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Non-covalent DNA binding and cytotoxicity of certain mixed-ligand ruthenium(II) complexes of 2,2'-dipyridylamine and diimines.
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PAPER
www.rsc.org/dalton | Dalton Transactions
Non-covalent DNA binding and cytotoxicity of certain mixed-ligand
ruthenium(II) complexes of 2,2 -dipyridylamine and diimines†
Published on 05 March 2008. Downloaded by Temple University on 27/10/2014 07:59:47.
Venugopal Rajendiran,a Mariappan Murali,‡a Eringathodi Suresh,b Mallayan Palaniandavar,*a
Vaiyapuri Subbarayan Periasamyc and Mohammad Abdulkader Akbarshac
Received 1st October 2007, Accepted 30th January 2008
First published as an Advance Article on the web 6th March 2008
DOI: 10.1039/b715077f
A series of mixed ligand ruthenium(II) complexes [Ru(Hdpa)2 (diimine)](ClO4 )2 1–5, where Hdpa is
2,2 -dipyridylamine and diimine is 1,10-phenanthroline (phen) and a modified/extended
1,10-phenanthroline such as, 5,6-dimethyl-1,10-phenanthroline (5,6-dmp),
dipyrido[3,2-d:2 ,3 -f ]quinoxaline (dpq), 5-methyldipyrido[3,2-d:2 ,3 -f ]quinoxaline (mdpq) and
dipyrido[3,2-a:2 ,3 -c]phenazine (dppz) have been isolated and characterized by analytical and spectral
methods. The complex [Ru(Hdpa)2 (phen)](PF6 )2 1 has been structurally characterized and the
coordination geometry around Ru(II) in it is described as distorted octahedral. 1 H NMR spectral data
reveal that 1–5 should have a C 2 symmetry lying on the diimine plane due to the rapid flapping of the
coordinated Hdpa ligands. The interaction of the complexes with calf thymus (CT) DNA has been
explored by using absorption and emission spectral and viscometry and electrochemical techniques and
the mode of DNA binding of the complexes has been proposed. The DNA binding affinity of the
complexes decreases with decrease in number of planar aromatic rings in the co-ligand supporting the
intercalation of the diimine co-ligands in between the DNA base pairs. Circular dichroic spectral
studies reveal that the complexes 3–5 exhibit induced circular dichroism upon binding to CT
DNA. Interestingly, upon interaction with CT DNA all the complexes show an increase in anodic
current in the cyclic voltammograms suggesting that they are involved in electrocatalytic guanine
oxidation. Interestingly, of all the complexes, only 5 alters the DNA superhelicity upon binding with
supercoiled pBR322 DNA, which is consistent with its higher DNA binding affinity. Further, the
cytotoxicities of the complexes against human cervical epidermoid carcinoma cell line (ME180) have
been examined. Interestingly, 5 exhibits a cytotoxicity against ME180 higher than other complexes with
potency approximately 8 times more than cisplatin for 24 h incubation but 4 times lower than cisplatin
for 48 h incubation.
Introduction
The designing of non-covalent DNA binding ruthenium(II)
anticancer drugs is growing in interest in the field of
metallopharmaceuticals1 because the covalently DNA binding
anticancer agent cisplatin possesses inherent limitations such as
high toxicity and low administration dosage.2 The mechanism of
action of DNA-targeting metal-based drugs has been thought
to involve covalent binding to nucleobase moieties and a low
degree of selectivity.3 So, there is considerable attention focused
on the design of new metal-based drugs that exhibit enhanced
selectivity and novel modes of DNA interaction such as nona
School of Chemistry, Bharathidasan University, Tiruchirappalli, 620 024,
Tamilnadu, India. E-mail: palanim51@yahoo.com
b
Analytical Science Discipline, Central Salt and Marine Chemical Research
Institute, Bhavnagar, 364 002, India
c
Department of Animal Science, Bharathidasan University, Tiruchirapalli,
620 024, India
† Electronic supplementary information (ESI) available: 1 H NMR spectral
data for ligands and Figs. S1 and S2. CCDC reference numbers 660567–
660570. For crystallographic data in CIF or other electronic format see
DOI: 10.1039/b715077f
‡ Present Address: Department of Chemistry, National College, Tiruchirappalli 620 001, Tamil Nadu, India.
This journal is © The Royal Society of Chemistry 2008
covalent interactions that mimic the mode of interaction of
proteins with DNA.4 Very recently non-covalent DNA binding
metal complexes,1 particularly, metallointercalators have received
attention in designing efficient anticancer drugs.5 Intercalation,
which is well-known to strongly influence the properties of DNA,
has been reported as a preliminary step in mutagenesis.6 Noncovalent interactions between transition-metal complexes and
DNA occur by intercalation, groove binding, or electrostatic
surface binding. In particular, metal complexes possessing planar
aromatic ligands, which bind to DNA by intercalation, is receiving
considerable attention.7 Such metallointercalators tend to be
strongly mutagenic and some have shown promising chemotherapeutic activity, which correlates well with their DNA binding
affinity.8 Also, complexes that contain the appropriately orientated
H-bonding functionalities permit effective binding to DNA either
in the major or minor grooves.9
Ruthenium complexes are regarded as promising alternatives to
platinum complexes and several ruthenium complexes have been
now proposed as potential anticancer substances,10 demonstrating
remarkable anticancer activity and showing general toxicity lower
than platinum compounds.11 Thus the Ru(III) complex ImH[transRuCl4 (DMSO)(Im)], NAMI-A, shows high selectivity for solid
tumor metastases12 (prevents spread of cancer) and low host
Dalton Trans., 2008, 2157–2170 | 2157
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toxicity13 and is the first ruthenium complex to enter clinical
trials.13b However, it fails to affect primary tumor growth14 and
does not exhibit cytotoxicity against tumor cells in vitro. A
related Ru(III) compound,15 KP1019, has also entered clinical
trials, as it was found to exhibit antiproliferative activity in
vitro in human colon carcinoma cell lines.16 Very recently, Sadler
and his co-workers have reported half-sandwich Ru(II) arene
complexes, which exhibit reproducible anticancer activity against
A2780 human ovarian cancer cell lines both in vitro and in
vivo.17 Since DNA has been identified as the possible primary
molecular target18 of metal-based anticancer agents such as
cisplatin,2a attention is mainly focused on interacting ruthenium
complexes with DNA to identify whether DNA binding is effective
and whether they can act as chemotherapeutic agents. These
ruthenium(III) complexes are considered as prodrugs because the
activity seems to be initiated by reduction of Ru(III) to Ru(II) once
the core complex is inside the anoxic cancer tissues, followed by
linkage to nucleic acids. Therefore, in the field of biocoordination
Chemistry development of Ru(II) anticancer complexes that have
both appreciable solubility in aqueous media and uptake by the
cells have received much attention because the reduction step
is skipped.19 In our earlier study we have shown8 that the noncovalently DNA binding complex [RuII (N2 S2 )(dppz)]2+ exhibits a
significant cytotoxicity against melanoma cancer cell line (A375).
In this report we explore the DNA binding properties of a series
of ruthenium(II) complexes of the type [Ru(Hdpa)2 (diimine)]2+
1–5, where Hdpa is 2,2 -dipyridylamine and diimine is a
modified/extended aromatic 1,10-phenanthroline (Scheme 1)
such as 1,10-phenanthroline (phen) (1), 5,6-dimethyl-1,10phenanthroline (5,6-dmp) (2), dipyrido[3,2-d:2 ,3 -f ]quinoxaline
(dpq) (3), 5-methyldipyrido[3,2-d:2 ,3 -f ]quinoxaline (5-mdpq) (4)
and dipyrido[3,2-a:2 ,3 -c]phenazine (dppz) (5). The rationale
behind the design of the complexes is that the bidentate primary
ligand Hdpa offers flexibility around the secondary amine nitrogen
bridging the two pyridine rings, the two pyridine planes adopting
either coplanar or tilted conformation upon coordination to
metal centers.20 Also, Hdpa possesses the potential to form
hydrogen bond with suitable DNA functionalities, which would
be expected to enhance the DNA binding affinity significantly.
The co-ligands phen and extended 1,10-phenanthrolines with
different numbers of aromatic rings to vary the surface area for
intercalation would be expected to act as affinity ligands, and
dictate the extent of DNA binding interaction of the complexes.
The complexes are coordinatively saturated and rigid and so
no covalent DNA binding is expected. The phen complex 1
has been structurally characterized. The DNA binding of the
Ru(II) complexes has been studied by using a variety of physical
methods such as absorption and emission spectroscopy, viscosity
and electrochemical techniques. The ability of the complexes to
unwind/untwist DNA has been studied by using supercoiled
pBR322 DNA. The cytotoxicity of the complexes 1–5 has been
studied by screening the complexes against human epidermoid
carcinoma (ME180) cell line. It is remarkable that the complex 5
exhibits a cytotoxicity higher than the other complexes. Also, it
displays cytotoxicity in the range of cisplatin, which is in clinical
use currently.
