ashis Biswas, Edward R. T.
c
d
d
e
f
Tiekink, Subhadip Mukhopadhyay, Sujit K. Bhutia, Werner Kaminsky and Ekkehard Sinn
a
Department of Chemistry, National Institute of Technology, Rourkela 769008, Odisha, India.
b
School of Basic Sciences, Indian Institute of Technology Bhubaneswar, Bhubaneswar 751 013, Odisha, India.
c
Department of Chemistry, University of Malaya, 50603 Kuala Lumpur, Malaysia.
d
Department of Life Science, National Institute of Technology, Rourkela 769008, Odisha, India.
e
Department of Chemistry, University of Washington, Box 351700, Seattle, WA 98195, USA.
f
Department of Chemistry, Western Michigan University, Kalamazoo, MI 49008, USA.
1

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____________________________________________________________________________
Abstract
Four neutral oxidovanadium(V) complexes VO L (1), VO L (2), VOL (OEt) (3) and VOL (OEt)EtOH (4) [where HL = 2ꢀ
1
2
3
4
1
2
2
2
3
thiophenoylhydrazone of 2ꢀacetylpyridine, HL = 2ꢀamino benzoylhydrazone of 2ꢀbenzoyl pyridine, H L =
2
4
isonicotinoylhydrazone of 2ꢀhydroxy acetophenone, H L = 2ꢀfuroylhydrazone of 2ꢀhydroxyꢀ1ꢀnapthaldehyde] with a hydrazone
2
scaffold containing either furan, thiophene and pyridine residues have been synthesised. All complexes were thoroughly
characterized by various spectroscopic (IR, UVꢀVis, NMR and ESIꢀMS) and single crystal Xꢀray diffraction techniques.
Crystallography establishes fiveꢀcoordinate geometries, distorted toward square pyramidal for each of 1 and 2, based on a
tridentateꢀO,N,N coordinating anion and two oxidoꢀO atoms. The dianion in 3 is tetradentate, coordinating one V atom as for 1
and 2, and bridging another via the pyridylꢀN atom, and the N O octahedral coordination geometry is completed by oxidoꢀ and
2
4
ethanolateꢀO atoms. The result of the V–N bridging is a helical coordination polymer. An NO octahedral geometry is found in 4
5
defined by a tridentateꢀO,N,O anion, as well as oxidoꢀ, ethanolateꢀ and ethanolꢀO atoms. Biological studies reveal that 1–4 have
DNA binding propensity and show these to interact with CTꢀDNA through minor groove binding mode, with binding constants
3
5
ꢀ1
ranging from 10 −10 M . All complexes show good photoꢀinduced cleavage of pUC19 supercoiled plasmid DNA with 3
showing the highest photoꢀinduced DNA cleavage activity of ~ 65%. Additionally, 1–4 are cytotoxic against the human cervical
cancer cell line (HeLa) with IC50 values ranging from 10 to 20 ꢀM.
______________________________________________________________________________
2

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1. Introduction
Metal complexes bearing a heterocyclic moiety open a new era of pharmaceutical research since their presence can
1–4
moderate physicochemical properties as well as can enhance the medicinal properties. Isoxazoles, pyrroles, and pyrazoles are
wellꢀknown examples of heteroaromatic organic compounds associated with diverse biological properties. Hence, it is not
surprising there are several reports in the literature where metal complexes have had their physicochemical and pharmacological
5–7
8
properties influenced by the presence of biologically active heterocycles containing nitrogen. In this context, thiophene,
9
10
furan, and pyridine moieties are especially regarded as desirable, due to their widespread use as a scaffold in medicinal
chemistry.
1
1
Metal coordination is one of the most efficient strategies in the design of repository, slowꢀrelease or longꢀacting drugs. It
12
was the discovery of cisplatin that gave momentum to metalꢀbased drug research. Despite its tremendous success as an antiꢀ
13–16
cancer drug, its use is restricted due to serious sideꢀeffects, general toxicity, and acquired drug resistance.
Therefore,
considerable effort is being made to synthesise new metalꢀbased drugs that effectively and specifically target cancer cells while
minimising toxic side effects. Among transition metals, vanadium complexes have attracted significant attention owing to their
17–20
21
22
diversified applications as models for the biological functions of vanadium,
such as haloperoxidation, phosphorylation,
2
3–26
27
28
insulin mimicking,
nitrogen fixation, tumour growth inhibition and prophylaxis against carcinogenesis. These complexes
29ꢀ32
have also numerous applications in DNA binding, cleavage and have shown the ability to act as antiꢀproliferative agents.
Hydrazones, –NH–N=CRR' (R and R′ = H, alkyl, aryl), are versatile ligands and have applications in the fields of
33
34
analytical and medicinal chemistry. Hydrazone moieties are important pharmacophoric cores of several antiꢀcancer, antiꢀ
35
inflammatory, antiꢀnociceptive and antiꢀplatelet drugs. Moreover, the electronic properties of hydrazone complexes are also
36
important in the design of complexes with better DNAꢀbinding and cleavage characteristics. Hydrazones in combination with
3
7–39
heterocycles display a broad range of biological activity.
heterocycles demonstrating medicinal properties are presented in Chart 1.
complexes having hydrazone ligands where a heterocyclic moiety enhances the pharmacological activity of these complexes.
However, to date, analogous vanadium complexes, and especially the study of their pharmacological activity, is still scarce and
Examples of hydrazide and hydrazone derivatives containing
40–44
There are several reports in the literature on metal
4
5
4
6
needs to be explored.
Considering the interesting pharmacological properties of ligand systems containing heterocycles and as a part of our studies
32,47
on design and synthesis of novel vanadium complexes with potential pharmaceutical activities,
we now focus our attention on
the synthesis of some new oxidovanadium complexes with a hydrazone scaffold containing either furan, thiophene and pyridine
residues. These show interesting DNA binding, photoꢀinduced DNA cleavage and cytotoxicity profiles. The interaction of the
complexes with calfꢀthymus DNA (CTꢀDNA) was also investigated using UVꢀVis absorption titration, thermal denaturation,
circular dicroism and fluorometric competitive binding studies. Their photoꢀcleavage reactions with pUC19 supercoiled plasmid
3

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DNA were investigated by gel electrophoresis. In addition, the cytotoxicity of the complexes against the cervical cancer cell line
HeLa) was assessed by the MTT assay.
(
4

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2. Experimental
2
.1 Materials and instrumentation
All chemicals were purchased from commercial sources and used without further purification. The VO(acac) complex was
2
4
8
prepared as described in the literature. MTT (3ꢀ[4,5ꢀDimethylthiazolꢀ2ꢀyl]ꢀ2,5ꢀdiphenyltetrazolium) and DAPI (4′,6ꢀdiamidinoꢀ
2
ꢀphenylindole dihydrochloride) were purchased from Sigma Aldrich (USA). Minimal essential medium (MEM) was purchased
from Gibco, India. Reagent grade solvents were dried and distilled prior to use. The supercoiled (SC) pUC19 DNA was purified
from E. coli cells with the aid of GeneJET Plasmid Isolation Kit (Thermo Scientific, USA). Biochemistry grade calf thymus (CT)
DNA was purchased from SRL (India). Molecular biology grade agarose was purchased from Sigma Aldrich (USA). Methyl
green and ethidium bromide were procured from HiMedia Laboratories. Elemental analyses were performed on a Vario ELcube
1
13
CHNS Elemental analyser and IR spectra were recorded on a PerkinꢀElmer Spectrum RXI spectrophotometer. H and C NMR
spectra were recorded with a Bruker Ultrashield 400 MHz spectrometer using SiMe as the internal standard. Electronic spectra
4
were recorded on a PerkinꢀElmer Lamda25 spectrophotometer. Cyclic voltammograms were recorded in CH Cl solutions,
2
2
containing 0.1 (M) TEAP (Tetraethyl ammonium perchlorate) as supporting electrolyte, using a CH1120A potentiostat, with
glassy carbon working electrode, Pt wire as counter electrode and Ag, AgCl/saturated KCl as reference electrode.
