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Cytotoxic Ruthenium(II) Complexes Containing a Dangling Pyridine: Selectivity for Diseased Cells Mediated by pH-Dependent DNA Binding.
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Article
Cytotoxic Ruthenium(II) Complexes Containing a Dangling Pyridine:
Selectivity for Diseased Cells Mediated by pH-Dependent DNA
Binding
Somasundaram Sangeetha and Mariappan Murali*
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sı Supporting Information
*
ABSTRACT: Ruthenium(II) complexes of the type [Ru(bpy)2(L1/L2/
L3)]PF6 [where bpy = 2,2′-bipyridine, H(L1) = N-(pyrid-2-yl)salicylaldimine
(1), H(L2) = N-(6-methylpyrid-2-yl)salicylaldimine (2), and H(L3) = N-(4,6dimethylpyrid-2-yl)salicylaldimine (3)] have been isolated. The X-ray
structures of 1−3 reveal distorted octahedral coordination geometry with a
planar ruthenium phenolate moiety. They exhibit interpair dimeric association
in their solid state such as (a) π−π-stacking interactions (1−3) and (b) C−
H···π interactions (2). The 1H NMR spectral data shed light on the
characteristics of metal−ligand bonding and chelate ring conformations. The
complexes exhibit strong metal-to-ligand charge-transfer transitions in the
visible region. The complexes also undergo two successive metal-based
oxidative processes corresponding to the RuII/RuIII and RuIII/RuIV couples.
Resonance Raman studies strongly suggest that the lowest unoccupied
molecular orbital of 1−3 is localized at the bpy ligand. Absorption, emission,
and circular dichroic spectral measurements for 1−3 with calf-thymus DNA reveal a groove binding mode of interaction.
Interestingly, all of the complexes exhibit pH-dependent DNA damage, and the pH at which the damage is highest corresponds to
the pH conditions of the cancer cells. The DNA damage is in the order of 3 > 2 > 1, in which a hydrolytic mechanism dominates.
The protein binding properties of the complexes examined by the tryptophan quenching measurements suggest a static mechanism.
The positive ΔH and ΔS values indicate that the force acting between the complexes and bovine serum albumin (BSA) is mainly a
hydrophobic interaction, and thus BSA may act as a targeted drug-delivery vehicle for ruthenium(II) complexes (K ∼ 105). It is
noteworthy that 3 exhibits selectivity with high cytotoxicity against breast cancer cells (EVSA-T and MCF-7), and its potency is
comparable to that of cisplatin.
■
INTRODUCTION
Cisplatin is one of the most extensively used antitumor drugs
to treat many cancers.1 It causes cytotoxicity by cross-linking
DNA, inducing alterations in the DNA structure that prevent
replication and protein synthesis. However, high general
toxicity, severe side effects, and increasing drug resistance
have restricted the clinical application of cisplatin. The
introduction of special drug-dosing protocols2 has lowered
the general toxicity of cisplatin, although there is still a scope
for further improvements. This gives the thrust to explore
metallodrugs bearing metal ions other than platinum. Under
physiological conditions, the oxidation states RuII, RuIII, and
RuIV are all accessible to ruthenium, making it unique among
the platinum group metals. Ruthenium(III) complexes tend to
be more biologically inert than related ruthenium(II) and
ruthenium(IV) complexes. Ruthenium-based anticancer drugs3
show low general toxicity and demonstrate remarkable
anticancer activity compared to platinum compounds and
accumulate particularly in cancer cells.4 The capacity of
ruthenium to mimic iron ion binding to particular proteins,
© 2022 American Chemical Society
such as serum transferrin and albumin, which are known to be
responsible for the solubilization, transport, and detoxification
of iron in mammals, explains the decreased toxicity.4 Rapidly
proliferating cancer cells have a higher demand for iron,
resulting in an overexpression of transferrin receptors on their
surfaces, causing ruthenium compounds to accumulate in the
cells.4
The discovery of two interesting ruthenium(III) coordination compounds, (ImH)trans-[Ru III Cl 4 (DMSO)(Im)]
(NAMI-A; Im = imidazole and DMSO = dimethyl sulfoxide)5
and (IndH)trans-[RuIIICl4(Ind)2] (KP1019; Ind = indazole),6
developed by the groups of Alessio and Keppler, respectively,
was the first breakthrough in the field of ruthenium-based
Received: October 31, 2021
Published: January 31, 2022
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Inorg. Chem. 2022, 61, 2864−2882
Inorganic Chemistry
pubs.acs.org/IC
Article
Scheme 1. Structure and Proton Numbering Scheme of Ligands
amino acids, and drugs.22,23 Generally, a medicinal drug with
SA is transported and distributed in four steps:24 (i) the drug is
transferred across a biological membrane from its original
binding site or plate; (ii) the drug interacts with SA under
equilibrium conditions to form a drug−SA complex; (iii) the
drug−SA complex is transported through blood plasma; (iv)
finally, it is transferred to its acceptor site or dose parts in a
competitive manner. Bovine serum albumin (BSA) is a SA
protein with several biochemical applications and is frequently
utilized in biological studies because it contains 76% of its
sequence with human serum albumin. This protein is noted for
obtaining larger conformational flexibility to a wide range of
ligands, and it is also inexpensive.