Experimental
Reagents and materials
RuCl3 ·3H2 O (Arora Matthey), 1,10-phenanthroline (Merck),
2,2 -dipyridylamine (Aldrich), calf thymus (CT) DNA (highly
polymerized stored at 4 ◦ C), Hoechst 33258 (Sigma), the
self-complementary oligonucleotides d(GCGCGCGCGCGC) referred as d(GC)12 , d(ATATATATATAT) referred as d(AT)12 and
d(CGCGATCGCG) referred as d(CGCGATCGCG)2 were purchased from Sigma and stored at −20 ◦ C. The lyophilized oligonucleotides were digested in Tris buffer and annealed using standard
procedures to make the double-stranded oligonucleotides and
stored at 4 ◦ C. The concentrations of the oligonucleotide solutions
were determined using the procedures provided by the supplier and
pBR322 supercoiled plasmid DNA (stored at −20 ◦ C) and agarose
(Genei) and cisplatin (Bristol-Myers Squibb Co., Princeton) were
used as received. Ultra-pure Milli-Q water (18.2 mX) was used in
all experiments. Reagent grade solvents were dried and distilled by
usual methods and the solvents were stored over molecular sieves
(4 Å).
Methods and instrumentation
Scheme 1 Possible coordination geometries of mixed ligand ruthenium(II) complexes of Hdpa and structures of diimine (N–N) co-ligands
and proton numbering pattern.
2158 | Dalton Trans., 2008, 2157–2170
Microanalysis (C, H and N) were carried out with Vario EL
elemental analyzer. An LCQ DECA XP electrospray mass spectrometer was employed for ESI-MS analysis. UV-Vis spectroscopy
was recorded on a Varian Cary 300 Bio UV-Vis spectrophotometer
using cuvettes of 1 cm path length. Emission intensity measurements were carried out using Jasco F 6500 spectrofluorometer.
Circular dichroic spectra of DNA were obtained by using JASCO
J-716 spectropolarimeter equipped with a Peltier temperature
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control device. Viscosity measurements were carried out using
Schott Gerate AVS 310 automated viscometer.
Solutions of DNA in the buffer 50 mM NaCl–5 mM Tris HCl
in water gave the ratio of UV absorbance at 260 and 280 nm,
A260 /A280 , as 1.921 indicating that the DNA was sufficiently free of
protein. Concentrated stock solutions of DNA (13.5 mM) were
prepared in buffer and sonicated for 25 cycles, where each cycle
consisted of 30 s with 1 min intervals. The concentration of DNA
in nucleotide phosphate (NP) was determined by UV absorbance
at 260 nm after 1 : 100 dilutions. The extinction coefficient, e260 ,
was taken as 6600 M−1 cm−1 . Stock solutions were stored at 4 ◦ C
and used after no more than 4 days. Supercoiled plasmid pBR322
DNA was stored at −20 ◦ C and the concentration of DNA in
base pairs was determined by UV absorbance at 260 nm after
appropriate dilutions taking e260 as 13 100 M−1 cm−1 . Concentrated
stock solutions of metal complexes were prepared by dissolving
calculated amounts of metal complexes in respective amount of
solvent and diluted suitably with corresponding buffer to required
concentrations for all the experiments.
Synthesis of ligands
The ligands dipyrido[3,2-d:2 ,3 -f ]quinoxaline (dpq),22 5-methyldipyrido[3,2-d:2 ,3 -f ]quinoxaline (mdpq)22 and dipyrido[3,2a:2 ,3 -c]phenazine (dppz)23 were synthesised according to literature methods.
Synthesis of Ru(II) complexes
CAUTION
During handling of the perchlorate salts of metal complexes with
organic ligands care should be taken because of the possibility of
explosion.
Synthesis of cis-[Ru(Hdpa)2 (Cl2 )]Cl·H2 O. A mixture of
RuCl3 ·3H2 O (0.52 g, 2 mmol) and Hdpa (4.2 mmol, 0.71 g) was
refluxed in 80 mL of ethanol for 8 h, and filtered. The filtrate was
concentrated to about 40 ml and a green product was obtained
after stored in a refrigerator for overnight.
Synthesis of [Ru(Hdpa)2 (phen)](ClO4 )2 1. A mixture of cis[Ru(Hdpa)2 (Cl2 )]Cl (0.15 g, 0.26 mmol) and phen (0.0515 g,
0.26 mmol) was heated to reflux in water (30 mL) for 15 min
under N2 atmosphere, after which the solution was cooled then
the reducing agent 30% H3 PO3 neutralized with NaOH was added
to the reaction mixture and the reflux were continued to 3 h and
treated with an excess of NaClO4 . The precipitated complex was
dried, dissolved in a small amount of acetonitrile, and purified by
chromatography over alumina using acetonitrile–methanol (3 : 1,
v/v) as an eluent. The red coloured compound was obtained after
drying in vacuo. Yield, 0.16 g, 73%. Red coloured single crystals
of 1, suitable for X-ray studies were obtained on slow evaporation
of the complex in an acetonitrile–n-butanol (1 : 3) mixture. Anal.
Calc. for RuC32 H26 N8 Cl2 O8 : C, 46.73; H, 3.19; N, 13.62. Found: C,
46.30; H, 3.20; N, 13.43%. kmax /nm (e/M−1 cm−1 ) (5% CH3 OH–
5 mM Tris-HCl–50 mM NaCl buffer (0.5 : 9.5, v/v) buffer at
pH 7.1): 505 (sh), 451 (6582), 423 (sh) (7113), 380 (7895), 283
(45 211), 265 (61 386), 230 (41 147). ESI-MS: [Ru(Hdpa)2 (phen)]2+
displays a peak at m/z = 312.0, calc. 311.8. 1 H NMR (DMSOd6 , 400 MHz): d (multiplicity, integration, assignment, J/Hz,
This journal is © The Royal Society of Chemistry 2008
coordination-induced shifts (c.i.s.), d complex – d ligand )/ppm. Group
I, 8.734 (d, 2H, H2 , 8.1, −0.514), 8.042 (t, 2H, H3 , 4.1, 0.178),
9.173 (d, 2H, H4 , 5.2, 0.603), 8.258 (s, 2H, H5 , 0.210). Group II,
7.410 (d, 2H, H3 , 8.3, −0.327), 7.986 (t, 2H, H4 , 4.4, 0.350), 6.983
(t, 2H, H5 , 6.5, 0.136), 7.762 (d, 2H, H6 , 4.8, −0.451). Group III,
7.007 (d, 2H, H3 , 8.3, −0.730), 7.528 (t, 2H, H4 , 4.3, −0.108), 6.485
(t, 2H, H5 , 6.5, −0.362), 7.118 (d, 2H, H6 , 4.4, −1.095) and 10.699
(s, 2H, NH, 1.067).
Synthesis of [Ru(Hdpa)2 (5,6-dmp)](ClO4 )2 2. This was prepared by refluxing cis-[Ru(Hdpa)2 (Cl2 )]Cl (0.15 g, 0.26 mmol) and
5,6-dmp (0.054 g, 0.26 mmol) using the procedure employed for 1.
Yield: 0.18 g (81%). Anal. Calc. for RuC34 H30 N8 Cl2 O8 : C, 48.01;
H, 3.55; N, 13.17. Found: C, 48.00; H, 3.42; N, 13.13%. kmax /nm
(e/M−1 cm−1 ) (5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer
(0.5 : 9.5, v/v) buffer at pH 7.1): 505 (sh), 453 (5673), 423 (sh)
(6067), 389 (6497), 290 (31 864), 272 (51 434), 240 (33 736). ESIMS: [Ru(Hdpa)2 (5,6-dmp)]2+ displays a peak at m/z = 325.6,
calc. 325.8. 1 H NMR (DMSO-d6 , 400 MHz): d (multiplicity,
integration, assignment, J/Hz, c.i.s.)/ppm. Group I, 8.829 (d,
2H, H2 , 8.1, −0.262), 8.031 (t, 2H, H3 , 4.2, 0.474), 9.089 (d, 2H,
H4 , 5.1, 0.784), 2.781 (s, 6H, CH3 , 0.207). Group II, 7.374 (d, 2H,
H3 , 8.2, −0.363), 7.988 (t, 2H, H4 , 7.9, 0.352), 6.985 (t, 2H, H5 ,
6.2, 0.138), 7.740 (d, 2H, H6 , 4.8, −0.473). Group III, 6.991 (d,
2H, H3 , 7.5, −0.746), 7.531 (t, 2H, H4 , 7.4, −0.105), 6.479 (t, 2H,
H5 , 6.2, −0.368), 7.044 (d, 2H, H6 , 4.9, −1.169) and 10.653 (s, 2H,
NH, 1.021).
Synthesis of [Ru(Hdpa)2 (dpq)](ClO4 )2 3. This was prepared
by refluxing cis-[Ru(Hdpa)2 (Cl2 )]Cl (0.2 g, 0.35 mmol) and dpq
(0.060 g, 0.35 mmol) using the procedure employed for 1. Yield:
0.26 g (88%). Anal. Calc. for RuC34 H26 N8 Cl2 O8 : C, 46.69; H,
3.00; N, 16.02. Found: C, 46.20; H, 2.95; N, 15.96%. kmax /nm
(e/M−1 cm−1 ) (5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer
(0.5 : 9.5, v/v) buffer at pH 7.1): 522 (sh), 466 (5225), 434
(sh) (5367), 284 (46 952), 269 (6359), 257 (55 008). ESI-MS:
[Ru(Hdpa)2 (dpq)]2+ displays a peak at m/z = 337.6, calc. 337.8.