2
.2 Synthesis of the ligands
1
3–4
Schiff base ligands, HL and H L , were prepared by condensation of acid hydrazides (10 mmol) and the corresponding
2
47
2
carbonyl compound (10 mmol) in stirring ethanol (15 mL) for 3 h following a standard procedure. HL was prepared by
condensation of 2ꢀamino benzoylhydrazide with 2ꢀbenzoyl pyridine in refluxing methanol for 5 h. The resulting white compound
was filtered, washed with ethanol and dried over fused CaCl . The synthesised ligands are shown in Scheme 1.
2
1
2
.2.1 HL . Yield: 74%. Anal. Calc. for C H N OS: C, 58.76; H, 4.52; N, 17.13; S, 13.07. Found: C, 58.79; H, 4.48; N,
1
2
11
3
–
1
1
1
7.11; S, 13.05%. FTIR (KBR, ν/cm ): 3164 (N–H), 1647 (C=O), 1572 (C=N). H NMR (400 MHz, DMSOꢀd , ppm): δ 11.09
6
13
(
s, 1H, NH), 8.61–7.20 (m, 7H, aromatic), 2.45 (s, 3H, CH3). C NMR (100 MHz, DMSOꢀd , ppm): δ 162.55, 155.32, 150.79,
6
1
49.12, 137.10, 135.59, 133.33, 127.16, 125.23, 124.45, 121.25, 12.88.
2
2
.2.2 HL . Yield: 78%. Anal. Calc. for C H N O: C, 72.14; H, 5.10; N, 17.71. Found: C, 72.15; H, 5.11; N, 17.69%. FTIR
1
9
16
4
–1
1
(
KBR, ν/cm ): 3472 (NH )sym, 3354 (NH ) , 3048 (N–H), 1654 (C=O), 1576 (C=N). H NMR (400 MHz, DMSOꢀd , ppm): δ
2
2
a
s
y
m
6
1
3
1
1
1
4.50 (s, 1H, NH), 8.91–6.63 (m, 13H, aromatic), 6.56 (s, 2H, NH2). C NMR (100 MHz, DMSOꢀd , ppm): δ 157.07, 155.75,
6
53.94, 153.81, 152.27, 143.60, 142.78, 141.95, 138.00, 134.30, 134.12, 134.05, 133.64, 132.51, 131.38, 130.09, 122.24, 120.67,
17.91.
3
2
.2.3 H L . Yield: 69%. Anal. Calc. for C H N O : C, 65.87; H, 5.13; N, 16.46. Found: C, 65.88; H, 5.11; N, 16.50%.
2
14 13
3
2
–
1
1
FTIR (KBR, ν/cm ): 3368 (O–H), 3151 (N–H), 1678 (C=O), 1607 (C=N). H NMR (400 MHz, DMSOꢀd , ppm): δ 13.32 (s, 1H,
6
5

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13
OH), 11.72 (s, 1H, NH), 8.20–6.89 (m, 8H, aromatic), 2.42 (s, 3H, CH3). C NMR (100 MHz, DMSOꢀd , ppm): δ 163.40,
6
1
60.01, 159.21, 150.56, 140.84, 132.05, 129.19, 122.84, 122.63, 122.43, 119.68, 119.10, 117.82, 14.78.
4
2
.2.4 H L . Yield: 71%. Anal. Calc. for C H N O : C, 68.57; H, 4.32; N, 9.99. Found: C, 68.59; H, 4.34; N, 9.96%. FTIR
2
16 12
2
3
–1
1
(
KBR, ν/cm ): 3445 (O–H), 3172 (N–H), 1640 (C=O), 1603 (C=N). H NMR (400 MHz, DMSOꢀd , ppm): δ 12.72 (s, 1H, OH),
6
1
3
1
1
2.26 (s, 1H, NH), 9.51 (s, 1H, CH), 8.21–6.71 (m, 9H, aromatic). C NMR (100 MHz, DMSOꢀd , ppm): δ 158.43, 154.24,
6
47.54, 146.70, 146.50, 133.20, 132.10, 129.37, 128.26, 128.16, 123.96, 121.04, 119.29, 115.95, 112.75, 109.00.
2
.3 Synthesis of complexes 1–4
1
1
2
.3.1 VO L (1). To a hot solution of HL (1 mmol) in methanol (30 mL), VO(acac) (1 mmol) in DMF (5 mL) was added,
2 2
the colour changed instantly from colourless to greenꢀbrown. After 6 h of reflux, the solution was cooled, filtered and kept for
crystallisation. Slow evaporation of the filtrate over 4 days produced orange crystals. Yield: 63%. Anal. Calc. for C H N O SV:
1
2
10
3
3
–1
C, 44.05; H, 3.08; N, 12.84; S, 9.80. Found: C, 44.06; H, 3.05; N, 12.83; S, 9.81%. FTIR (KBR, ν/cm ): 1600 (C=N), 1255 (C–
–1
–1
1
O)enolic, 1036 (N–N), 946 (V=O). UVꢀVis in DMF [λ/nm (ε/M cm )]: 399 (14121), 277 (8371). H NMR (400 MHz, DMSOꢀd ,
6
13
ppm): δ 8.80–7.21 (m, 7H, aromatic), 2.71 (s, 3H, CH3). C NMR (100 MHz, DMSOꢀd , ppm): δ 171.54, 165.30, 159.69,
6
51
1
52.14, 148.11, 145.19, 144.33, 132.16, 127.23, 126.40, 124.22, 13.48. V NMR (DMSOꢀd ppm): δ –507.
6,
2
2
2
.3.2 VO L (2). To a hot solution of HL (1 mmol) in ethanol (30 mL), VO(acac) (1 mmol) was added, the colour changed
2 2
instantly from pale yellow to orange. After 5 h of reflux, the solution was cooled, filtered off and kept for crystallisation. Slow
evaporation of the filtrate over 4 days produced dark red crystals. Yield: 68%. Anal. Calc. for C H N O V: C, 57.30; H, 3.80;
19
15
4
3
–1
N, 14.07. Found: C, 57.31; H, 3.84; N, 14.08%. FTIR (KBR, ν/cm ): 3393 (NH )sym, 3284 (NH2)asym, 1594 (C=N), 1255 (C–
2
–1
–1
1
O)enolic, 1028 (N–N), 943 (V=O). UVꢀVis in DMF [λ/nm (ε/M cm )]: 460 (8945), 298 (11114), 258 (12770). H NMR (400
13
MHz, DMSOꢀd , ppm): δ 8.93–6.53 (m, 13H, aromatic), 6.73 (s, 2H, NH ). C NMR (100 MHz, DMSOꢀd , ppm): δ 178.68,
6
2
6
5
1
1
58.22, 155.08, 154.13, 150.71, 143.56, 134.04, 131.61, 130.99, 129.96, 129.32, 129.21, 127.12, 116.69, 115.45, 110.75.
V
NMR (DMSOꢀd , ppm): δ –512.
6
3
3
2
.3.3 VOL (OEt) (3). To the (1 mmol) sample of ligand H L in ethanol (20 mL), VO(acac) (1 mmol) was added under
2
2
refluxing conditions. After 3 h, the resulting deepꢀbrown solution was filtered and slow evaporation of the filtrate over 4–5 days
produced black crystals. The crystals were filtered and washed with ethanol. Yield: 64%. Anal. Calc. for C H N O V: C, 52.61;
16
16
3
4
–1
H, 4.42; N, 11.50. Found: C, 52.63; H, 4.44; N, 11.48%. FTIR (KBR, ν/cm ): 1601 (C=N), 1239 (C–O)enolic, 1037 (N–N), 964
–
1
–1
1
(
V=O). UVꢀVis in DMF [λ/nm (ε/M cm )]: 399 (7783), 326 (18601), 267 (22547). H NMR (400 MHz, DMSOꢀd , ppm): δ
6
13
8
.75–6.92 (m, 8H, aromatic), 5.59 (m, 2H, CH (OEt)), 1.54 (t, 3H, CH (OEt)), 2.93 (s, 3H, CH ). C NMR (100 MHz, DMSOꢀ
2 3 3
d6, ppm): δ 167.52, 164.69, 164.32, 161.80, 150.22, 140.15, 133.79, 130.69, 123.00, 122.51, 120.72, 117.00, 82.18, 56.50, 18.98,