In this study, we explore the DNA binding properties and
BSA interaction capabilities of mixed-ligand ruthenium(II)
complexes of the types [Ru(bpy)2(L1-L3)](PF6) (1−3), where
bpy is 2,2′-bipyridine, and salicylaldimine-type bidentate
ligands (Scheme 1) such as N-(pyrid-2-yl)salicylaldimine
[H(L1)], N-(6-methylpyrid-2-yl)salicylaldimine [H(L2)], and
N-(4,6-dimethylpyrid-2-yl)salicylaldimine [H(L3)]. The salicylaldimine-type bidentate ligands containing coordinative
Nimine and Ophenolate donors constitute similar RuN 5O
chromophores in 1−3 with two bpy ligands and exhibit subtle
variation in the dangling pyridine ring due to the absence or
presence of electron-donating methyl substitution [H(L1), no
methyl; H(L2), 6-methyl; H(L3), 4,6-dimethyl]. It is believed
that the (stereochemical) connection between the ligand
surroundings of chemical nucleases and DNA25,26 is essential
for obtaining high DNA cleavage efficiency and cytotoxicity.
Electronic effects have been demonstrated to be quite crucial
in the cleavage mechanism even though steric effects play a
major role. Therefore, the ability to efficiently synthesize a few
structurally identical ligands with small differences in their
steric and electronic configurations is necessary to optimize
and completely comprehend a chemical nuclease system. As a
group, salicylaldimine-type ligands are made up of a flexible
and kinetically nonlabile ligand template that allows both the
steric and electronic properties of the central metal ion to be
modified in a synthetically simple way.25,26 Although
salicylaldimine-type ligands have a wide range of synthetic
and mechanistic possibilities, electronic tuning has been nearly
completely restricted to aliphatic or aromatic groups bonded to
an imine nitrogen atom. Until recently, there has not been any
practical synthesis of salicylaldimine-type ligands containing a
dangling heterocycle, in which both sterics and electronics can
be manipulated in the same ligand environment. Interestingly,
the present ruthenium(II) complexes (1−3) of salicylaldiminetype ligands containing a dangling pyridine display similar
structural, spectral, and electrochemical properties as well as
possess more or less the same DNA binding and BSA
interaction behavior, but there is distinct DNA cleavage
activity and cytotoxicity because of the presence of both steric
and electronic properties in the same ligand.
anticancer drug prospects. Despite the various studies
conducted to date, the mechanisms of action of NAMI-A
and KP-1019 remain mostly unclear. Alessio described a
variety of unproven myths that have grown up around the
NAMI-A and KP-1019 compounds over the years, significantly
impacting the entire field of ruthenium anticancer drugs.7
Rather, these substances have significantly boosted interest in
nonplatinum anticancer metal compounds and broadened the
logical approach to this field. The principal molecular target of
metal-based anticancer drugs like cisplatin has been identified
as DNA.8−10 So, the interaction of ruthenium complexes with
DNA is being studied to see if DNA binding is effective and if
they may be employed as chemotherapeutic drugs. The
ruthenium(III) complexes are thought to have a different
mode of action than platinum(II) drugs. These ruthenium(III)
complexes are reduced to reactive ruthenium(II) species in
hypoxic tumor tissues and form DNA adducts, adding to the
cytostatic effects. As a result, the development of ruthenium(II) anticancer complexes that have both substantial solubility
in aqueous solutions and cell uptake has received a lot of
interest in the field of biocoordination chemistry to circumvent
the reduction step.11 New classes of cationic and neutral halfsandwich ruthenium(II) arene compounds (also known as
piano-stool compounds) discovered by the Sadler {[(η6biphenyl)Ru(en)Cl](PF6) (RM175; en = ethane-1,2-diamine)} and Dyson {[(η6-p-cymene)RuCl2(pta)] (RAPTA-C;
pta = 1,3,5-triaza-7-phosphatricyclo[3.3.1.1]-decane)}12,13
groups in recent years have been revealed to have promise
anticancer action in vivo and in vitro. All of the rutheniumbased drugs have developed with the intent of covalently
binding the metallic center to its destination (DNA).
Noncovalent interactions between DNA and other molecules
are common in nature and have a significant impact. It can be
demonstrated in the realms of recognition14 and antibiotics15
and useful in the development of new drugs. Groove binding
(major and minor) and intercalation16 are the two main types
of noncovalent DNA bindings that have been recognized for
the last 60 years. Metallodrugs that bind noncovalently to
DNA in these two ways have been developed and have
different actions to cisplatin.16 Intercalation appreciably
influences the DNA characteristics and is described as an
early step in mutagenesis.17 In particular, metal complexes with
planar aromatic ligands that intercalate to DNA are gaining a
great deal of current research.18 These metallointercalators are
highly mutagenic, and some have demonstrated potential
chemotherapeutic efficacy, which is based on their propensity
for DNA binding.19 Moreover, complexes with properly
oriented hydrogen-bonding capabilities can effectively bind
to DNA in either the major or minor grooves.20
Serum albumin (SA) in blood plasma is the most prevalent
soluble protein.21 It is involved in the transportation,
distribution, and metabolism of a wide range of exogenous
and endogenous compounds, including metal ions, fatty acids,
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754, 729, 616, 576, 562, 536, 520 cm−1. ESI-MS: m/z 227 [(M +
H)+]. Anal. Calcd for C14H14N2O: C, 74.31; H, 6.24; N, 12.38.
Found: C, 74.31; H, 6.26; N, 12.57.