1
H NMR (DMSO-d6 , 400 MHz): d (multiplicity, integration,
assignment, J/Hz, c.i.s.)/ppm. Group I, 9.129 (d, 2H, H2 , 7.5,
−0.303), 9.123 (t, 2H, H3 , 4.4, 1.200), 9.559 (d, 2H, H4 , 7.7, 0.338),
9.374 (s, 2H, H6 , 0.240). Group II, 7.315 (d, 2H, H3 , 8.3, −0.422),
8.163 (t, 2H, H4 , 4.1, 0.527), 7.588 (t, 2H, H5 , 6.3, 0.741), 7.685 (d,
2H, H6 , 4.8, −0.528). Group III, 7.023 (d, 2H, H3 , 8.1, −0.714),
7.985 (t, 2H, H4 , 4.2, 0.349), 6.483 (t, 2H, H5 , 6.3, −0.364), 7.038
(d, 2H, H6 , 4.4, −1.175) and 10.611 (s, 2H, NH, 0.979).
Synthesis of [Ru(Hdpa)2 (mdpq)](ClO4 )2 4. This was prepared
by refluxing cis-[Ru(Hdpa)2 (Cl2 )]Cl (0.15 g, 0.26 mmol) and mdpq
(0.064 g, 0.26 mmol) using the procedure employed for 1. Yield:
0.16 g (72%). Anal. Calc. for RuC35 H28 N8 Cl2 O8 : C, 47.31; H,
3.18; N, 15.76. Found: C, 47.10; H, 3.08; N, 15.25%. kmax /nm
(e/M−1 cm−1 ) (5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer
(0.5 : 9.5, v/v) buffer at pH 7.1): 525 (sh), 463 (5895), 435
(sh) (6094), 371 (7889), 282 (52 982), 260 (61 621). ESI-MS:
[Ru(Hdpa)2 (mdpq)]2+ displays a peak at m/z = 344.6, calc. 344.8.
1
H NMR (DMSO-d6 , 400 MHz): d (multiplicity, integration,
assignment, J/Hz, c.i.s.)/ppm. Group I, 9.123 (d, 2H, H2 , 8.0,
−0.343), 9.119 (t, 2H, H3 , 4.5, 1.257), 9.536 (d, 2H, H4 , 6.3, 0.325),
9.507 (d, 2H, H4 , 6.1, 0.296), 9.274 (s, 1H, H6 , 0.238), 2.941 (s,
3H, CH3 , 0.143). Group II, 7.300 (d, 2H, H3 , 8.2, −0.437), 8.137
Dalton Trans., 2008, 2157–2170 | 2159
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(t, 2H, H4 , 5.4, 0.501), 7.581 (t, 2H, H5 , 6.4, 0.734), 7.693 (d, 2H,
H6 , 4.9, −0.520). Group III, 7.000 (d, 2H, H3 , 8.2, −0.737), 7.985
(t, 2H, H4 , 5.5, 0.349), 6.481 (t, 2H, H5 , 6.3, −0.366), 7.022 (d, 2H,
H6 , 4.6, −1.191) and 10.574 (s, 2H, NH, 0.942).
Synthesis of [Ru(Hdpa)2 (dppz)](ClO4 )2 5. This was prepared
by refluxing cis-[Ru(Hdpa)2 (Cl2 )]Cl (0.2 g, 0.35 mmol) and dppz
(0.099 g, 0.35 mmol) using the procedure employed for 1. Yield:
0.25 g (80%). Anal. Calc. for RuC38 H28 N8 Cl2 O8 : C, 49.36; H,
3.05; N, 15.15. Found: C, 49.15; H, 3.03; N, 15.05%. kmax /nm
(e/M−1 cm−1 ) (5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer
(0.5 : 9.5, v/v) buffer at pH 7.1): 479 (4279), 437 (sh) (4623), 373
(13 814), 360 (14 081), 279 (56 256). ESI-MS: [Ru(Hdpa)2 (dppz)]2+
displays a peak at m/z = 362.6, calc. 362.9. 1 H NMR (DMSOd6 , 400 MHz): d (multiplicity, integration, assignment, J/Hz,
c.i.s.)/ppm. Group I, 9.081 (d, 2H, H2 , 8.0, −0.376), 8.641 (t,
2H, H3 , 4.5, 0.736), 9.643 (d, 2H, H4 , 5.3, 0.461), 8.226 (d, 2H,
H7 , 5.8, −0.103), 8.143 (t, 2H, H8 , 6.3, 0.113). Group II, 7.326 (d,
2H, H3 , 8.2, −0.411), 7.987 (t, 2H, H4 , 4.4, 0.351), 7.043 (t, 2H,
H5 , 6.4, 0.196), 7.678 (d, 2H, H6 , 4.5, −0.535). Group III, 7.074
(d, 2H, H3 , 8.2, −0.663), 7.613 (t, 2H, H4 , 4.5, −0.023), 6.513 (t,
2H, H5 , 6.5, −0.334), 7.109 (d, 2H, H6 , 4.8, −1.104) and 10.686 (s,
2H, NH, 1.054). ESI-MS: [Ru(Hdpa)2 (dppz)]2+ displays a peak at
m/z = 362.6, calc. 362.9.
X-Ray crystallography
A crystal of 1 of suitable size was selected from the motherliquor and immersed in paraffin oil, then mounted on the tip
of a glass fiber and cemented using epoxy resin. Intensity data
for crystal was collected using Mo-Ka (k = 0.71073 Å) radiation
on a Bruker SMART APEX diffractometer equipped with CCD
area detector at 100 K. The data integration and reduction
was processed with SAINT24 software. An empirical absorption
correction was applied to the collected reflections with SADABS.25
The structure was solved by direct methods using SHELXTL26 and
was refined on F 2 by the full-matrix least-squares technique using
the SHELXL-9727 program package. All non-hydrogen atoms
were refined anisotropically till convergence is reached. Hydrogen
atoms attached to the ligand moieties are stereochemically fixed.
The crystallographic data and details of data collection for 1 is
given in Table 1. Graphical representations of the structure were
made with POV-Ray v3.6.28
DNA binding experiments
Concentrated stock solutions of metal complexes were prepared
by dissolving them in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl
buffer (0.5 : 9.5, v/v) buffer at pH 7.1 of metal complexes and
diluting suitably with corresponding buffer to required concentrations for all the experiments. For absorption and emission
spectral experiments the DNA solutions were pretreated with the
solutions of metal complexes to ensure no change in concentration
of the metal complexes. The absorption spectra were recorded on
a Varian Cary 300 Bio UV-Vis spectrophotometer using cuvettes
of 1 cm path length.
Absorption spectral titration experiments were performed by
maintaining a constant concentration of the complex and varying
the nucleic acid concentration. This was achieved by dissolving
an appropriate amount of the metal complex and DNA stock
2160 | Dalton Trans., 2008, 2157–2170
Table 1 Crystal data and structure refinement details for 1
Empirical formula
Mr
Crystal system
Space group
a/Å
b/Å
c/Å
V /Å3
Z
Dc /g cm−3
l/cm−1
k(Mo-Ka)/Å
T/K
F(000)
Reflection collected
Independent reflections
Rint
Goodness-of-fit on F 2
Ra (I ≥ 2r(I))
wRb (I ≥ 2r(I))
a
R =
F o | − |F c / |F o |.
2 1/2
[w(|F o |) ]} .
RuC32 H25 N8 Cl2 O8
821.57
Orthorhombic
Pbca
18.642(10)
15.952(9)
22.343(12)
6644(6)
8
1.643
6.98
0.71073
293(2)
3320
24624
4333
0.2366
1.057
0.0936
0.2158
b
R = (
[w(|F o | − |F c |)2 /
solutions while maintaining the total volume constant (1 mL).
This results in a series of solutions with varying concentrations
of DNA but with a constant concentration of the complex. The
absorbance (A) was recorded after successive additions of CT
DNA.
For viscosity measurements a Schott Gerate AVS 310 automated
viscometer was thermostated at 25 ± 1 ◦ C in a constant temperature bath. DNA concentration was kept constant (200 lM in
NP) and the concentration of metal complexes varied (1/R =
[Ru]/[DNA] = 0.0–0.50). The flow times were noted from the
digital timer attached with the viscometer. Data are presented as
g/g0 vs. 1/R, where g is the viscosity of DNA in the presence of
the ruthenium(II) complex and g0 is the relative viscosity of DNA
alone. Relative viscosity values were calculated from the observed
flow time of DNA solution (t) corrected for the flow time of buffer
alone (t0 ), using the expression g0 = (t − t0 )/t0 .