51
1
7.32. V NMR (DMSOꢀd , ppm): δ –541.
6
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4
3
4
2
.3.4 VOL (OEt)EtOH (4). Complex 4 was prepared and crystallised similarly to 3, replacing H L by H L to yield (red)
2
2
crystals. Yield: 69%. Anal. Calc. for C H N O V: C, 55.05; H, 4.85; N, 6.42. Found: C, 55.02; H, 4.86; N, 6.38%. FTIR (KBR,
2
0
21
2
6
–
1
–1
–1
ν/cm ): 3567
(
O–H), 1612 (C=N), 1238 (C–O)enolic, 1052 (N–N), 998 (V=O). UVꢀVis in DMF [λ/nm (ε/M cm )]: 436 (7621),
1
3
5
26 (8168), 284 (11506), 260 (13128). H NMR (400 MHz, DMSOꢀd , ppm): δ 9.61 (s, 1H, CH), 8.40–6.65 (m, 9H, aromatic),
6
13
.67 (m, 2H, CH (OEt)), 3.44 (q, 2H, CH (EtOH)), 1.55 (t, 3H, CH (OEt)), 1.06 (t, 3H, CH (EtOH)). C NMR (100 MHz,
2
2
3
3
DMSOꢀd , ppm): δ 164.45, 163.24, 148.60, 146.19, 145.88, 135.40, 132.74, 129.24, 128.70, 128.61, 124.43, 121.75, 119.55,
6
51
1
15.27, 112.53, 111.50, 74.38, 56.61, 48.94, 18.57. V NMR (DMSOꢀd , ppm): δ –537.
6
2
.4 Xꢀray crystallography
Intensity data for 1, 2 and 4 were collected on a Bruker APEXII diffractometer equipped with a CCD area detector and graphiteꢀ
monochromated Mo Kα radiation (λ = 0.71069 Å), and those for 3 were measured on an Oxford Diffraction Xcalibur Ruby
Gemini system employing graphiteꢀmonochromated Cu Kα radiation (λ = 1.54178 Å). Data reduction and empirical absorption
49,50
correction, based on the multiꢀscan method, was by standard methods.
The structures were solved by direct methods using
51
52
SHELXS97 through the WinGX Interface and refinement (anisotropic displacement parameters, Cꢀbound hydrogen atoms in
2
2
2
2
2
c
idealised positions and a weighting scheme of the form w = 1/[σ (F ) + aP + bP] where P = (F + 2F )/3) of each structure
o
o
2
51
was carried out on F by fullꢀmatrix leastꢀsquares procedures ). The amineꢀH atoms in 2 were located from a difference Fourier
map and refined with the distance restraint N‒H = 0.88±0.01 Å. The thienyl ring in 1 was found to be disordered over two
coplanar dispositions of opposite orientation: the major component refined to a site occupancy factor of 0.836(2). For 1, the
ꢀ
3
maximum and minimum residual electron density peaks of 1.03 and 0.41 e Å , respectively, were located 0.78 and 0.51 Å,
respectively, from the V atom. The absolute structure of 3 was determined on the basis of 539 Friedel pairs included in the data
5
3
set; the value of the Flack parameter was 0.012(6). Crystal data and refinement details are given in Table 1. The molecular
52
structures shown in Figs 1–4, were drawn with 70% (35% for 2) displacement ellipsoids . The overlay diagrams shown in the
5
4
55
ESI were drawn with QMol, and the crystal packing diagrams with DIAMOND. Data interpretation was accomplished using
5
6
PLATON.
2
.5 DNA binding experiments
.5.1 Absorption spectral studies. Binding of the oxidovanadium(V) complexes to calf thymus (CT) DNA was carried out
2
4
7
as previously described. The UVꢀVis titration experiments were performed using a fixed concentration of metal complex (25
ꢀM) but variable CTꢀDNA concentrations ranging from 0 to 70 ꢀM in 10 mM Tris–HCl buffer (pH 8.0) containing 1% DMF.
7

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Binding of ligands to CTꢀDNA was also studied. For this, a fixed concentration of ligand [25 ꢀM in 10 mM Tris–HCl buffer (pH
.0) containing 1% DMF] was titrated with variable DNA concentration ranging from 0 to 350 ꢀM.
8
2
.5.2 Thermal denaturation studies. Thermal denaturation studies of CTꢀDNA (160 ꢀM) in the absence and presence of
o
complexes (150 ꢀM) were carried out by monitoring the absorbance at 260 nm in the temperature range of 30–90 C with a ramp
o
rate of 0.5 C/min in 10 mM TrisꢀHCl buffer (pH 8.0) containing 1% DMF. The experiments were carried out using a Chirascan
CD spectropolarimeter (Applied Photophysics, UK) in absorbance mode equipped with a peltier temperature controller. The
57
melting temperature (T ) was determined from the derivative plot (dA260/dT vs T) of the melting profile.
m
2
.5.3 Circular dichroism studies. Circular Dichroism (CD) spectroscopy was studied using Chirascan CD
o
spectropolarimeter (Applied Photophysics, UK) at 25 C. CD spectra of CTꢀDNA (100 ꢀM) in the absence and presence of
complexes (100 ꢀM) were obtained in the wavelength range of 240–400 nm in 10 mM TrisꢀHCl buffer (pH 8.0) containing 1%
4
7
DMF using quartz cell with 10 mm path length.
2
2
.5.4 Competitive DNA binding by fluorescence measurements
.5.4.1 4',6ꢀdiamidinoꢀ2ꢀphenylindole (DAPI) displacement experiment. The DNA binding probe, 4',6ꢀdiamidinoꢀ2ꢀ
58
phenylindole (DAPI), binds to the minor groove of the DNA. The competitive binding of the complexes 1–4 to the minor
groove of CTꢀDNA was studied by measuring the fluorescence of DAPI (2 ꢁM) bound CT−DNA (50 ꢁM) with increasing
concentration of the complexes. The fluorescence intensities of DAPI at 455 nm (excitation 358 nm) with an increasing amount
of the complex concentration (0−90 ꢁM) was measured with the aid of Fluoromax 4P spectrofluorimeter (Horiba Jobin Mayer,
USA).
2
.5.4.2 Methyl Green (MG) displacement experiment. The DNA binding probe, methyl green (MG), binds to the major
5
9
groove of the DNA. The competitive binding of the complex 1–4 to the major groove of CTꢀDNA was studied by measuring
the fluorescence of MG (2 ꢁM) bound CT−DNA (50 ꢁM) with increasing concentration of the complexes. The fluorescence
intensities of MG at 672 nm (excitation 633 nm) with an increasing amount of the complex concentration (0−90 ꢁM) was
measured with the aid of the same spectrofluorimeter as used above.
2
.5.4.3 Ethidium bromide (EB) displacement experiment. The DNA binding probe, ethidium bromide (EB), binds to the
6
0
DNA by intercalation. The competitive binding of the complex 1–4 by intercalation to CTꢀDNA was studied by measuring the
fluorescence of EB (2 ꢁM) bound CT−DNA (50 ꢁM) with increasing concentration of the complexes. The fluorescence
intensities of EB at 597 nm (excitation 510 nm) with an increasing amount of the complex concentration (0−90 ꢁM) was
measured with the aid of the same spectrofluorimeter as used above.
8

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2
.6 DNA cleavage experiments
For DNA cleavage experiments, 300 ng supercoiled (SC) pUC19 DNA was used and all experiments were carried out in 50 mM
TrisꢀHCl buffer (pH 8.0) containing 1% DMF and 10 mM phosphate buffer (pH 7.8) containing 1% DMF.
2
.6.1 Chemicalꢀinduced DNA cleavage. For chemical nuclease studies, the reactions were performed in the dark using
hydrogen peroxide (0.5 mM) as the oxidising agent in the absence and presence of complexes (1–500 µM). The solutions were
incubated at 37 ºC for 3 h and analysed for DNA cleaved products by agarose gel electrophoresis.