Synthesis of Ruthenium(II) Complexes. The complexes
[Ru(bpy) 2 (L 1 )]PF 6 (1), [Ru(bpy) 2 (L 2 )]PF 6 (2), and [Ru(bpy)2(L3)]PF6 (3) were prepared by using the following general
procedure. A 0.260 g sample of [Ru(bpy)2Cl2]·2H2O was heated to
reflux in ethanol (25 mL) with stirring for 1 h. The solid ligand H(L1)
(0.099 g, 0.5 mmol), H(L2) (0.106 g, 0.5 mmol), or H(L3) (0.113 g,
0.5 mmol) and a base, N,N-diisopropylethylamine (0.1 mL), were
then added to the above solution. Stirring was continued, and the
mixture was heated under reflux for 6 h. The color of the solution was
changed from violet to red-brown. An excess of solid NH4PF6 was
added to precipitate the product, and the red-brown crystalline solids
thus formed were filtered off and washed with small amounts of cold
ethanol. The solid complexes were dried in vacuo over P4O10. Finally,
the products were recrystallized from ethanol. Yield: 1, 0.231 g
(61%); 2, 0.246 g (64%); 3, 0.263 g (67%).
Anal. Calcd for C32H25N6OPF6Ru (1): C, 50.87; H, 3.34; N, 11.12.
Found: C, 50.39; H, 3.06; N, 11.02. ΛM in acetonitrile (MeCN) at 25
°C: 141 Ω−1 cm2 mol−1. 1H NMR (DMSO-d6, 300 MHz): δ
[multiplicity, integration, assignment, coupling constant in hertz,
coordination-induced shifts (c.i.s.), δcomplex − δligand] 8.722 (d, 1H,
H3A, 8.1, 0.339), 8.095 (t, 1H, H4A, 7.5, 0.164), 7.693 (t, 1H, H5A,
6.5, 0.257), 9.516 (d, 1H, H6A, 5.1, 0.839), 8.585 (d, 1H, H3B, 8.1,
0.202), 8.185 (t, 1H, H4B, 8.4, 0.254), 7.781 (t, 1H, H5B, 6.6, 0.345),
8.653 (d, 1H, H6B, 6.9, −0.024), 7.498 (d, 1H, H3C, 5.4, −0.885),
7.264 (t, 1H, H4C, 6.6, −0.667), 7.925 (t, 1H, H5C, 5.9, 0.489),
8.653 (d, 1H, H6C, 6.9, −0.024), 7.498 (d, 1H, H3D, 5.4, −0.885),
6.926 (t, 1H, H4D, 6.2, −1.005), 7.563 (t, 1H, H5D, 7.8, 0.127),
8.200 (d, 1H, H6D, 8.1, −0.477), 5.937 (d, 1H, H3E, 7.8, −1.049),
7.344 (t, 1H, H4E, 8.4, −0.112), 6.903 (t, 1H, H5E, 6.5, −0.072),
7.918 (d, 1H, H6E, 7.8, 0.153), 6.445 (d, 1H, H3F, 8.4, −0.981),
7.047 (t, 1H, H4F, 7.7, −0.846), 6.361 (t, 1H, H5F, 7.2, −0.981),
7.276 (d, 1H, H6F, 6.3, −1.248), 8.405 (s, 1H, −CHN−, −1.080)
ppm. Electronic spectrum in MeCN [λmax/nm (εmax/dm3 mol−1
cm−1)]: 245 (34600), 295 (45250), 342 (9980), 371 (9560), 491
(8970), 571 sh. CV (scan rate, 50 mV s−1) and DPV (2 mV s−1) data
in MeCN. Ligand oxidation: Epa, 1.922 V; E1/2 in DPV, 2.089 V.
RuIII/RuII couple: Epa, 0.591 V; Epc, 0.513 V; ΔEp, 68 mV; ipc/ipa, 0.9;
D, 7.4 × 106 cm2 s−1; E1/2 in CV, 0.547 V; E1/2 in DPV, 0.548 V.
RuIV/RuIII couple: Epa, 1.841 V; E1/2 in DPV, 1.706 V. Reduction of
bpy: Epa, −1.651 and −2.074 V; Epc, −1.740 and −1.951 V; ΔEp, 89
and 123 mV; E1/2 in CV, −1.700 and −2.013 V; E1/2 in DPV, −1.719
and −2.021 V.
Anal. Calcd for C33H27N6OPF6Ru (2): C, 51.50; H, 3.54; N, 10.92.