Circular dichroic spectra of DNA were obtained by using
JASCO J-716 spectropolarimeter equipped with a Peltier temperature control device. All experiments were done using a quartz
cell of 1 cm path length for DNA or 0.2 cm path length for
oligonucleotides. Each CD spectrum was collected after averaging
over at least four accumulations using a scan speed of 100 nm min−1
and a 1 s response time. Machine plus cuvette baselines were
subtracted and the resultant spectrum zeroed 50 nm outside the
absorption bands. Emission intensity measurements were carried
out using Jasco F 6500 spectrofluorimeter. The tris buffer was
used as a blank to make preliminary adjustments. The excitation
wavelength was fixed and the emission range was adjusted before
measurements. DNA was pretreated with ethidium bromide in the
ratio [NP/EthBr] = 10 for 30 min at 27 ◦ C. The metal complexes
(1/R = [metal]/[DNA] = 10) were then added to this mixture
and their effect on the emission intensity was measured. Cyclic
voltammetry (CV) and differential pulse voltammetry (DPV) were
performed in a single compartment cell with a three-electrode
configuration on a EG & G PAR 273 potentiostat-galvanostat
equipped with an PIV computer. The working electrode was a
glassy carbon disk (0.384 cm2 ) and the reference electrode a
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saturated calomel electrode. A platinum plate was used as the
counter electrode. The supporting electrolyte was 50 mM NaCl–
5 mM Tris-HCl buffer at pH 7.1. Solutions were deoxygenated
by purging with nitrogen gas for 15 min prior to measurements;
during measurements a stream of N2 gas was passed over the
solution. All the experiments were carried out at 25.0 ± 0.2 ◦ C
maintained by a Haake D8-G circulating bath. The redox potential
E 1/2 was calculated from the anodic (E pa ) and cathodic (E pc ) peak
potentials of CV traces as (E pa + E pc )/2 and also from the peak
potential (E pa ) of DPV response as E p + DE/2 (DE is the pulse
height).
The interaction of complexes with supercoiled pBR322 DNA
was monitored using agarose gel electrophoresis. In reactions
using supercoiled pBR322 plasmid DNA (Form I, 40 lM) in 5%
CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer (0.5 : 9.5, v/v)
at pH 7.1 was treated with metal complexes in the same buffer.
For photocleavage studies, the reactions were carried out under
illuminated conditions at 365 nm (12 W) monochromatic light
source. In each experiment supercoiled pBR322 DNA (Form I,
40 lM) was treated with metal complexes (60 lM) and irradiated
at 365 nm monochromatic wavelength for 5 min. The samples
were then incubated for 1 h in the dark for 37 ◦ C and analysed for
the photocleaved products using gel electrophoresis as discussed
below. A loading buffer containing 25% bromophenol blue,
0.25% xylene cyanol and 30% glycerol (3 ll) was added and
electrophoresis performed at 60 V for 5 h in Tris–acetate–EDTA
(TAE) buffer (40 mM Tris-base, 20 mM acetic acid, 1 mM EDTA)
using 1% agarose gel containing 1.0 lg mL−1 ethidium bromide.29
The gels were viewed in a Gel doc system and photographed using
a CCD camera (Alpha Innotech Corporation).
Cell viability assay
MTT assay was carried out as described previously.30 The complexes 1–5, in the concentration 0.05–100 lM, dissolved in DMSO
(Sigma-Aldrich, St. Louis, MO, USA), were added to the wells 24 h
after seeding of 5 × 103 cells per well in 200 lL of fresh culture
medium. DMSO was used as the vehicle control. After 24 and
48 h, 20 lL of MTT solution [5 mg/mL in phosphate-buffered
saline (PBS)] was added to each well and the plates were wrapped
with aluminum foil and incubated for 4 h at 37 ◦ C. The purple
formazan product was dissolved by addition of 100 lL of 100%
DMSO to each well. The absorbance was monitored at 570 nm
(measurement) and 630 nm (reference) using a 96 well plate reader
(Bio-Rad, Hercules, CA, USA). The stock solutions of the metal
complexes were prepared in DMSO and in all the experiments
the percentage of DMSO was maintained in the range of 0.1–
1%. DMSO by itself was found to be non-toxic to the cells till
1% concentration. Data were collected for four replicates each
and used to calculate the mean. The percentage inhibition was
calculated, from this data, using the formula:
The IC50 values were calculated using Table Curve 2D version
5.01.
This journal is © The Royal Society of Chemistry 2008
Hoechst 33258 staining
Cell pathology was detected by staining the nuclear chromatin
of trypsinized cells (4.0 × 104 ml−1 ) with 1 ll of Hoechst 33258
(1 mg ml−1 , aqueous) for 10 min at 37 ◦ C. Protocol: Staining of
suspension cells with Hoechst 33258 to detect apoptosis.31 A drop
of cell suspension was placed on a glass slide and a cover-slip was
laid over to reduce light diffraction. At random 300 cells were
observed in a fluorescent microscope (Carl Zeiss, Jena, Germany)
fitted with a 377–355 nm filter, and observed at ×400 magnification
and the percentage of cells reflecting pathological changes were
calculated. Data were collected for four replicates and used to
calculate the mean and the standard deviation.
Results and discussion
Synthesis and characterisation of Ru(II) complexes
The mixed ligand complexes [Ru(Hdpa)2 (diimine)](ClO4 )2
1–5, where Hdpa is 2,2 -dipyridylamine and diimine is 1,10phenanthroline (phen) (1), 5,6-dimethyl-1,10-phenanthroline
(5,6-dmp) (2), dipyrido[3,2-d:2 ,3 -f ]quinoxaline (dpq) (3),
5-methyldipyrido[3,2-d:2 ,3 -f ]quinoxaline (mdpq) (4) and
dipyrido[3,2-a:2 ,3 -c]phenazine (dppz) (5) have been isolated by
reacting the complex [Ru(Hdpa)2 Cl2 ]Cl with the corresponding
diimine ligands. The CHN analyses of the complexes were
consistent with the formula [Ru(Hdpa)2 (diimine)](ClO4 )2 . The
complexes are soluble in both polar and non-polar solvents. The
perchlorate salts of 3–5 are highly soluble in water but those of 1
and 2 are soluble in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl
buffer (0.5 : 9.5, v/v) at pH 7.1. So solutions of the all complexes
were prepared in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl
buffer at pH 7.1 for DNA binding and other studies.
Description of the structure of [Ru(Hdpa)2 (phen)](ClO4 )2 (1).
The ball-and-stick representation of the structure of the cation
[Ru(Hdpa)2 (phen)]2+ of complex 1 is illustrated in Fig. 1 with
atom numbering scheme. The crystallographic data are given in
Fig. 1 Ball-and-stick representation of the crystal structure of
[Ru(Hdpa)2 (phen)]2+ cation 1; atoms as spheres of arbitrary diameter.
Hydrogen atoms are omitted for clarity.
Dalton Trans., 2008, 2157–2170 | 2161
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Table 2 Selected interatomic distances (Å) and angles (◦ ) for 1
Ru(1)–N(1)
Ru(1)–N(2)
Ru(1)–N(3)
2.086(10)
2.084(9)
2.073(11)
Ru(1)–N(4)
Ru(1)–N(5)
Ru(1)–N(6)
2.051(11)
2.072(11)
2.071(11)
N(1)–Ru(1)–N(2)
N(1)–Ru(1)–N(3)
N(1)–Ru(1)–N(5)
N(1)–Ru(1)–N(6)
N(1)–Ru(1)–N(8)
N(2)–Ru(1)–N(3)
N(2)–Ru(1)–N(5)
N(2)–Ru(1)–N(6)
79.7(4)
88.5(4)
175.0(4)
93.6(4)
74.1(4)
89.7(4)
96.4(4)
90.3(4)
N(2)–Ru(1)–N(8)
N(3)–Ru(1)–N(5)
N(3)–Ru(1)–N(6)
N(3)–Ru(1)–N(8)
N(5)–Ru(1)–N(6)
N(5)–Ru(1)–N8
N(6)–Ru(1)–N8
173.0(5)
88.5(5)
177.9(5)
93.3(5)
89.4(4)
89.9(4)
86.9(5)
Table 1 while the selected bond lengths and bond angles are given
in Table 2. The complex cation possesses a distorted octahedral
coordination geometry constituted by two dipyridylamine (Hdpa)
and 1,10-phenanthroline ligands chelated to Ru(II). The presence
of distorted coordination geometry around Ru(II) is evident from
the values of the bond angles N1–Ru1–N2 [79.6(4)], N1–Ru1–
N3 [88.5(4)], N6–Ru1–N8 [87.0(5)], N3–Ru–N6 [177.9(5)], N2–
Ru1–N8 [173.0(6)] and N1–Ru1–N5 [175.0(4)◦ ], which deviate
from the ideal bond angles of 90◦ and 180◦ . The pyridyl rings
of the coordinated Hdpa ligands are non-planar to each other.