4
7
2
.6.2 Photoꢀinduced DNA cleavage. The photoꢀinduced DNA cleavage activity was done as described previously. The
photolytic DNA cleavage experiments were carried out on supercoiled (SC) pUC19 DNA (300 ng) with complexes (1–500 µM)
in 50 mM Tris HCl buffer (pH 8.0) containing 1% DMF and in 10 mM phosphate buffer (pH 7.8) containing 1% DMF. The
extent of DNA cleavage was measured from the intensities of the bands using the UVP Gel Documentation System. The
observed error in measuring the band intensities ranged between 3–6%. The mechanistic studies were performed using four
different additives: two are singlet oxygen quenchers (sodium azide and Lꢀhistidine) and other two are hydroxyl radical
quenchers (KI and Dꢀmannitol) prior to the addition of the complex. The concentration of each additive was 0.5 mM.
2
.7 Cytotoxicity studies
.7.1 Cell Culture. Human cervical cells HeLa were obtained from National Centre of Cell Science (NCCS), Pune, India,
2
and were maintained in minimal essential medium supplemented with 10% fetal bovine serum, penicillinꢀstreptomycin solution
and incubated at 37°C in 5% CO and 95% humidified incubator. The complexes were dissolved in DMSO at a concentration of
2
1
00 mM as stock solution, and diluted in culture medium at concentrations of 12.5, 25.0, 50.0 and 100.0 ꢁM as working
solutions. To avoid DMSO toxicity, the concentration of DMSO was less than 0.1% (v/v) in all experiments.
2
.7.2 MTT assay. HeLa cells were harvested from maintenance cultures in logarithmic phase, after counting in a
4
hemocytometer using trypan blue solution. The cell concentration was adjusted to 5x10 cells/ml and the cells were plated in 96
well flat bottom culture plates and incubated for 72 h with various concentrations of the test compounds. The effect of the drugs
on the cancer cell viability was studied using MTT dye reduction assay by measuring the optical density at 595 nm using microꢀ
61
plate reader spectrophotometer (PerkinꢀElmer 2030).
.7.3 Nuclear Staining. Nuclear staining using DAPI stain was performed according to the method previously described.
Briefly, HeLa cells either treated or untreated with test compounds were smeared on a clean glass slide, cells were fixed with
.7% formaldehyde for 15 min, permeabilised with 0.1% Triton Xꢀ100 and stained with 1 ꢁg/ml DAPI for 5 min at 37 °C. The
6
2
2
3
cells were then washed with PBS and examined by fluorescence microscopy (Olympus IX 71) to ascertain any condensation or
fragmentation of the nuclei indicating cells undergoing apoptosis.
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3. Results and discussion
3
.1 Synthesis
Four oxidovanadium(V) complexes (1–4) have been prepared using different aroylhydrazones as ligands (Scheme 1), i.e. with
varying heterocycleꢀderivatives in their hydrazone moieties. Reactions of the selected aroylhydrazones with VO(acac) proceed in
2
refluxing ethanol / DMF+MeOH to afford crystalline products in good yields.
Complexes (1–4) were completely soluble in DMF, DMSO and partially soluble in CH Cl and H O. All the complexes were
2
2
2
stable in both solid and solution phases. The solution phase stability over the period of the biological assays, i.e. 72 h, of the
complexes was confirmed by electronic absorption and NMR spectral studies (Fig. S1†).
3
.2 Spectral Characteristics
1
–2
3–4
The spectral (IR, UVꢀVis and NMR) data of HL , H2L and their corresponding oxidovanadium(V) complexes (1–4) are given
in the Experimental Section.
3
.2.1 IR spectroscopy. The disappearance of characteristic bands due –NH and –C=O in the respective ligand spectra, and
–1
the appearance of new bands in the range 1255–1238 cm in the spectra of the resulting complexes indicate the enolisation of
these two groups to form a –N=C–O bond sequence. The strong and sharp peak displayed by the complexes in the range 1612–
ꢀ
1
63,64
ꢀ1
1
594 cm is likely to be associated with the –C=N–N=C– moiety.
The presence of bands at 946 and 943 cm for 1 and 2,
4
7,65
respectively, is assigned to V=O stretching,
which indicates the presence of dioxido group whereas a sharp band in the range
ꢀ
1
63a
9
98–964 cm is assigned to V=O stretching of a oxidoalkoxido vanadium(V) complex, as for 3 and 4.
–4
3
.2.2 Electronic spectra. The electronic spectra of 1–4 were recorded in DMF (1.48 x 10 M) and are similar. A
representative spectrum of 3 is shown in Fig. 5. Strong absorptions in the range 460–399 nm are assignable to the ligandꢀtoꢀmetal
charge transfer transitions whereas the bands in the higher energy region (326–258 nm) are likely to be due to ligand centred
6
3a,b
transitions.
1
13
3
.2.3 NMR spectra. The H and C NMR data of ligands and their complexes were recorded in DMSOꢀd . The spectra of
6
1–2
3–4
HL and H2L exhibit resonances due to –NH in the range 14.50–11.09 ppm. These disappear in the spectra of the complexes
confirming enolisation of the ligands. For 3 and 4, the absence of signals in the range 13.32–12.72 ppm indicates deprotonation
of the –OH groups and its coordination to the vanadium centre after metallation. Complexes 1 and 3 each displays a singlet in the
range δ = 2.93–2.71 ppm due to –CH protons. Complexes 3 and 4 exhibit separate resonances corresponding to bound ethanolate
3
63b,66,67
at 5.59 and 5.67 ppm due to –CH and 1.54 and 1.55 ppm due to –CH , respectively.
In addition, 4, exhibits a quartet at
2
3
3
.44 ppm due to –CH and a triplet at 1.06 ppm due to –CH , respectively which indicates the presence of the coordinated ethanol
2
3
molecule. Finally, 4 displays a resonance at δ = 9.61 ppm ascribed to –CH.
51
The V NMR spectra of 1–4 each displays a singlet in the range δ= –541 to –507 ppm. These chemical shifts are usual for
6
8
complexes containing oxidovanadium(V) species.
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0

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3
.3. Electrochemical properties
The cyclic voltammogram of 1−4 were taken in CH Cl solution (0.1 M TEAP) and the redox potentials are summarized in Table
2
2
2
. The dioxidovanadium(V) complexes (1 and 2) are found to display two quasi reversible reduction wave at E1/2 values within
the potential window −0.87 to −0.70 V and −1.11 to −1.02 V, which are assigned to the V(V)/V(IV) and V(IV)/V(III) redox pair
6
9
of the metal respectively. Fig. 6a depicts a representative voltammogram of 2. Similarly the monooxido vanadium(V)
3+
2+
63b,70
complexes (3 and 4) display a wellꢀdefined quasi reversible cyclic voltammetric response due to the VO −VO couple
at
the range E1/2 = 0.40 − 0.36V (ꢁE = 164 − 150 mV), A representative voltammogram of 4 is displayed in Fig. 6b.
P
3
.4 Description of the Xꢀray crystal structures
.4.1 Structural commentary. Crystal structures were obtained for each of 1–4 and these are discussed in turn below. The
molecular structures are shown in Figs 1–4, Figs S2a and S3a†, and selected geometric parameters are listed in Table 3.
.4.2 Molecular structures. In 1, Fig. 1, the pentaꢀcoordinated vanadium atom is bound by a tridentate uninegative anion,
3
3
coordinating via enolateꢀO, azoꢀN and pyridylꢀN atoms, and two oxidoꢀO atoms. Confirmation that the multidentate ligand is
functioning as a 2ꢀacetylpyridine 2ꢀthiophenoylhydrazone is found in the magnitudes of the two C=N bond lengths, Table 3; a
similar mode of coordination for this specific anion toward vanadium was found in a sevenꢀcoordinated species where one oxidoꢀ
71
O atom is replaced by peroxoꢀ and methanolꢀO atoms. The resulting N O donor set in 1 is distorted toward square pyramidal
2
3
based on a τ value of 0.30 which compares to ideal values of 0.0 and 1.0 for square pyramidal and trigonal bipyramidal
7
2
geometries, respectively. Each fiveꢀmembered chelate ring is planar with r.m.s. deviations of 0.021 and 0.023 Å for the
V1,O1,N2,N3,C7) and (V1,N1,N2,C1,C6) rings, respectively, and the dihedral angle between them is 1.91(6)º. The dihedral
(
angle between the terminal pyridyl and thienyl (major component) rings is 4.96(19)º suggesting that the entire ligand is
approximately planar. The overall molecular structure in 1 matches closely to the most relevant structure available for
comparison, namely the furanyl derivative, which exhibits crystallographically imposed mirror symmetry with the tridentate
7
3
ligand lying on the plane.