Found: C, 51.21; H, 3.41; N, 10.86. ΛM in MeCN at 25 °C:140 Ω−1
cm2 mol−1. 1H NMR (DMSO-d6, 400 MHz): δ (multiplicity,
integration, assignment, coupling constant in hertz, c.i.s., δcomplex −
δligand) 8.717 (d, 1H, H3A, 8.0, 0.334), 8.095 (t, 1H, H4A, 8.0, 0.164),
7.687 (t, 1H, H5A, 5.2, 0.251), 9.514 (d, 1H, H6A, 4.8, 0.837), 8.602
(d, 1H, H3B, 8.0, 0.219), 8.178 (t, 1H, H4B, 7.8, 0.247), 7.771 (t, 1H,
H5B, 6.6, 0.335), 8.648 (d, 1H, H6B, 6.4, −0.029), 7.506 (d, 1H,
H3C, 5.2, −0.877), 7.256 (t, 1H, H4C, 5.2, −0.675), 7.929 (t, 1H,
H5C, 8.0, 0.493), 8.648 (d, 1H, H6C, 6.4, −0.029), 7.465 (d, 1H,
H3D, 5.2, −0.918), 7.036 (t, 1H, H4D, 7.6, −0.895), 7.552 (t, 1H,
H5D, 7.8, 0.116), 8.222 (d, 1H, H6D, 8.0, −0.455), 6.401 (d, 1H,
H3E, 7.2, −0.570), 6.955 (t, 1H, H4E, 7.6, −0.465), 6.343 (t, 1H,
H5E, 6.4, −0.596), 7.265 (d, 1H, H6E, 6.0, −0.520), 5.930 (d, 1H,
H3F, 8.0, −1.266), 7.274 (t, 1H, H4F, 6.6, −0.478), 6.731 (d, 1H,
H5F, 7.6, −0.487), 2.079 (s, 3H, 6-CH3, −0.393), 8.364 (s, 1H,
−CHN−, −1.058) ppm. Electronic spectrum in MeCN [λmax/nm
(εmax/dm3 mol−1 cm−1)]: 245 (34410), 295 (44650), 342 (10220),
371 (9190), 491 (8970), 571 sh. CV (scan rate, 50 mV s−1) and DPV
(2 mV s−1) data in MeCN. Ligand oxidation: Epa, 1.943 V; E1/2 in
DPV, 1.889 V. RuIII/RuII couple: Epa, 0.612 V; Epc, 0.544 V; ΔEp, 68
mV; ipc/ipa, 0.8; D, 7.4 × 106 cm2 s−1; E1/2 in CV, 0.578 V; E1/2 in
DPV, 0.568 V. RuIV/RuIII couple: Epa, 1.833 V; E1/2 in DPV, 1.698 V.
Reduction of bpy: Epa, −1.741 and −2.072 V; Epc, −1.649 and −1.953
EXPERIMENTAL SECTION
Materials and Methods. Ruthenium trichloride trihydrate (Arora
Mathey), agarose (Genei), and ethidium bromide (EthBr; Merck)
were utilized as supplied. 2,2′-Bipyridine, salicylaldehyde, 2-aminopyridine, 2-amino-6-picoline, 4,6-dimethyl-2-aminopyridine, and calfthymus (CT) DNA (stored at −20 °C) were purchased from SigmaAldrich. Invitrogen Life Technologies provided the ΦX174 supercoiled phage DNA (0.25 μg μL−1), which was kept at −20 °C. Bovine
serum albumin (BSA) was acquired from Sigma-Aldrich and kept at 4
°C. Ultrapure Milli-Q water (18.2 mΩ) was utilized for all studies.
The solvents were acquired from Biosolve (AR grade) and employed
immediately in the syntheses. The precursor cis-[Ru(bpy)2Cl2]·2H2O
was synthesized following the procedure in the literature.27
MCF7 (breast cancer), EVSA-T (breast cancer), WIDR (colon
cancer), IGROV (ovarian cancer), M19 (melanoma), A498 (renal
cancer), and H226 (nonsmall cell lung cancer) were among the seven
human tumor cell lines employed. The WIDR, M19, A498, IGROV,
and H226 cell lines are from an anticancer screening panel of the
National Cancer Institute, USA.28 MCF7 and EVSA-T are estrogen
receptor (ER)+/progesterone receptor (PgR)+ and (ER)−/(PgR)−
human breast cancer cell lines, respectively. A mycoplasma test was
performed on all cell lines prior to the tests, and this turned out to be
negative. All cell lines were grown in a continuous logarithmic culture
in RPMI 1640 (Gibco, Invitrogen, Paisley, Scotland) media with 4-(2hydroxyethyl)-1-piperazineethanesulfonic acid and phenol red. The
medium was supplemented with 10% fetal calf serum (Gibco,
Invitrogen, Paisley, Scotland), 100 units mL−1 penicillin (Sigma, St.
Louis, MO), and 100 g mL−1 streptomycin (Sigma, St. Louis, MO).
For passage and use in the study, the cells were slightly trypsinized.
Physical Measurements. Kindly consult the Supporting
Information for the details of physical measurements.
Synthesis of Ligands. A solution of salicylaldehyde (3.1 g, 0.025
mol) in methanol (10 mL) was added, under constant stirring, to a
solution of 2-aminopyridine (2.4 g, 0.025 mol), 2-amino-6-picoline
(2.6 g, 0.025 mol), or 4,6-dimethyl-2-aminopyridine (3.1 g, 0.025
mol) in methanol (30 mL). The reaction mixture was refluxed (70
°C) for 2 h. Methanol was evaporated to dryness under reduced
pressure to yield an orange oily residue. The oily material was
redissolved in warm CH2Cl2 (20 mL) and left at room temperature
for 12 h. The orange-yellow crystals were collected, washed with
diethyl ether, and dried in a vacuum.