The least squares planes of the pyridyl rings of the first Hdpa
ligand coordinated to the metal center shows an interplanar
angle (N3–C13 to C17 and N5–C19 to C22) of 35.19◦ , which is
higher than that (39.66◦ ) between the pyridyl rings of the second
Hdpa ligand (N6–C23 to C27 and N8–C28 to C32) suggesting
that the first Hdpa ligand makes a more effective coordination
than the second one and that the steric crowding of ligands is
significant. The flexible non-planar Hdpa ligands approach Ru(II)
more closely and r-coordinates to Ru(II) more strongly than the
rigid 1,10-phenanthroline ligand. A similar observation has been
made earlier.32 The Ru–NHdpa and Ru–Nphen bond distances are very
similar to those observed in other Hdpa and phen complexes.33,34
1
H NMR spectra
The 1 H NMR spectra35,36 of 1–5 contain mainly eleven distinct
resonances (six doublets and five triplets), which could be classified
into three groups (group I, H2 –H4 , group II and III, H3 –H6 ;
Scheme 1 and Fig. S1, ESI†). The group I has three resonances
which arise from coupling of H2 , H3 and H4 protons of phen
ring with each other. Similarly, the groups II and III have each
four resonances, which arise from coupling of H3 , H4 , H5 and H6
protons of Hdpa with each other. When coordinated to Ru(II) the
planar phen, 5,6-dmp, dpq, mdpq and dppz rings are expected to
form a five-membered chelate ring with an envelope conformation
and the non-planar Hdpa ligands form six-membered chelate
rings32 with a boat conformation, as observed in the X-ray crystal
structure of 1. A combination of conformations of the two Hdpa
ligands gives four cases as shown in Fig. 2. For cases 2 and 3,
the chemical shifts should be different among the two halves of
the phen ring and also among the four pyridine rings of Hdpa
ligand because of absence of flapping of the pyridine rings of
the ligand. On the other hand, for cases 1 and 4 three groups of
magnetic environments – two equivalent halves of phen and four
pyridine rings of Hdpa – would arise because of flapping of the
py rings of Hdpa and hence the presence of a C 2 symmetry. The
2162 | Dalton Trans., 2008, 2157–2170
Fig. 2 The numbering of the phen and pyridine rings of Hdpa ligands and four cases due to the conformation of the Hdpa ligands in
[Ru(Hdpa)2 (phen)]2+ (1)
observed 1 H NMR spectral pattern is consistent with cases 1 and
4, suggesting the presence of C 2 symmetry and hence the rapid
flapping of the py rings of Hdpa ligand. Also, the X-ray crystal
structure of 1 reveals no steric interaction between H6 and H6
protons of the Hdpa ligands occupying cis positions and supports
the presence of rapid flapping of Hdpa ligands with no inhibition
from ligand crowdedness. Thus, the two halves of phen/5,6dmp/dpq/mdpq/dppz ring (R1 and R1 ) in 1–5 with an apparent
C 2 symmetry become equivalent and also the two pyridine rings
of Hdpa1-R2 and Hdpa1-R3 become equivalent to Hdpa2-R2
and Hdpa2-R3, respectively. However, the current-loop model
calculation37 of aromatic ring-current shifts of [Ru(Hdpa)2 (bpy)]2+
indicates that the flapping of Hdpa ligands makes each proton of
the complex experience an averaged magnetic environment among
cases 1–4.
The chemical shifts of each proton in 5,6-dmp (2) or dpq (3) or
mdpq (4) or dppz (5) are related (Fig. S2B, ESI†) to that of phen (1)
whereas the proton signals of Hdpa (2–5) are in good agreement
(Fig. S2C and S2D, ESI†) with those of Hdpa in 1. Judging by
the similarity of the patterns with the homoleptic complexes38
[Ru(phen)3 ]2+ and [Ru(Hdpa)3 ]2+ as well as 1 (Fig. S2A, ESI†),
group I can be assigned to R1 of the phen or 5,6-dmp or dpq
or mdpq or dppz ligand, while groups II and III are assigned to
either R2 or R3 of Hdpa ligands. Apart from the three main
resonances of group I, the complexes 1, 2 and 3 show one
individual resonance for the benzene ring (H5 ) of phen, dimethyl
substituted benzene ring (CH3 ) of 5,6-dmp and quinoxaline ring
(H6 ) of dpq. Similarly, 4 and 5 exhibit two individual resonances
for the methyl substituted quinoxaline ring (H6 and CH3 ) of mdpq
and phenazine ring (H7 and H8 ) of dppz.
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The complexes 1–5 exhibit negative c.i.s. values for H2 (group I)
and H6 (group II and III) protons adjacent to the coordinated
nitrogen atoms, which result from through space ring-current
anisotropy effects upon coordination, with the protons lying
directly over the shielding plane of another aromatic pyridine ring.
Also, upon coordination to Ru(II), the signals of H7 (5; group I),
H3 (1–5; group II and III), H4 (1, 2, 5; group III) and H5 (1–5;
group III) protons are shifted upfield due to Ru(II)-to-ligand pback donation. The positive c.i.s. values observed for H3 and H4
(1–5; group I), H5 (1, 2; group I), H6 (3, 4; group I), H8 (5; group
I), H4 and H5 (1–5; group II) and H4 (3, 4; group III) protons
arise from a r-effect based on electron donation to Ru(II) via the
nitrogen lone pairs.
Interaction of Ru(II) complexes with DNA
Absorption spectral studies
In 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1
the complexes 1–5 display a higher energy band in the range 230–
380 nm, which arises from intra-ligand p–p* type transitions39
involving ligand energy levels higher in energy than the ligand
LUMO levels. The MLCT bands for 1–5 all appear in the region
of the spectrum typical for Ru(II) complexes with coordinated
polyimine ligands. The MLCT band energy decreases from phen
complex 1 (451 nm) to dppz complex 5 (484 nm) with 33 nm redshifted, as the diimine ligand is replaced by one with enhanced
p-delocalization revealing stabilization of the p* (diimine-based
LUMO).40 The methyl substitution on phen co-ligand would be
expected to build a negative charge on metal by r-donating methyl
groups to destabilize the p* (diimine-based LUMO), but it is not
observed in the dmp complex 2. However, methyl substitution on
dpq co-ligand destabilizes the p* orbital (diimine-based LUMO)
and decreases (3 nm) the metal dp(RuII ) → p* MLCT band energy
relative to the analogous dpq complex 3. Except 5, the complexes
1–4 display a new lower energy band in the range 505–525 nm,
which arises from inter-ligand nHdpa –p*diimine .39 The complexes 1–5
exhibit no detectable luminescence in 5% CH3 OH–5 mM TrisHCl–50 mM NaCl buffer at pH 7.1.
Upon the addition of calf thymus (CT) DNA to 1–5 (R =
[NP]/[Ru complex] = 0–25) in 5% CH3 OH–5 mM Tris-HCl–
50 mM NaCl buffer at pH 7.1 at 25 ◦ C interesting changes in
intensity of the intense intraligand (IL) absorption bands (264–
387 nm) and metal-to-ligand (pM → pL *) charge transfer (MLCT)
band of the complexes in the visible region (453–484 nm, Table 3)
are observed. All the complexes exhibit uniform hypochromism
at lower DNA concentrations but hyperchromism41 at higher
Fig. 3 Effect of addition of DNA on the absorption intensity of the
complexes 1, 453; 2, 453; 3, 446; 4, 464; 5, 484 nm in a 5% CH3 OH–5 mM
Tris-HCl–50 mM NaCl buffer at pH 7.1 and 25 ◦ C.
DNA concentrations revealing that the DNA binding modes
of all the complexes are similar (Fig. 3). It appears that the
complexes undergo distortion in the coordination sphere at higher
DNA concentrations resulting in enhanced allowedness of the
intraligand as well as MLCT bands. The mixed spectral behaviour
reveals the presence of more than one DNA binding mode for the
complexes. Also, while the well defined MLCT bands observed for
1–4 were used to monitor absorption titration of the complexes,
the intraligand transition was used for 5. So no attempt was
made to quantitatively compare the DNA binding affinities of the
complexes. The higher hypochromism exhibited by 5 is obviously
due to involvement of the extended aromatic ring of dppz ligand
in intercalative interaction with the DNA base pairs. Also, as
expected, the methyl substituted 5,6-dmp and mdpq complexes 2
and 4 show lower hypochromism than their analogous phen (1)
and dpq (3) complexes (cf . below).
Ethidium displacement assay
All the present complexes fail to show steady-state emission in
5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1
at 25 ◦ C and lack emission even in the presence of CT DNA
(R = 25). So the extent of DNA binding of these complexes
has been evaluated using competitive binding studies involving
ethidium bromide (EthBr), which is known to emit strongly (kex ,
450; kem , 595 nm) due to its intercalative interaction with DNA. On
adding the complexes 1–5 to DNA (R = 1) pretreated with EthBr
([DNA]/[EthBr]) = 1) the emission intensity decreases (Fig. 4).