The molecular structures of the two independent molecules comprising the asymmetric unit of 2 are shown in Figs 2 and
S2a†, and these are similar to that just described for 1 except that the methyl and thienyl groups in 1 have been substituted by
phenyl and anilinyl groups, respectively. The uninegative, tridentate ligand adopts a similar mode of coordination as just
described for 1 in each molecule of 2. The independent molecules differ most notably in the relative orientations of the phenyl
and anilinyl substituents as highlighted in the overlay diagram, Fig. S2b†, and quantified in the dihedral angles formed between
the pyridyl and each of the phenyl and anilinyl rings of 54.39(8) and 23.36(8)º, respectively, for the first independent molecule in
2
, and 47.77(7) and 6.18(8)º for the second molecule. From these data, the tridentate ligand in the second molecule adopts a more
planar conformation cf. the first independent molecule. The fiveꢀmembered chelate rings in each molecule are almost planar with
1
1

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the r.m.s. deviations for the (V1,O1,N2,N3,C7) and (V1,N1,N2,C1,C6) rings being 0.046 and 0.009 Å for the first molecule,
respectively, and 0.044 and 0.028 Å for the second molecule, respectively. The values of τ calculated for the two independent
molecules are 0.35 and 0.28, respectively, again indicating a distortion toward a square pyramidal geometry. Each molecule
7
4
features an intramolecular N–H...N hydrogen bond which closes an S(6) loop. The analogous ligand where the phenyl group
75
76
has been substituted for a methyl group has been characterized in two crystal structures, i.e. mononuclear cobalt and uranium
complexes, each with a nearly planar conformation of the tridentate ligand.
An increase in coordination number at vanadium in 3, Fig. 3a, is evident owing to the formation of intermolecular V–
N1(pyridyl) bonds. The vanadium atom is coordinated by the enolateꢀO1, phenoxideꢀO2 and azoꢀN2 atoms of the dinegative
ligand, as well as oxidoꢀO3, ethanolateꢀO4 and bridging pyridylꢀN1 atoms; the dianion is therefore, tetradentate. The result of the
V–N bridging is the formation of a supramolecular helical chain along the cꢀaxis, Fig. 3b. The N O donor set defines a distorted
2
4
octahedral geometry with the twist angle between the trigonal faces defined by the O1,O4,N1 atoms and the O2,O3,N2 atoms
being approximately 51º, cf. 60º for an ideal octahedron and 0º for an ideal trigonal prism. While the fiveꢀmembered chelate ring
is essentially planar, r.m.s. deviation = 0.033 Å, the sixꢀmembered chelate ring is best described as being based on a halfꢀchair
conformation with the vanadium atom lying 0.812(3) Å out of the plane defined by the remaining chelate atoms, i.e.
O2,N2,C1,C6,C7, which have a r.m.s. deviation of 0.0787 Å from their leastꢀsquares plane. There are no literature precedents for
crystal structures containing the ligand as found in 3.
The fourth structure, 4, is illustrated in Figs 4 and S3a†, as the asymmetric unit comprises two independent complex
molecules; for the second independent molecule, the ethanol molecule is statistically disordered over two positions (Fig. S3a†).
The overlay diagram of the two independent molecules is shown in Fig. S3b†.The coordination geometry is based on an
octahedral NO donor set defined by enolateꢀO1, azoꢀN2 and naphthylꢀoxideꢀO2 atoms of the tridentate dinegative anion, as well
5
as oxidoꢀO4, ethanolateꢀO5 and ethanolꢀO6 atoms. The V=O4 bond lengths in 4 are the shortest across the series owing to the
trans disposition of the oxidoꢀO to the relatively weakly bound ethanol molecules. Each fiveꢀmembered chelate ring is planar but
the sixꢀmembered rings are best described as having halfꢀchair conformations with the O2 atom lying 0.454(3) Å out of the plane
defined by the remaining five atoms (r.m.s. deviation = 0.1037 Å); the equivalent values for the V2ꢀmolecule are 0.348(3) and
0
.0826 Å, respectively. The dihedral angle between the chelate rings for the V1ꢀcontaining molecule is 6.88(18)º, and this opens
up to 10.06(12)º for the species with V2. Overall, the multidentate ligand exhibits a small twist as seen in the dihedral angles
formed between the outermost ring of the naphthalyl system and furanyl ring of 13.90(19) (V1) and 10.42(18)º (V2). A search of
7
7
the Cambridge Structural Database revealed 42 vanadiumꢀcontaining structures with an NO donor set defined by the same
5
donor atoms as for 4. In each structure there was a trans arrangement of oxidoꢀO and ethanolꢀO atoms as well as between
ethanolateꢀO and nitrogen donor atoms. The most closely related structure is one where the furanyl ring has been replaced by a
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78
3
ꢀpyridyl ring and the N1ꢀimine C atom carries a methyl group. In the latter, the tridentate ligand has a decidedly curved
conformation with the dihedral angle between the terminal rings being 34.56(7)°.
.4.3 Supramolecular features. In the absence of functional groups capable of conventional hydrogen bonding, the most
3
significant intermolecular interactions operating in the structure of 1 are of the type π…π and C–H…O. The former connect
molecules into centrosymmetric dimers via interactions involving the thienyl and pyridyl rings. These are connected into a threeꢀ
dimensional architecture via pyridylꢀC–H interacting with a bifurcated oxidoꢀO3 atom. A view of the unit cell contents for 1 is
given in Fig. S4† and full geometric details describing the aforementioned interactions are collected in the figure caption.
The most prominent feature of the crystal packing in 2 is the formation of N–H…O hydrogen bonds between the aminoꢀH
and oxidoꢀO atoms so that supramolecular chains aligned along [101] are formed comprising alternating V1ꢀ and V2ꢀcontaining
molecules, Fig. S5a†. These are linked by a network of C–H…O interactions involving anilinylꢀ, methylꢀ, phenylꢀ and pyridylꢀH
as donors and oxidoꢀO as acceptors, see Fig. S5b† for geometric data.
In the crystal structure of 3, the coordination polymers are connected into a supramolecular layer in the acꢀplane with the
links being of the type C–H…π(chelate) ring with the latter comprising the (V,O1,N2,N3,C8) atoms of the fiveꢀmembered ring,
see Fig. S6a† for geometric data. Such C–H…π(chelate) ring interactions are relatively rare, or more likely not always
79–81
recognised, but are increasingly attracting attention in the crystallographic literature.
..ABA... pattern with no specific interactions between them, see Fig. S6b†.
Finally, in the crystal packing of 4, each of the two independent molecules comprising the asymmetric unit selfꢀassociate via
methanolꢀO–H...N1(imine) hydrogen bonds to form twoꢀmolecule aggregates and 10ꢀmembered {...HOVN } supramolecular
The layers stack along the bꢀaxis in an
.
2
2
synthons, see Figs S7a, b†. Within the dimeric aggregates there are also π...π interactions between the thienyl and inner ring of
the naphthalyl group. The dimers stack into columns aligned along the bꢀaxis, so that layers of alternating V1ꢀ and V2ꢀcontaining
molecules stack along the aꢀaxis. In the layers comprising V2ꢀcontaining molecules, naphthalylꢀC–H...O(naphthalyl) interactions
stabilise the architecture but there are no specific interactions between V1ꢀcontaining dimers or between layers, Fig. S7c†.