N-(Pyrid-2-yl)salicylaldimine [H(L1)]. Yield: 4.0 g, 80%. 1H NMR
(DMSO-d6, 300 MHz): group I, δ 6.986 (d, 1H, H3E), 7.456 (t, 1H,
H4E), 6.975 (t, 1H, H5E), 7.765 (d, 1H, H6E), 13.041 (s, 1H,
−OH); group II, δ 7.426 (d, 1H, H3F), 7.893 (t, 1H, H4F), 7.342 (t,
1H, H5F), 8.524 (d, 1H, H6F), 9.485 (s, 1H, −CHN−) ppm. IR
(neat): 3424, 3054, 2563, 1651, 1607, 1588, 1575, 1558, 1497, 1466,
1455, 1430, 1352, 1278, 1232, 1185, 1146, 1111, 1044, 1030, 1004,
994, 960, 915, 879, 845, 789, 780, 744, 733, 676, 624, 578, 563, 528,
503 cm−1. ESI-MS: m/z 199 [(M + H)+]. Anal. Calcd for
C12H10N2O: C, 72.71; H, 5.08; N, 14.13. Found: C, 72.36; H,
5.14; N, 14.03.
N-(6-Methylpyrid-2-yl)salicylaldimine [H(L2)]. Yield: 4.5 g, 85%.
1
H NMR (DMSO-d6, 300 MHz): group I, δ 6.971 (d, 1H, H3E),
7.420 (t, 1H, H4E), 6.939 (t, 1H, H5E), 7.785 (d, 1H, H6E), 13.109
(s, 1H, −OH); group II, δ 7.196 (d, 1H, H3F), 7.745 (t, 1H, H4F),
7.218 (d, 1H, H5F), 2.472 (s, 3H, 4-CH3), 9.423 (s, 1H, −CHN−)
ppm. IR (neat): 3418, 3052, 2918, 1651, 1610, 1552, 1575, 1498,
1450, 1282, 1232, 1185, 1144, 992, 896, 821, 794, 757, 723, 579, 564,
548 cm−1. ESI-MS: m/z 213 [(M + H)+]. Anal. Calcd for
C13H12N2O: C, 73.57; H, 5.70; N, 13.20. Found: C, 73.57; H,
5.75; N, 13.39.
N-(4,6-Dimethylpyrid-2-yl)salicylaldimine [H(L3)]. Yield: 5.0 g,
88%. 1H NMR (DMSO-d6, 300 MHz): group I, δ 6.983 (d, 1H,
H3E), 7.432 (t, 1H, H4E), 6.952 (t, 1H, H5E), 7.747 (d, 1H, H6E),
13.215 (s, 1H, −OH); group II, δ 7.074 (s, 1H, H3F), 7.047 (s, 1H,
H5F), 2.315 (s, 3H, 4-CH3), 2.455 (s, 3H, 6-CH3), 9.432 (s, 1H,
−CHN−) ppm. IR (neat): 3477, 3064, 2918, 1648, 1603, 1542,
1498, 1452, 1370, 1279, 1196, 1145, 1118, 1040, 994, 899, 846, 794,
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Table 1. Selected Crystal Data and Structure Refinement Parameters for 1−3
formula
fw (g mol−1)
temperature (K)
wavelength (Å)
cryst syst
space group
a (Å)
b (Å)
c (Å)
α (deg)
β (deg)
γ (deg)
V (Å)3, Z
Dcalc (g cm−3)
μ (mm−1)
F(000)
cryst size (mm3)
θ (deg)
index ranges
reflns collected
indep reflns
reflns obsd [I > 2σ(I)]
Rint
GOF
R1, wR2[I > 2σ(I)]
R1, wR2 (all data)
[Ru(bpy)2(L1)](PF6) (1)
[Ru(bpy)2(L2)](PF6) (2)
[Ru(bpy)2(L3)](PF6) (3)
C32H25F6N6OPRu
755.62
293(2)
0.71069
orthorhombic
Pbca
16.5791(8)
18.4378(9)
20.3836(10)
90
90(10)
90
6230.9(5), 8
1.611
0.628
3040
0.27 × 0.24 × 0.20
1.93−29.60
−22 ≤ h ≤ 22, −24 ≤ k ≤ 25, −27 ≤ l ≤ 26
75188
8264
4928
0.0458
1.008
0.0487, 0.0890
0.0887, 0.1612
C33H27F6N6OPRu
769.65
293(2)
0.71069
monoclinic
P21/n
10.4489(7)
20.8610(13)
15.1307(9)
90
102(10)
90
3218.7(4), 4
1.588
0.610
1552
0.23 × 0.18 × 0.16
1.69−28.22
−13 ≤ h ≤ 13, −27 ≤ k ≤ 27, −19 ≤ l ≤ 19
34523
7398
5031
0.0311
1.018
0.0835, 0.2550
0.1089, 0.2765
C34H29F6N6OPRu
783.67
293(2)
0.71069
monoclinic
P21/n
10.9642(9)
20.9054(16)
15.1969(12)
90
105.891(2)
90
3350.2(5), 4
1.554
0.587
1584
0.20 × 0.16 × 0.15
1.70−29.00
−14 ≤ h ≤ 14, −27 ≤ k ≤ 28, −20 ≤ l ≤ 19
43292
8707
4573
0.0468
1.003
0.0510, 0.1354
0.1039, 0.1518
V; ΔEp, 92 and 119 mV; E1/2 in CV, −1.695 and −2.013 V; E1/2 in
DPV, −1.719 and −2.032 V.
Anal. Calcd for C34H29N6OPF6Ru (3): C, 52.11; H, 3.73; N, 10.72.