The observed fluorescence intensities of DNA-bound ethidium
bromide were plotted against complex concentration and the
Table 3 Absorption spectral properties of Ru(II) complexes bounda to CT DNA
Complex
kmax /nm
R
Change in absorbance
De (%)
Red shift/nm
[Ru(Hdpa)2 (phen)]2+ 1
[Ru(Hdpa)2 (5,6-dmp)]2+ 2
[Ru(Hdpa)2 (dpq)]2+ 3
[Ru(Hdpa)2 (mdpq)]2+ 4
[Ru(Hdpa)2 (dppz)]2+ 5
453
453
446
464
484
25
25
25
25
25
Hypo- and hyperchromism
Hypo- and hyperchromism
Hypo- and hyperchromism
Hypo- and hyperchromism
Hypo- and hyperchromism
6
5
18
11
14
5
2
1
9
2
Measurement were made at R = 25, where R = [DNA]/[Ru complex], concentration of Ru(II) complex solutions = 1 × 10−4 M (1, 2 and 4) and 5 ×
10−5 M (3 and 5).
a
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Circular dichroic spectral studies
Fig. 4 Effect of addition of 1–5 on the emission intensity of the CT
DNA-bound ethidium bromide (125 lM) at different complex concentrations in a 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1 and
25 ◦ C.
values of apparent DNA binding constant (K app ) were calculated42
using the relation
K EthBr [EthBr] = K app [Complex]
where,42 K EthBr is 1.0 × 107 M−1 , the concentration of EthBr is
12.5 lM and the concentration of the complex is that used to
obtain a 50% reduction of fluorescence intensity of EthBr. The
DNA binding abilities of the complexes follow the order, 5 dppz
(K app , 9.78 × 107 M−1 ) > 3 dpq (5.84 × 107 M−1 ) > 4 mdpq (5.00 ×
107 M−1 ) > 2 dmp (4.77 × 107 M−1 ) > 1 phen (0.97 × 107 M−1 )).
Both the electron transfer from excited EthBr to ruthenium(II) and
the EthBr displacement mechanisms would account for the highest
K app value of the dppz complex (cf. below) and hence its DNA
binding affinity as well (cf . above). The dppz complex 5, which
is involved in a strong DNA intercalation, would compete with
the intercalatively bound EthBr for DNA binding and quench the
EthBr emission to a greater extent (100%) than other complexes.
It appears that the DNA helix simultaneously accommodates
both the complex and EthBr in the grooves and the enhanced
hydrophobicity of the fused benzene ring in DNA-bound dppz
complex perturbs the DNA helix more than the other complexes,
to displace the bound EthBr more efficiently. This is attributed
to the intramolecular photoinduced electron transfer exhibited
by the flanking phenazine moiety in DNA-bound or free dppz
complex quenches the emission of the DNA-bound EthBr.43 The
phen complex 1, which is involved in partial DNA intercalation,
would be expected to show a K app value higher than 2. However, 2
shows a K app value higher than 1 indicating that the hydrophobic
interaction34 of 2 with DNA involving 5,6 methyl groups rather
than the partial intercalation of phen ring is more important in
determining the EthBr quenching. The complex 4 displays a K app
value lower than 3 even though it possesses one methyl group
on the quinoxaline moiety. So two adjacent methyl groups are
needed to place them effectively in the grooves and displace the
DNA-bound EthBr.44 The dicationic complex 5 is expected to
be involved in strong partial intercalation with DNA and compete
with the intercalatively bound monopositive ethidium cation more
strongly than other complexes. We propose that hydrogen bonding
interactions, which would occur between the–NH- of Hdpa in 5
and functional groups present on the edge of the DNA,1c would
also contribute significantly to the higher DNA binding affinity
of 5.
2164 | Dalton Trans., 2008, 2157–2170
Circular dichroic spectra provide information about the chirality
of spectroscopically active species in solution and this is a useful
technique in diagnosing changes in DNA morphology during
drug–DNA interactions, as the band due to base stacking (275 nm)
and that due to right-handed helicity (248 nm) are quite sensitive to
the mode of DNA interactions with small molecules.45 The changes
in CD signals of DNA observed on interaction with drugs may often be assigned to the corresponding changes in DNA structure.46
Thus simple groove binding and electrostatic interaction of small
molecules show less or no perturbation on the base-stacking
and helicity bands, while intercalation enhances the intensities
of both the bands stabilizing the right-handed B conformation
of CT DNA as observed for the classical intercalator methylene
blue.47 Thus rac-metal complexes give a zero CD but show induced
circular dichroic (ICD) signals on enantiopreferential binding to
DNA providing further and definitive confirmation for their DNA
binding.48,49 Thus, the CD spectral technique has been used to
study the enantiopreferential DNA binding of the present racmetal complexes. Also, the technique is useful in diagnosing
changes in DNA morphology during drug-DNA interaction as
CD signals are quite sensitive to the mode of DNA interactions of
small molecules.43,50
When the present rac-complexes are incubated with CT DNA
at 1/R (= [Ru complex]/[DNA]) value of 1, the CD spectrum of
DNA undergoes changes in both the positive and negative bands
(Table 4). Upon adding complex 1 slight changes in intensities
of the negative and positive bands, and shifts in band positions
are observed. This reveals that 1 merely binds with DNA electrostatically. Interestingly, 2 shows a sharp B to W conformational
change (Fig. 5), which is similar to that observed for [Co(NH3 )6 ]3+
bound to DNA of short lengths with 160 base pairs.51 On the
other hand, upon addition of rac complexes 3 (Fig. 6), 4 (Fig. 7)
and 5 (Fig. 8) to DNA both these bands disappear and an
inverted CD signal is observed with intensity much higher than
free DNA. The latter consists of a positive (262 nm) and a
negative band (282 nm) with a zero cross-over at 272 nm. This is
typical of exciton coupled ICD arising due to enantiopreferential
binding of the D-enantiomer of the rac-complex and/or ligand–
ligand interactions among DNA bound/unbound complexes.
The complexes 3–5 which exhibits ICD interact with different
polynucleotides and the binding efficiency varies as CT DNA >
poly(AT)12 ∼
= d(CGCGATCGCG)2 > poly(GC)12 (Fig. 6, 7 and
Table 4 CD parameters for the interaction of calf thymus DNA with
complexes 1–5
Sample
CD spectral band,a k/nm
50 lM CT DNA
DNA + 5 lM [Ru(Hdpa)2 (phen)]2+ 1
DNA + 5 lM [Ru(Hdpa)2 (5,6-dmp)]2+ 2
DNA + 5 lM [Ru(Hdpa)2 (dpq)]2+ 3
DNA + 5 lM [Ru(Hdpa)2 (mdpq)]2+ 4
DNA + 5 lM [Ru(Hdpa)2 (dppz)]2+ 5
245
245
245
241, 286
239, 286
243,289
276
269
267, 298
262
263
264
a
Measurement made at 1/R value of 1 for complexes 1–5 where 1/R =
[Ru]/[NP]; concentration of DNA solutions = 5 × 10−5 M. Cell path
length = 1 cm.
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Fig. 5 (A) Circular dichroism spectra of CT DNA in the absence
(a) and presence of [Ru(Hdpa)2 (phen)]2+ (b) at 1/R = 1. (B) Circular
dichroism spectra of CT DNA in the absence (a) and presence of
[Ru(Hdpa)2 (5,6-dmp)]2+ (b) at 1/R = 1. [DNA] = 2 × 10−5 M.
8). These observations reveal the preferential binding of 3–5 to AT
and mixed rather than GC sequences.
Viscometry studies
The hydrodynamic changes of DNA biopolymer are the consequence of lengthening of the molecule, the diminished bending
between layers, and the diminished length-specific mass.52 The
DNA viscosity is enhanced significantly due to complete or partial
intercalation of drugs into the DNA base stacking but it is slightly
disturbed by electrostatic or covalent binding of molecules. When
1–5 are treated with CT DNA (200 lM) and their concentration
is increased from 1/R = 0–0.5 (1/R = [Ru]/[DNA]), the relative
viscosity of CT DNA increases (Fig. 9), which follows the order
5 > 3 > 4 > 2 > 1. These results parallel the K app values observed
for the complexes and illustrate the lengthening of DNA duplex
upon intercalation of the extended planar aromatic ligands of
the complexes on DNA binding. The highest increase in DNA
viscosity effected by 5 is similar to those observed for proven
intercalators, and can be explained by a model used for the DNA
binding of acridine orange and proflavine.53 Also, incorporation
of a methyl group on dpq in 3 decreases the partial intercalation of
dpq leading to a lower increase in viscosity of DNA upon binding
to 4. On the other hand, the incorporation of two methyl group
on phen ring as in 5,6-dmp leads to a higher increase in viscosity
revealing the importance of hydrophobic interaction of 2 in DNA
grooves in enhancing the length of DNA.