3
.5 DNA binding studies
.5.1 Absorption spectroscopic studies. The binding propensity of 1–4 to CTꢀDNA was studied using different
spectroscopic techniques. Complexes 1–4 show absorption bands in the regions 440–400 and 340–300 nm which are attributed to
3
63a
L–V(dπ) LMCT and intraꢀligand transitions, respectively. UVꢀVis titration experiments were carried out to determine the
equilibrium binding constant (K ) of the complexes to CTꢀDNA (Table 4 and Fig. 7). Upon addition of CTꢀDNA, the UVꢀVis
b
absorption bands in the region 340–300 nm showed a hypochromic shift for 1 whereas 2–4 exhibited hyperchromism (Fig. 7),
indicating interaction between each complex and CTꢀDNA. The observed hypochromicity of complex 1, may be due to the
interaction between the electronic states of the chromophores and the DNA bases. On the other hand, the observation of
1
3

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hyperchromism in the intraꢀligand transition band near 300 nm for complexes 2–4 may be due to the electrostatic binding of the
complex on the DNA surface which results in a distortion of coordination sphere which leads to an enhancement of the
5,82,83
absorption intensity in the intraꢀligand transition bands.
The binding affinity of the interaction between CTꢀDNA and each of 1–4 is indicated by the binding constant, K , which
b
3
6e
was calculated using the following equation.
[
ꢀꢁꢂ]
[ꢀꢁꢂ]
ꢉ
=
ꢇ
+ꢇꢊ
Eq. 1
ꢃ
ꢄ
ꢅꢃ
ꢆ
ꢃ
ꢈ
ꢅꢃ
ꢆ
(ꢃ ꢅꢃ )
ꢈ ꢈ ꢆ
where [DNA] is the concentration of DNA base pairs, ε , ε , and ε correspond to apparent extinction coefficient for the complex
a
f
b
i.e. Abs/[complex] in the presence of DNA, in absence of DNA and to fully bound DNA, respectively. A plot of [DNA]/(ε – ε )
a
f
vs [DNA] gave a slope and intercept equal to 1/(ε – ε ) and 1/K (ε – ε ), respectively; K is calculated from the ratio of the slope
b
f
b
b
f
b
5
3
−1
to the intercept. The data reported in Table 4 reveal K has values for 1–4 from 1.13 × 10 to 5.03 × 10 M with the order of
b
DNA binding strength being 2 > 1 > 3 > 4. The binding affinity of the ligands to CTꢀDNA were also measured and gave values
lower than their corresponding complexes (Fig. S8 and Table S1†).
3
.5.2 Thermal denaturation studies. To investigate whether the stability of DNA alters upon interaction with the
57
complexes, DNA melting experiments were performed in the absence and presence of 1–4. The midpoint transition temperature
or melting temperature (T ) of CTꢀDNA in absence of any complex is ~65.7 °C. The T increased very slightly (~1.3ꢀ1.8 °C)
m
m
upon interaction with 1–3 whereas 4 showed a slight decrease (1.7 °C) in the thermal melting of CTꢀDNA (Fig. 8 and Table 4).
The low ꢂT values for 1–3 primarily suggest groove binding of the complexes to CTꢀDNA rather than an intercalative mode of
m
84,85
binding to DNA which usually gives a large positive ꢂT value.
The slight negative ∆T value of ꢀ1.70 °C for 4 (Table 4 and
m
m
86
Fig. 8) may be due to the destabilisation of the DNA double helix by this complex.
3
.5.3 Circular dichroism studies. The conformational changes in CTꢀDNA upon the interaction with complexes was
87
examined using circular dichroism (CD) spectroscopy. The Bꢀconformation of CTꢀDNA shows two conservative CD bands in
the UV region, a positive band at 275 nm due to base stacking and a negative band at 245 nm due to rightꢀhanded helicity.
Groove binding and electrostatic interaction of small molecules shows less or no perturbation on the base stacking and helicity
bands whereas an intercalation mode can induce intensity changes of both bands, thus modulating the rightꢀhanded Bꢀ
8
7
conformation of DNA. The CD spectra of CTꢀDNA (100 ꢀM) in the presence of the complexes showed very slight changes, for
the positive band at 275 nm as well as for the negative band at 245 nm, implying that the interactions were possibly groove
binding in nature (Fig. 9).
3.5.4 Competitive binding studies. In order to ascertain the binding mode of the complexes with CTꢀDNA, we have
performed the displacement assay or competitive binding experiment with three fluorescent dyes i.e. 4',6ꢀdiamidinoꢀ2ꢀ
phenylindole (DAPI), methyl green (MG) and ethidium bromide (EB). Among these dyes, DAPI binds to the minor groove of
1
4

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58ꢀ60,88
DNA, MG binds to the major groove of DNA while EB binds to DNA though intercalation.
The binding of these dyes to
DNA leads to an increase in the quantum yield of the dye due to which they give enhanced fluorescence upon binding to
58ꢀ60
DNA.
The minor groove competitive binding experiment was performed by titrating DAPI bound CTꢀDNA solution with increasing
amount of complexes. The fluorescence intensities of DAPI bound CTꢀDNA at 455 nm (excitation 358 nm) were measured with
an increase of complex concentration (0–90 µM). The competitive binding of the complexes to CTꢀDNA led to the quenching of
the emission intensity of the DAPI bound CTꢀDNA at 455 nm which is possibly due to the displacement of the bound DAPI to
CTꢀDNA by the complexes. Complex 4 showed the highest quenching of DAPI bound to CTꢀDNA fluorescence intensity at 455
nm i.e. ~ 80% , followed by complex 2 and 3 (~60%) (Fig.10). Complex 1 showed ~20% quenching of the fluorescence intensity
at 455 nm which is least among the four complexes (Fig. 10). Therefore our data clearly suggest that all the complexes bind to
CTꢀDNA through minor groove binding mode, as depicted from the decrease in the fluorescence intensity of DAPI bound to CTꢀ
DNA at 455 nm.
Similar competitive binding experiments were also performed to ascertain the major groove and intercalative binding mode of
complexes with CTꢀDNA by monitoring the fluorescence intensity of MG bound CTꢀDNA at 672 nm and EB bound CTꢀDNA at
5
97 nm respectively (Figs. S9 and S10). On addition of the complexes to MG bound CTꢀDNA, the emission intensity at 672 nm
did not quench significantly (≤2−4%) with the increasing concentration of the complexes from 0−90 µM (Fig. S9). This showed
that the complexes 1−4 were not able to displace MG bound to CTꢀDNA. This clearly suggests that the complexes do not interact
with CTꢀDNA by major groove binding mode. Similar results were also obtained when EB bound CTꢀDNA was titrated with
increasing concentration of the complexes (0–90 µM) (Fig. S10). As there was no significant decrease (≤2−4%) in the emission
intensity of EB bound CTꢀDNA at 597 nm, it reveals that the complexes do not bind to CTꢀDNA through intercalation.
Therefore, from the fluorometric competitive binding studies it is concluded that the complexes (1−4) interact with CTꢀDNA via
minor groove binding mode.
3
.6 DNA cleavage studies
.6.1 Chemicalꢀinduced DNA cleavage. The DNA cleavage activity of the complexes 1–4 (1–500 ꢁM) was studied in the
3
dark in the presence of hydrogen peroxide (500 ꢁM) as the oxidising agent using supercoiled pUC19 DNA (300 ng) in 50 mM
TrisꢀHCl buffer (pH 8.0) containing 1% DMF. The complexes do not show any chemicalꢀinduced DNA cleavage activity.