Found: C, 51.85; H, 3.54; N, 10.70. ΛM in MeCN at 25 °C: 141 Ω−1
cm2 mol−1. 1H NMR (DMSO-d6, 400 MHz): δ (multiplicity,
integration, assignment, coupling constant in hertz, c.i.s., δcomplex −
δligand) 8.714 (d, 1H, H3A, 8.0, 0.331), 8.091 (t, 1H, H4A, 8.0, 0.160),
7.689 (t, 1H, H5A, 6.6, 0.253), 9.516 (d, 1H, H6A, 5.6, 0.839), 8.632
(d, 1H, H3B, 8.0, 0.249), 8.191 (t, 1H, H4B, 7.8, 0.260), 7.776 (t, 1H,
H5B, 6.6, 0.340), 8.653 (d, 1H, H6B, 7.6, −0.024), 7.534 (d, 1H,
H3C, 5.2, −0.849), 7.266 (t, 1H, H4C, 6.6, −0.665), 7.933 (t, 1H,
H5C, 7.8, 0.497), 8.653 (d, 1H, H6C, 7.6, −0.024), 7.444 (d, 1H,
H3D, 5.2, −0.939), 6.954 (t, 1H, H4D, 6.6, −0.977), 7.559 (t, 1H,
H5D, 7.6, 0.123), 8.246 (d, 1H, H6D, 8.0, −0.431), 6.423 (d, 1H,
H3E, 8.4, −0.560), 7.025 (t, 1H, H4E, 7.8, −0.407), 6.347 (t, 1H,
H5E, 7.6, −0.605), 7.273 (d, 1H, H6E, 7.2, −0.474), 5.532 (s, 1H,
H3F, −1.542), 6.550 (s, 1H, H5F, −0.497), 1.914 (s, 3H, 4-CH3,
−0.401), 2.038 (s, 3H, 6-CH3, −0.417), 8.363 (s, 1H, −CHN−,
−1.069) ppm. Electronic spectrum in MeCN [λmax/nm (εmax/dm3
mol−1 cm−1)]: 245 (37550), 295 (46840), 342 (10590), 371 (10320),
491 (9310), 571 sh. CV (scan rate, 50 mV s−1) and DPV (2 mV s−1)
data in MeCN. Ligand oxidation: Epa, 2.080 V; E1/2 in DPV, 1.940 V.
RuIII/RuII couple: Epa, 0.569 V; Epc, 0.502 V; ΔEp, 67 mV; ipc/ipa, 0.9;
D, 7.2 × 106 cm2 s−1; E1/2 in CV, 0.536 V; E1/2 in DPV, 0.545 V.
RuIV/RuIII couple: Epa, 1.803 V; E1/2 in DPV, 1.668 V. Reduction of
bpy: Epa, −1.734 and −2.071 V; Epc, −1.642 and −1.952 V; ΔEp, 92
and 119 mV; E1/2 in CV, −1.688 and −2.012 V; E1/2 in DPV, −1.719
and −2.032 V.
X-ray Crystallographic Procedures. Upon slow evaporation of
an ethanolic solution of the respective complex, single crystals of 1−3
were formed. A crystal of 1, 2, or 3 was mounted on a glass fiber using
epoxy cement. A Bruker SMART APEX CCD diffractometer with a
fine-focus sealed-tube Mo Kα X-ray source was used to collect X-ray
diffraction data. The intensity data for all of the crystals were recorded
using the ω-scan technique. Data acquisition was done using SMART
software, and data were extracted using SAINT. Empirical absorption
corrections were made on the intensity data.29 The space group for 1
was assigned as Pbca, while 2 and 3 were assigned as P21/n using
systematic absences and E statistics of the data set. These space
groups were later confirmed by successful structure solution and
refinement. The details of data collection and structure analysis are
given in Table 1. A direct method was used to solve the structures of
1, 2, or 3 using the SIR9730 program and refined with the SHELXL 97
program31 using the full-matrix least-squares method on F2.
Anisotropic refinement was applied to all non-hydrogen atoms. We
found no disorder in PF6− for 2. Six fluorine atoms in the PF6− anion
for 1 and 3 were disordered. Site occupancies of the disordered
fluorine atoms were refined, and their sum was limited to one. The
hydrogen atoms in the complex were fixed in place and refined with
the help of a riding model. The goodness-of-fit (GOF) values of
1.008(1), 1.018(2), and 1.003(3) were achieved during final
refinement of the structures of 1−3. Selected bond lengths and
angles are furnished in Table 2.
DNA and Protein Binding Experiments. The ratio of UV
absorbances at 260 and 280 nm, A260/A280, of DNA solutions in a 5
mM Tris HCl/50 mM NaCl buffer was 1.9,32 suggesting that the
DNA was adequately free of protein.
The stock solution of protein (1.0 × 10−4 mol L−1) was prepared
by dissolving solid BSA in a 0.05 M phosphate buffer at pH 7.4, stored
at 0−4 °C in the dark for about 1 week, and then diluted to 1.0 ×
10−6 mol L−1 using a phosphate buffer (pH 7.4, 0.05 M) upon use.
The concentration of BSA was determined from optical density
measurements, using a value of the molar absorptivity ε280 of 44720
M−1 cm−1.33 The detailed procedure for the spectroscopic experiments of DNA and protein binding as well as the electrochemical
experiments for DNA binding is described in the Supporting
Information.
DNA Cleavage Experiments. In the pH-dependent DNA
cleavage test, a 2.5 mM ruthenium(II) complex in 0.1 M phosphate
buffer, pH 6.0−8.0 in 0.5 step increments, was incubated for 4 h with
0.05 μg μL−1 ΦX174 RF supercoiled phage DNA. The DNA damage
was evaluated by comparing the electrophoretic movement of the
species in a 1% agarose gel, prepared in a 1 × TBE buffer, to those of
the control incubations: control 2 examined the activity of the
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RESULTS AND DISCUSSION
Syntheses of Ligands and Ruthenium(II) Complexes.