Electrochemical behaviour of complexes in the absence and
presence of DNA
In 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer solution
at pH 7.1 the complexes 1 and 2 exhibit a quasi-reversible
Ru(II)/Ru(III) redox wave (DE p , 98 mV) while 3–5 exhibit irreversible (DE p , 172–232 mV) response (Table 5) with the E 1/2 values
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Fig. 6 (A) Circular dichroism spectra of CT DNA in the absence (a) and
presence of [Ru(Hdpa)2 (dpq)]2+ at 1/R = 1; [DNA] = 2 × 10−5 M. (B)
poly(GC)12 , (C) poly(AT)12 and (D) d(CGCGATCGCG)2 in the absence
(a) and presence of [Ru(Hdpa)2 (dpq)]2+ (b) at 1/R = 4; poly(GC)12 ,
poly(AT)12 and d(CGCGATCGCG)2 = 1 × 10−5 M.
falling in the range of 0.555–0.630 V. In addition, all the complexes
exhibit one more metal-based oxidative response in DPV with E 1/2
values falling in the range of 0.789–1.040 V. This is attributed to the
Ru(II)/Ru(III) couple of deprotonated mixed-ligand complexes54
[Ru(dpa)2 (diimine)] as illustrated in Fig. 14 and Table 5. The
low Ru(II)/Ru(III) redox potentials of the present chelates may
be attributed to the strong r-donating and less p-accepting ability
of Hdpa ligand to stabilize the higher Ru(III) oxidation state. The
formal potentials of the Ru(II)/Ru(III) couple E ◦ (or voltammetric
E 1/2 ) follows the order 5 > 3 > 4 > 1 > 2, which is in agreement with
the MLCT band energies of the complexes (cf. above). The number
of methyl groups and extended aromatic rings in the diimine
ligands dictate the E 1/2 value and thus, incorporation of electronrepelling methyl groups on phen ring as in the 5,6-dmp complex
2 renders the oxidation of Ru(II) to Ru(III) facile leading to a E 1/2
values lower than the phen complex 1. Similarly, the mdpq complex
4 exhibits the Ru(II)/Ru(III) redox couple at a potential lower than
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Fig. 7 (A) Circular dichroism spectra of CT DNA in the absence (a) and
presence of [Ru(Hdpa)2 (mdpq)]2+ at 1/R = 1; [DNA] = 2 × 10−5 M. (B)
poly(GC)12 , (C) poly(AT)12 and (D) d(CGCGATCGCG)2 in the absence
(a) and presence of [Ru(Hdpa)2 (mdpq)]2+ (b) at 1/R = 4; poly(GC)12 ,
poly(AT)12 and d(CGCGATCGCG)2 = 1 × 10−5 M.
the dpq complex 3. The Ru(II)/Ru(III) potentials become more
positive as the diimine ligand is replaced by one with enhanced pdelocalization revealing the higher stabilization of the lower Ru(II)
oxidation state, which is consistent with the decrease in MLCT
band energy. All the complexes display one or two irreversible
reduction waves in the potential range −0.349 to −1.559 V, which
arise from addition of electrons in the electrochemically accessible
LUMO55 of the coordinated co-ligands rather than that of the
Hdpa ligand.
The application of electrochemical methods to monitor the
binding of metal complexes to DNA provides useful complements
to the above methods of investigations such as UV-Vis and CD
spectroscopy.56 The cyclic (CV) and differential pulse voltammetric
(DPV) responses have been obtained for 1–5 in 5% CH3 OH–
5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1 in the presence
of DNA also and the well-behaved DPV responses are used to
monitor the interaction of complexes with DNA (Table 5). On
the incremental addition of CT DNA to the metal complexes
2166 | Dalton Trans., 2008, 2157–2170
Fig. 8 (A) Circular dichroism spectra of CT DNA in the absence (a) and
presence of [Ru(Hdpa)2 (dppz)]2+ at 1/R = 1; [DNA] = 2 × 10−5 M. (B)
poly(GC)12 , (C) poly(AT)12 and (D) d(CGCGATCGCG)2 in the absence
(a) and presence of [Ru(Hdpa)2 (dppz)]2+ (b) at 1/R = 4; poly(GC)12 ,
poly(AT)12 and d(CGCGATCGCG)2 = 1 × 10−5 M.
(R = 0.25–3), an increase in peak currents of the anodic waves
in the CV and DPV responses for 1–5 (Fig. 10) is observed
at lower DNA concentrations. In contrast, at higher DNA
concentrations the value of ipa decreases. This indicates that
multiple turnovers of oxidation of DNA by the oxidized Ru(III)
form of the metal complexes occur during a single voltammetric
sweep at lower DNA concentrations. The drop in peak currents
at higher concentrations of DNA reveal the slower mass transfer
of the complexes bound to DNA fragments, which leads to a
decrease in concentration of the unbound redox-active species
in solution. Similar catalytic enhancement of the anodic current
has been observed on ITO electrode when complexes such as
[Ru(diimine)3 ]2+ , where diimine = bpy, phen and 5,6-dmp,33,57
are interacted with CT DNA. A summary of the voltammetric
results is given in Table 5 and the typical CV responses of the dpq
complex in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer in
the presence and absence of CT DNA are shown in Fig. 10. It is
This journal is © The Royal Society of Chemistry 2008
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Table 5 Cyclic voltammetric behavioura of Ru(II) complexes in the absence and presence of DNA at 25.0 ± 0.2 ◦ C
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E 1/2 /V
Complex
R
E pa /V
E pc /V
CV
DPV (I)
DPV (II)
DE p /mV
106 D/cm2 s−1
[Ru(Hdpa)2 (phen)]2+ 1
0.00
3.00
0.631
0.647
0.533
0.547
0.582
0.597
0.580
0.594
−0.349b
1.040
1.053
—
98
100
7.0
[Ru(Hdpa)2 (5,6-dmp)]2+ 2
0.00
3.00
0.620
0.620
0.522
0.528
0.571
0.574
0.577
0.575
−1.311b
0.798
0.799
−1.559b
98
92
0.14
[Ru(Hdpa)2 (dpq)]2+ 3
0.00
3.00
0.696
0.710
0.556
0.522
0.626
0.616
0.637
0.667
−0.429b
0.813
0.839
−0.987b
232
188
5.7
[Ru(Hdpa)2 (mdpq)]2+ 4
0.00
3.00
0.672
0.704
0.550
0.500
0.611
0.602
0.613
0.681
−0.419b
0.811
0.816
−1.021b
122
204
7.3
[Ru(Hdpa)2 (dppz)]2+ 5
0.00
0.25
0.730
0.696
0.558
0.600
0.644
0.648
0.655
0.637
−0.557b
0.829
0.823
−0.757b
172
96
0.12
Measured vs. saturated calomel electrode; add 0.244 to convert to normal hydrogen electrode (NHE); scan rate 50 mV s−1 ; supporting electrolyte 50 mM
NaCl; complex concentration 0.25 × 10−3 ; differential pulse voltammetry (DPV), scan rate 2 mV s−1 ; pulse height 50 mV; R = [DNA]/[Ru(II) complex].
b
Ligand reduction potential.
a
Fig. 9 The effect of addition of 1–5 on the viscosity of CT DNA in 5%
CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1; relative specific
viscosity vs. 1/R; [CT DNA] = 200 lM.
obvious that at higher concentrations of DNA the strongly DNAbound Ru(III) species are immobilized on DNA rendering them
incapable of moving to other parts of the biopolymer and effect
oxidation of guanine of DNA. Further, the shift in E 1/2 of 1–4 to
a more positive potential (4–70 mV) on binding to DNA reveals
that the oxidation of Ru(II) to Ru(III) is rendered difficult. On the
other hand, the E 1/2 of 5 becomes less positive (20 mV) on binding
to DNA revealing that the oxidation of Ru(II) to Ru(III) becomes
facile. Thus the DNA surface-bound complexes 1 and 2 exhibit
guanine oxidation and in fact it has been previously reported that
DNA surface-bound complexes such as [Ru(diimine)3 ]2+ exhibit
electrocatalytic oxidation of guanine.33,57 However, the complexes
3, 4 and 5 bound to DNA through partial intercalation are involved
in electrocatalytic guanine oxidation and it is possible that the
partial intercalation of the dpq, mdpq and dppz complexes enables
their closer approach to guanine of DNA.
This journal is © The Royal Society of Chemistry 2008
Fig. 10 Differential pulse voltammograms of 0.25 mM [Ru(Hdpa)2 (dpq)]2+ 5 in the absence (R = 0) and presence (R = 0.25–3.0) of CT
DNA in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1 and
25 ◦ C. Scan rate 5 mV s−1 ; pulse height 50 mV; supporting electrolyte,
50 mM NaCl; working electrode glassy carbon.
Interaction with supercoiled pBR322 DNA
The interaction of complexes 1–5 with supercoiled pBR322 DNA
in 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1 was
studied by agarose gel electrophoresis. The complexes (40 lM)
were incubated with DNA (40 lM in base pairs) for 1 h and
then subjected to gel electrophoresis. The electrophorogram shows
(lanes 2–5, Fig. 11(A)) a pattern for 1–4, which is very similar
to the control indicating that the plasmid DNA is not cleaved.