3
.6.2 Photoꢀinduced DNA cleavage. To investigate whether the DNA binding properties of the complexes were
accompanied with photoꢀnuclease activity, a photoꢀinduced DNA cleavage activity assay was performed. The photoꢀinduced
DNA nuclease activity of 1–4 was studied using supercoiled (SC) pUC19 DNA in 50 mM TrisꢀHCl buffer (pH 8.0) containing
1
% DMF upon irradiation of UVA light of 350 nm in the presence and absence of the complexes (Fig. 11). DNA cleavage was
1
5

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indicated by the decrease in the supercoiled pUC19 DNA (Form I) and subsequent formation of nicked circular DNA (Form II)
and linear DNA (Form III). The percentage of net DNA cleavage by the complexes was estimated using the following equation:
ꢋꢌꢍꢎꢇꢏꢏꢐꢑꢒꢇ×ꢋꢌꢍꢎꢇꢏꢏꢏꢐ
ꢋꢌꢍꢎꢇꢏꢏꢓꢑꢒꢇ×ꢋꢌꢍꢎꢇꢏꢏꢏꢓ
NetꢇDNAꢇcleavageꢇ% =
−
Eq. 2
ꢋꢌꢍꢎꢇꢏꢐꢑꢇꢋꢌꢍꢎꢇꢏꢏꢐꢑꢒꢇ×ꢋꢌꢍꢎꢇꢏꢏꢏꢐ
ꢋꢌꢍꢎꢇꢏꢓꢑꢇꢋꢌꢍꢎꢇꢏꢏꢓꢑꢒꢇ×ꢋꢌꢍꢎꢇꢏꢏꢏꢓ
89
The subscripts “s” and “c” refers to the sample and control, respectively. The DNA cleavage activity of the complexes were
carried out with increasing concentration of the complexes from 1 –500 µM. The net DNA cleavage percent by 1–4 was plotted
in a concentration dependent manner in Fig. 12. Approximately 10% DNA cleavage was observed in the presence of 10 ꢀM of 1–
4
(Fig. 12, inset). The cleavage activity was saturated for each of the complexes at a concentration of 100 ꢀM. At this
concentration, the photoꢀnuclease activity of 1, 2, 3 and 4 was 50, 33, 65 and 45%, respectively, indicating that 3 has the highest
photoꢀnuclease activity over the series. Further at 500 µM of complex concentration, 2−4 showed a slight decrease in the net
DNA cleavage activity while for complex 1 no such change was observed. Cleavage activity of the complexes (1−4) varies
4
7
because of the different functional group present in their ligand moieties. Different heterocycles present in the ligand
90
environment may also have influence on their observed cleavage efficiency. The photoꢀinduced DNA cleavage experiments
were also carried out with phosphate buffer returning similar activity as for the experiments in tris buffer, an observation in
47
accord with previous studies with dioxidovanadium complexes. Control experiments revealed that neither DMF (1%) nor the
ligand molecules showed any photoꢀinduced DNA cleavage activity, implying that DMF and the ligands are cleavage inactive
under similar conditions (Fig. S11†).
In order to understand the mechanistic pathways involved in the photoꢀcleavage reactions, the photoꢀinduced DNA cleavage
activity of 1–4 was investigated in the presence of various additives. The DNA cleavage reaction involving molecular oxygen can
1
proceed in two mechanistic pathways: (a) a typeꢀII process involving singlet oxygen species ( O ), or (b) by a photoꢀredox
2
91
pathway involving reactive hydroxyl radicals (OH). The addition of NaN (singlet oxygen quencher) inhibited the DNA
3
cleavage activity of 1, 3 and 4 by 8, 14 and 10 %, respectively (Fig. 13). By contrast, the other singlet oxygen quencher (Lꢀ
histidine) slightly inhibited the photoꢀinduced DNA cleavage activity of complexes 1, 3 and 4. Surprisingly, it was observed that
the DNA cleavage activity of 2 did not alter in the presence of these two additives. The hydroxyl radical scavenger, KI, inhibited
the photoꢀnuclease activity of 1, 3 and 4 by 10, 15 and 13 %, respectively, whereas the other hydroxyl radical scavenger Dꢀ
mannitol inhibited the DNA cleavage activity of these three complexes moderately (Figs 13 and S12†). However, no inhibition of
DNA cleavage was observed for 2 in the presence of these two additives (Fig. 13). Therefore, it is concluded that 1, 3 and 4
exhibit photoꢀinduced DNA cleavage activity via both singlet oxygen and hydroxyl radical pathways, while the mechanistic
pathways involved in the photoꢀinduced DNA cleavage by 2 cannot be stated with certainty.
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3
.7 Cytotoxicity studies
.7.1 MTT assay. The in vitro cytotoxicities of 1–4 were evaluated by the MTT assay against the cervical cancer cell line
3
1–2
3–4
HeLa; the IC50 values are presented in Table 5. The species HL , H2L and VO(acac) gave high IC values of > 200 ꢁM,
2
50
whereas 1–4 gave values in the range 10–20 ꢁM. Under the same experimental conditions, the commonly used chemotherapeutic
drugs cisplatin, gefitinib, gemcitabine, 5ꢀfluorouracil and vinorelbine are comparably effective in HeLa cells with IC50 values of
9
2
1
3, 20, 35, 40 and 48 ꢁM, respectively. The significant decrease in the inhibitory ability of the ligand molecules as well as their
lower binding affinity to CTꢀDNA compared to their metal complexes clearly indicates that incorporation of vanadium has a
marked effect on cytotoxicity. A possible explanation is that by coordination, the polarity of the ligand and the central metal ion
are reduced through charge equilibration which favours permeation of the complexes through the lipid layer of the cell
93,94
membrane.
The results of DNA binding/cleavage ability for the ligands (Figs S8 and S11†) have been considered and are
95
consistent with the observation that metal complexes can exhibit greater biological activities than the free ligands.
Within the series 1–4, the cytotoxicities of 1, 2 and 3 are approximately similar whereas 4 is more potent, Table 5. Their
dose dependencies are illustrated in Fig. 14. The variation in cytotoxicity may be affected by the various functional groups
attached to the aroylhydrazone derivative. Very recently, the antiꢀproliferative activity of some vanadium complexes were
9
6
32,47
reported by Yamaguchi et al. and us
using U937 cells and HeLa cells, respectively. The present results are in accord with
these results.
3
.7.2 Nuclear Staining Assay. To investigate the apoptotic potential of the test complexes in HeLa cells, DAPI staining was
performed. Chromatin condensation during the process of apoptosis (type I programmed cell death) is a characterizing marker of
nuclear alteration. HeLa cells were treated with 16, 15, 16, and 5 ꢁM of 1–4, respectively. The cells were incubated for 24 h
before DAPI nuclear staining and were examined under a fluorescent microscope fitted with a DAPI filter; all images are shown
in grayꢀscale. As shown in Fig. 15, control cells hardly showed any sort of condensation in comparison to the treated cells. The
treated cells demonstrate the brightly condensed chromatin bodies and the nuclear blebbings as marked by arrows in Fig. 15.
Besides showing nuclear change, the treated cells exhibited a shrinking morphology which is another important hallmark of
apoptosis.
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4
. Conclusions
The synthesis and characterization of four neutral oxidovanadium(V) complexes (1−4) with hydrazone scaffolds containing either
furan, thiophene and pyridine residues has been achieved. Xꢀray crystallography confirm the spectroscopic analyses and show
N O fiveꢀcoordinate geometries, distorted toward square pyramidal, for each of 1 and 2. The dianion in 3 is tetradentate as the
2
3
pyridylꢀN atom also bridges a second V atom leading to a helical coordination polymer and an N O octahedral geometry. A NO
5
2
4
octahedral geometry is found in mononuclear 4. Biological studies reveal that 1–4 show considerable DNA binding propensity.
DNA binding activities were investigated using UVꢀVis absorption titration, circular dichroism, thermal denaturation and
fluorometric competitive binding studies, and shows that the complexes interact with CTꢀDNA via minor groove binding mode,
3
5
ꢀ1
with binding constants ranging from 10 −10 M . All complexes show good photoꢀinduced cleavage of pUC19 supercoiled
plasmid DNA with 3 showing the highest photoꢀinduced DNA cleavage activity of ~65%. The results from the mechanistic study
suggest that the photolytic DNA cleavage of 1, 3 and 4 proceeded via both singlet oxygen and hydroxyl radical pathways.
Additionally, the cytotoxic activities of 1–4 were evaluated against a human cervical cancer cell line (HeLa). All are appreciably
cytotoxic compared with standard drugs (cisplatin, gefitinib, gemcitabine, 5ꢀfluorouracil and vinorelbine) with IC50 values
ranging from 10 to 20 µM. This potency correlates with the presence of a heterocycle attached to the hydrazone moiety. Despite
having a lower binding affinity, the higher cytotoxicity of 4 may be due to the destabilising interaction between the complex and
the DNA as evidenced from the thermal denaturation data. The results reported herein will inspire further work on
oxidovanadium(V) complexes for the development of metalꢀbased agents for antiꢀcancer applications.