Condensation reactions of 2-aminopyridine, 2-amino-6-picoline, or 4,6-dimethyl-2-aminopyridine with salicylaldehyde in
refluxing methanol for 2 h lead to new heterofunctionalized
phenol−imine bidentate ligands containing imine nitrogen
(Nimine) and phenolate oxygen (Ophenolate) donors and dangling
pyridine [H(L1)], 6-methylpyridine [H(L2)], or 4,6-dimethylpyridine [H(L3)] in high yields. The three ligands primarily
differ concerning the nature of the substituent(s) present in
the pyridine ring connected to the Nimine atom. The anionic
forms of the ligands (L1−3) are expected to bind to the metal
ion in a bidentate N,O manner, forming a six-membered
chelate ring. The ligands are soluble in a range of organic
solvents and are stable to hydrolysis and aerial oxidation. The
strong broad band observed at 3418−3477 cm−1 in the spectra
is assigned to the ν(OH) vibration. The Schiff bases display
their CN band within the wavenumber range 1648−1651
cm−1. The phenolic C−O of H(L1)−H(L3) is observed at
1278−1282 cm−1. A weak band observed at 992−994 cm−1 is
ascribed to the aldehydic ν(CH) (−CHN−) vibration.
The reaction of H(L1), H(L2), or H(L3) in the presence of a
base (N,N-diisopropylethylamine) with 1 equiv of [Ru(bpy)2Cl2]·2H2O in ethanol at reflux gave a red-brown
solution, from which a pure crystalline red-brown solid
precipitated upon the addition of NH4PF6; the addition of a
base was necessary to deprotonate the hydroxyl group and
render it a better coordination site. The microanalytical data
were as expected in accordance with the formation of
[Ru(bpy)2(L1/L2/L3)]PF6, in which L1, L2, or L3 acts as a
monoanionic phenolate−imine bidentate donor, and they were
confirmed by X-ray structure analysis. Conductivity experiments indicate that they behave as 1:1 electrolytes in a MeCN
solution. The maximum molecular peaks are observed from
electrospray ionization mass spectrometry (ESI-MS) spectra at
m/z 610.7 (1), 624.7 (2), and 638.7 (3), which match well
with the corresponding calculated masses (1, m/z 611.1; 2, m/
z 625.1; 3, m/z 639.1) of the monocations. As expected, the
free Schiff-base phenolic OH stretching35 (∼3400 cm−1) is not
observed in the IR spectrum of the complex. The CN
stretching36 is observed as a strong peak around 1600 cm−1
with a bathochromic shift between 37 and 49 cm−1, which is
typical for coordinated imines of this type.37 The diamagnetic
complexes 1−3 correspond to the bivalent state of ruthenium
(low-spin d6, S = 0).
Description of the Crystal Structures. The ORTEP
views of [Ru(bpy)2(L1)]+ (1), [Ru(bpy)2(L2)]+ (2), and
[Ru(bpy)2(L3)]+ (3) are illustrated in Figure 1A−C. The
crystal structures of the complexes contain a racemic mixture
of Λ/Δ enantiomeric pairs. In the structures of 1−3, the
ruthenium(II) ion is coordinated by the two bidentate bpy
ligands and a monoanionic phenolate−imine bidentate Schiffbase ligand, which forms a RuN5O chromophore in a distorted
octahedral geometry, as can be seen from the angles subtended
at the metal. The average cis and trans angles are 88.65(12)°
(1), 88.15(5)° (2), or 88.94(13)° (3) and 174.41(11)° (1),
173.75(8)° (2), or 173.44(13)° (3), respectively. Distortion of
the coordination sphere is primarily caused by the two acute
(∼79°) bite angles of the two juxtaposed bpy chelate rings.