In contrast, the lane 6 for 5 contains no band corresponding
to SC or any cleaved NC (form II) or LC form (form III);
however, a smearing of DNA is discerned in the gel. When
the concentration of 5 is varied from 0 to 100 lM keeping
the DNA concentration (40 lM) as constant, the SC form is
discernible as a faint band with smearing compared to control
Dalton Trans., 2008, 2157–2170 | 2167
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Table 6 In vitro cytotoxicity assays for complexes 1–5, cisplatin against
cervical epidermoid carcinoma cell line (ME180) (data are mean ±SD
of four replicates each). IC50 = concentration (mM) of drug required to
inhibit growth of 50% of the cancer cells
Fig. 11 (A) Gel electrophoresis diagram showing the interaction of
complexes 1–5 (40 lM) with supercoiled pBR322 DNA (40 lM in base
pair) without added reductant and irradiation in 5% CH3 OH–5 mM
Tris-HCl–50 mM NaCl buffer at pH 7.1 and 37 ◦ C with an incubation time
of 1 h. lane 1, DNA; lane 2, DNA + 1; lane 3, DNA + 2; lane 4, DNA + 3;
lane 5, DNA + 4; lane 6, DNA + 5. (B) Various concentrations of 5 were
reacted with supercoiled pBR322 plasmid DNA (40 lM in base pair) with
an incubation time of 1 h in a 5% CH3 OH–5 mM Tris-HCl–50 mM NaCl
buffer at pH 7.1. Lane 1, DNA control; lane 2, DNA + 5 (20 lM); lane 3,
DNA + 5 (40 lM); lane 4, DNA + 5 (60 lM); lane 5, DNA + 5 (80 lM);
lane 6, DNA + 5 (100 lM). Forms I and II are supercoiled and nicked
circular forms of DNA respectively.
at a low complex concentration (20 lM, Fig. 11(B)). At 40–
60 lM concentrations of the complex no SC form is detected
but, the DNA smear is discerned. At complex concentrations of
80 lM and above DNA smears become faint revealing that EthBr
is completely expelled out of the plasmid due to DNA binding
of the complex at higher concentrations leading to quenching
of EthBr emission (cf. above, competitive binding studies). We
reason out that the complex molecule 5 intercalates with DNA very
strongly and dramatically untwists the plasmid DNA (cf. above)
leading to structural changes and alteration in the superhelicity58
of closed circular DNA. Similar inaccessibility of EthBr to plasmid
DNA upon addition of [Ru(bbdo)(dppz)]2+ has been observed
previously.8 The ability of 1–5 (40 lM) to effect photo-induced
DNA strand scission was also examined by incubating them with
supercoiled pBR322 DNA (40 lM base pairs) in 5% CH3 OH–
5 mM Tris-HCl–50 mM NaCl buffer at pH 7.1 for 1 h. After
irradiation with a monochromatic radiation of 365 nm it is found
that all the complexes fail to cleave the DNA (result not shown).
Cytotoxicity against cervical epidermoid carcinoma cell line
(ME180)
The cytotoxicity of 1–5 against ME180 human cervical epidermoid
carcinoma cell line has been investigated in comparison with
the widely used drug cisplatin and the structurally analogous
and efficient DNA intercalator [Ru(bpy)2 (dppz)]2+ under identical
conditions by using MTT assay. The observed IC50 values (Table 6)
reveal that the complexes 1–4 exhibit cytotoxicity lower than
cisplatin but higher than [Ru(bpy)2 (dppz)]2+ for both 24 and
2168 | Dalton Trans., 2008, 2157–2170
Complex
IC50 (24 h)/lM
IC50 (48 h)/lM
[Ru(Hdpa)2 (phen)]2+ 1
[Ru(Hdpa)2 (5,6-dmp)]2+ 2
[Ru(Hdpa)2 (dpq)]2+ 3
[Ru(Hdpa)2 (mdpq)]2+ 4
[Ru(Hdpa)2 (dppz)]2+ 5
[Ru(bpy)2 (dppz)]2+
cisplatin
30.0 ± 5.0
15.0 ± 2.5
5.5 ± 1.9
5.6 ± 1.2
2.50 ± 0.8
42.0 ± 5.5
45.74 ± 5.00
10.0 ± 2.1
10.0 ± 1.2
4.25 ± 1.53
4.8 ± 1.1
2.50 ± 0.9
37.0 ± 3.9
1.89 ± 0.06
48 h incubation times. The ability of the complexes to exhibit
cytotoxicity follows the order 5 > 3 > 4 > 1 ∼
= 2. Interestingly,
the dppz complex 5 exhibits a potency approximately 8 times
more than cisplatin for 24 h incubation but 4 times lower
activity than cisplatin for 48 h incubation (Table 6). Remarkably,
it exhibits approximately 17 and 15 times more potency than
the structurally analogous [Ru(bpy)2 (dppz)]2+ for 24 and 48 h
incubation respectively. The only structural difference between
[Ru(Hdpa)2 (dppz)]2+ and [Ru(bpy)2 (dppz)]2+ is the amine –NH–
between two pyridine moieties but its incorporation significantly
increases the cytotoxicity of the complex. Thus it is clear that the
–NH– group in the primary ligand Hdpa would make significant
contributions to the cytotoxicity of the complex. Also, the data on
the manual counting of cells with normal and abnormal nuclear
features are shown in Fig. 12(A) and it is evident that the number
of abnormal cells increases in a time-dependent manner and
the complex 5 exhibits a higher percentage of abnormal nuclear
features.
After treating the cells with IC50 concentrations of 5, which
exhibits higher cytotoxicity, for 24 h the cells were observed2b,31
for cytological changes such as fragmented multinucleation, cell
blebbing without micronucleus externalization and cell blebbing
with micronucleus externalization adopting Hoechst 33258 staining (Fig. 12(B)). These interesting cytological changes need further
investigation to find the mode of cell death.
All the above observations clearly show that the dppz complex
5 exhibits DNA binding affinity higher than the other analogous
diimine complexes 1–4. Also, it is the only complex which alters
the DNA superhelicity and causes smearing of the supercoiled
pBR322 DNA in the absence of any external reagent or light. We
have already shown8 that [Ru(N2 S2 )(dppz)]2+ alters the superhelicity of supercoiled pBR322 DNA, forms DNA-intercalator-EthBr
adduct and exhibits a higher cytotoxicity. Further, the complex
5 exhibits a cytotoxicity higher than 1–4. Similarly, apart from
complex 5, the complex 3 exhibits a higher DNA binding affinity
and also a higher cytotoxicity. Finally, the cytotoxicities of the
complexes 1–5 are consistent with their abilities to bind with DNA.
Conclusions
Among the mixed ligand complexes [Ru(Hdpa)2 (diimine)]2+ the
5,6-dmp complex is involved in hydrophobic interaction while the
phen, dpq, mdpq and dppz complexes are involved in intercalation
of the diimine ligands into the DNA base pairs. All the DNA
binding studies such as absorption, emission and circular dichroic
spectral and viscosity experiments suggest that the dppz complex
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Fig. 12 (A) The percentage of cells in each phase is indicated as a graph. Data are mean values obtained from three independent experiments and bars
represent standard deviations. (B) Photomicrograph showing the features of Hoechst 33258 staining of ME180 cervical carcinoma cells: (a–c) untreated
cells (control), (d–l) treated with 5; (d–f) fragmented multinucleation; (g–i) cell blebbing without micronucleus externalization; (j–l) cell blebbing with
micronucleus externalization was observed. Cells were treated with 5 after 24 h seeding.
exhibits the highest DNA binding affinity among the present
complexes. Interestingly, the complexes 3, 4 and 5 exhibit ICD
and exhibit sequence selectivity by binding more strongly to AT
than GC sequence, and the D-enantiomers of the rac-complexes
show the potential to bind preferentially to the right-handed
B DNA. In contrast, the 5,6-dmp complex induces a B to W
conformational change on CT DNA. Also, all the complexes effect
the electrochemical oxidation of guanine moiety of DNA. It is
noteworthy that, of all the complexes, the dppz complex alone alters the DNA superhelicity and causes smearing of the supercoiled
pBR322 DNA in the absence of any external reagent or light. It is
remarkable to find that the same complex exhibits a cytotoxicity
higher than the other analogous diimine complexes and also the
efficient intercalator [Ru(bpy)2 (dppz)]2+ against human cervical
epidermoid carcinoma cell line. So it is concluded from this study
that both Hdpa as a primary ligand with the H-bonding motif
and dppz as an affinity co-ligand have significant potentials as
ligand moieties for designing newer metal-based anticancer drugs.
We are optimistic that further investigation on the mechanism of
action of the dppz compound towards the cell lines would help
to rationally design a promising non-covalent and target specific
anticancer drug.
Acknowledgements
Council of Scientific and Industrial Research, New Delhi, India
(Grant No. 01(2101)/07/EMR-II and SRF to V. R.) is gratefully
acknowledged for financial support. Professor M. Palaniandavar
is a recipient of Ramanna Fellowship, Department of Science and
Technology, New Delhi, India. The Chairman, Molecular BioThis journal is © The Royal Society of Chemistry 2008
physics Unit, Indian Institute of Science, Bangalore is gratefully
acknowledged for Circular Dichroism Spectral facility. University
Grants Commission (UGC), New Delhi and Department of
Science and Technology, New Delhi are gratefully acknowledged
for funding to generate Instruments Facility in the Department
through Special Assistance Program (SAP) of UGC and Funds
for Improvement of S & T Infrastructure (DST-FIST) program of
Department of Science and Technology, New Delhi respectively.
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