Acknowledgments
The authors thank the reviewers for their comments and suggestions, which were helpful in preparing the revised version of the
manuscript. Funding for this research was provided by the Department of Science and Technology, India [Grants SR/WOSꢀ
A/CSꢀ145/2011 (S.P.D.) and SR/FT/CSꢀ016/2008 (R.D.)] and the Council of Scientific and Industrial Research, India [Grant
3
7(1535)/12/EMRꢀII) (A.B.)]. A.K.P. acknowledges the receipt of a fellowship from ICMR, India [Grant 45/25/2012ꢀBio/BMS].
R. D. thanks Prof. S. K. Chattopadhyay for electrochemical studies. The support from the High Impact Research MoE Grant
UM.C/625/1/HIR/MoE/SC/12 from the Ministry of Higher Education, Malaysia, is also gratefully acknowledged.
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Notes and references
Corresponding Author
*
Eꢀmail: rupamdinda@nitrkl.ac.in
Fax: +91 6612462022; Tel: +91 6612462657
Electronic supplementary information (ESI) available (Table S1, Figs S1−S12): CCDC reference numbers 1007034ꢀ1007037.
For ESI and crystallographic data in CIF format see DOI: XXXXX
†
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7 L. Li, Q. Guo, J. Dong, T. Xu and J. Li, Photochem. Photobiol. B, 2013, 125, 56−62.
8 A. K. Krey and F. E. Hahn, Biochemistry, 1975, 14, 5061−5067.
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1 P. K. Sasmal, S. Saha, R. Majumdar, S. De, R. R. Dighe and A. R. Chakravarty, Dalton Trans., 2010, 39, 2147−2158.
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4 P. G. Avaji, C. H. V. Kumar, S. A. Patil, K. N. Shivananda and C. Nagaraju, Eur. J. Med. Chem., 2009, 44, 3552−3559.
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6 T. Yamaguchi, S. Watanabe, Y. Matsumura, Y. Tokuoka and A. Yokoyama, Bioorg. Med. Chem., 2012, 20, 3058−3064.
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DOI: 10.1039/C4RA14369H
Table 1 Crystallographic and refinement details for 1–4
Compound
1
2
3
4
Formula
C H N O SV
C H N O V
C H N O V
C H N O V
20 21 2 6
1
2
10
3
3
19 15
4
3
16 16
3
4
Formula weight
327.23
398.29
365.26
436.33
Crystal colour/habit
Orange/prism
0.11 x 0.15 x 0.20
100
Darkꢀred/prism
0.14 x 0.17 x 0.34
100
Black/prism
0.06 x 0.11 x 0.25
293
Red/needle
0.03 x 0.07 x 0.40
293
Crystal dimensions/mm
Temperature/K
Crystal system
monoclinic
triclinic
orthorhombic
Pna21
monoclinic
C2/c
Space group
P2 /c
P¯1
1
a/Å
b/Å
c/Å
a/°
7.1876(3)
10.6882(4)
16.6167(6)
90
7.2745(2)
14.3309(3)
16.7694(4)
87.92(10)
80.07(10)
83.41(10)
1710.41(7)
4
12.4265(2)
10.8273(2)
12.0750(2)
90
41.893(4)
8.7854(10)
21.729(2)
90
β/°
92.567(2)
90
90
98.017(5)
90
γ
/°
90
3
V/Å
1275.26(9)
4
1624.64(5)
4
7919.2(14)
16
Z
ꢀ
3
D /g cm
c
1.704
1.547
1.493
1.464
F(000)
664
816
752
3616
ꢀ
1
ꢁ/mm
0.951
0.609
5.339
0.541
Measured data
θ range/°
43228
2.3–27.5
2933
29305
4415
114454
1.9–25.6
7421
1.9–27.5
7826
5.4–77.5
2329
Unique data
Observed data [I ≥ 2.0σ(I)] 2760
6698
2119
5085
R, obs. data; all data
0.028; 0.030
0.033; 0.040
0.046, 0.865
0.086; 0.091
0.030; 0.033
0.061, 0
0.081; 0.082
0.045; 0.80
0.052, 15.124
0.106, 0.130
a, b in weighting scheme
Rw, obs. data; all data
Residual electron density
0.035, 1.109
0.075; 0.076
3
peaks/e Å
1.03, ꢀ0.41
0.39, ꢀ0.42
0.22, ꢀ0.22
0.65, ꢀ0.39
2
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a
Table 2 Cyclic voltammetric results for 1−4 at 298 K
_________________________________________________________________
_
Complex
E1/2 (ꢂE ) / V(mV)
P
_
_________________________________________________________________
1
VO L (1)
−0.87 (240), −1.11 (100)
−0.70 (300), −1.02 (130)
0.36 (164)
2
2
VO L (2)
2
3
VOL (OEt) (3)
VOL (OEt)EtOH (4)
4
0.40 (150)
_
_________________________________________________________________
a
–1
In CH Cl at a scan rate of 100 mV s . E = (Epa + E )/2, where E and Epc are anodic and cathodic peak potentials vs.
2
2
1/2
pc
pa
Ag/AgCl, respectively. ꢂE = E – Epc..
P
pa
2
5

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Table 3 Selected bond lengths and angles (Å, º) for 1–4
_
_______________________________________________________________________________________
a
b
Complex
1
2a
2b
3
4a
4b
_
_______________________________________________________________________________________
Parameter
V–O1
1.9684(11)
1.6213(12)
1.6196(13)
–
1.9458(12)
1.6196(13)
1.6206(13)
–
1.9604(12)
1.6203(12)
1.6171(12)
–
1.969(2)
1.859(2)
1.594(2)
1.777(2)
–
1.958(2)
1.938(2)
V–O2
1.856(2)
1.848(2)
V–O3
–
–
V–O4
1.577(2)
1.575(3)
V–O5
–
–
–
1.775(2)
1.753(2)
V–O6
–
–
–
–
2.325(2)
2.46(3)
V–N1
2.1112(13)
2.1200(13)
–
2.0939(15)
2.1067(14)
–
2.1015(14)
2.1200(14)
–
2.434(2)
2.141(2)
–
–
–
V–N2
2.111(2)
2.109(2)
V–N3
–
–
N2–N3
C=N1
1.3840(18)
–
1.381(2)
–
1.3786(19)
–
1.400(3)
–
1.392(3)
1.395(3)
1.299(4)
1.297(4)
C=N2
1.291(2)
1.312(2)
102.55(6)
102.16(6)
109.99(7)
–
1.303(2)
1.324(2)
103.92(6)
104.12(6)
109.89(7)
–
1.304(2)
1.325(2)
101.83(6)
102.57(6)
109.15(7)
–
1.298(4)
1.297(4)
149.59(9)
100.86(11)
99.62(11)
–
1.285(4)
1.283(4)
C=N3
–
–
O1–V–O2
O1–V–O3
O2–V–O3
O4–V–O6
O1–V–N1
O3–V–N1
O4–V–N2
O5–V–N2
N1–V–N2
150.57(9)
151.97(10)
–
–
–
–
177.03(10)
171.5(6)
146.50(5)
97.60(6)
–
147.47(6)
95.25(6)
–
146.72(5)
97.00(6)
–
79.19(9)
178.81(11)
163.35(9)
–
–
–
–
–
93.61(11)
163.36(11)
–
96.30(11)
157.32(12)
–
–
–
–
73.08(5)
73.32(6)
72.94(5)
85.55(9)
_
__________________________________________________________________________
2
6

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a The N1 atom in 3 is related by the symmetry operation 2ꢀx, 2ꢀy, ½+z.
b For the second component of the disordered ethanol molecule in 4, the V2–O6b bond length is 2.52(3) Å and the O4a–V2–O6b
bond angle is 174.8(6)º.
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Table 4 DNA binding parameters for 1–4
a
ꢀ1
b o
Complex
Binding constant (K ) (M )
∆Tm ( C)
b
4
1
2
3
4
8.56 × 10
+1.31
5
1.13 × 10
+1.83
4
4.95 × 10
+1.32
3
5.03 × 10
ꢀ1.70
a
DNA binding constants were determined by the UVꢀvis spectral method.
Change in the melting temperature of CTꢀDNA.
b
2
8