The halves of the bpy ligands are nearly coplanar with dihedral
angles of 5.5 and 4.2° (1), 5.4 and 1.7° (2), and 1.4 and 5.3°
(3) about the 2,2′ bond. The bite angles of the phenolate−
Table 2. Selected Bond Lengths (Å) and Bond Angles (deg)
for 1−3
N1−Ru1
N4−Ru1
N6−Ru1
O1−Ru1−N1
O1−Ru1−N4
O1−Ru1−N6
N1−Ru1−N4
N1−Ru1−N6
N3−Ru1−N5
N4−Ru1−N5
N5−Ru1−N6
N1−Ru1
N4−Ru1
N6−Ru1
O1−Ru1−N1
O1−Ru1−N4
O1−Ru1−N6
N1−Ru1−N4
N1−Ru1−N6
N3−Ru1−N5
N4−Ru1−N5
N5−Ru1−N6
N1−Ru1
N4−Ru1
N6−Ru1
O1−Ru1−N1
O1−Ru1−N4
O1−Ru1−N6
N1−Ru1−N4
N1−Ru1−N6
N3−Ru1−N5
N4−Ru1−N5
N5−Ru1−N6
[Ru(bpy)2(L1)](PF6) (1)
2.067(3)
N3−Ru1
2.059(3)
N5−Ru1
2.043(3)
O1−Ru1
89.53(11)
O1−Ru1−N3
88.72(11)
O1−Ru1−N5
94.49(11)
N1−Ru1−N3
175.45(12)
N1−Ru1−N5
86.62(11)
N3−Ru1−N4
98.38(12)
N3−Ru1−N6
87.66(12)
N4−Ru1−N6
78.92(12)
[Ru(bpy)2(L2)](PF6) (2)
2.066(5)
N3−Ru1
2.032(5)
N5−Ru1
2.041(5)
O1−Ru1
88.98(19)
O1−Ru1−N3
89.30(19)
O1−Ru1−N5
94.6(2)
N1−Ru1−N3
174.2(2)
N1−Ru1−N5
89.1(2)
N3−Ru1−N4
97.5(2)
N3−Ru1−N6
88.5(2)
N4−Ru1−N6
78.8(2)
[Ru(bpy)2(L3)](PF6) (3)
2.065(3)
N3−Ru1
2.049(3)
N5−Ru1
2.038(3)
O1−Ru1
89.38(12)
O1−Ru1−N3
87.14(11)
O1−Ru1−N5
93.45(13)
N1−Ru1−N3
173.29(13)
N1−Ru1−N5
86.86(13)
N3−Ru1−N4
97.05(12)
N3−Ru1−N6
89.34(12)
N4−Ru1−N6
79.21(14)
Article
2.066(3)
2.035(3)
2.074(2)
87.94(11)
171.98(11)
96.84(11)
94.59(12)
78.90(12)
175.80(11)
97.71(12)
2.059(5)
2.018(5)
2.083(4)
88.9(2)
172.7(2)
95.4(2)
93.9(2)
79.0(2)
174.36(19)
96.6(2)
2.057(3)
2.028(3)
2.074(2)
90.11(12)
171.26(13)
95.48(13)
94.89(13)
78.80(14)
175.76(13)
99.07(14)
complex under assay settings that are known to cause DNA
retardation (as above but with no phosphate buffer), and control 1
was DNA in conditions identical with those of control 2 but without
the complex. From each test, 20 μL was added with 4 μL of dye
(0.025 mg of bromophenol blue, 1 mL of glycerol, and 1 mL of MilliQ water) and pipetted into wells on the horizontal gel. A potential
difference of 40 mV was applied for 3 h to the gel, and the bands were
observed using EthBr staining. A similar experiment was carried out in
the absence of a complex to evaluate the effect of pH alone on DNA.
T4 Ligase-Religation Studies. An enzymatic study was carried
out using T4 DNA ligase to find out whether the cleaved products
were consistent with hydrolysis of the phosphodiester linkages in
DNA. For the religation tests, the solution was incubated for 18 h at
16 °C prior to gel electrophoresis. The nicked circular (NC) DNA
obtained from the hydrolytic cleavage reaction was recovered from
agarose gel by a phenol extraction method and purified by ethanol
precipitation. This was followed by the addition of a 10× ligation
buffer and T4 DNA ligase (4 units) to the purified NC DNA.
Sulforhodamine B (SRB) Test. The complexes were dissolved to
a concentration of 5 mg mL−1 in DMSO (Sigma, St. Louis, MO) and
subsequently diluted to a final concentration of 250000 ng mL−1 in a
full medium. The cytotoxicity was estimated by a microculture SRB
test.34 The Supporting Information contains a detailed procedure for
the SRB test.
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Figure 1. ORTEP views of 1 (A), 2 (B), and 3 (C) with atom numbering of the complex cation and thermal ellipsoids at 40% probability. Packing
diagrams D−F showing the π−π interaction (dotted lines) in [Ru(bpy)2(L1)]+ (1), the π−π and C−H···π interactions (dotted lines) in
[Ru(bpy)2(L2)]+ (2), and the π−π interaction (dotted lines) in [Ru(bpy)2(L3)]+ (3), respectively.
1−3 are probably due to the steric effect arising from the sixmembered chelate ring or may be a steric hindrance of the
dangling pyridine of L1−L3. The Ru−N2 distance is 3.86 Å
(1), 3.63 Å (2), or 3.81 Å (3), revealing that there is no
interaction with the seventh coordinating atom.
The Ru−Nimine bond length is shorter than the Ru−Ophenolate
despite the fact that Ophenolate is a better σ-donor. The bond
distance of N1−C7 is 1.305(5) (1) or 1.278(9) (2) or
1.301(5) Å (3), which is normal for a C = N double bond.40
However, closer examination of salicylaldimine of L1 (1) bond
distances show that the C2−C3 [1.372(6) Å] and C4−C5
[1.373(6) Å] are shorter than C1−C2 [1.416(5) Å], C3−C4
[1.377(7) Å], C5−C6 [1.414(6) Å] and C6−C1 [1.427(5) Å].
Also, C6−C7 [1.426(5) Å] is shorter than that observed for
the free ligand (1.448 Å).38 Similarly, the imine bond C7−N1
[1.305(5) Å] is longer than in the free ligand (1.285 Å) and
the phenolate bond C1−O1 [1.299(6) Å] is shorter than in
the free ligand (1.348 Å). These trends in Ru−Ophenolate and
Ru−Nimine distances and distances within the salicylaldimine
imine chelate, 89.53(11)° (1), 88.98(19)° (2), and 89.38(12)°
(3), are very close to the ideal value.
The Ru−Nbpy bond lengths within each complex are slightly
different and lie in the ranges 2.035(3)−2.066(3) Å (1),
2.018(5)−2.059(5) Å (2), and 2.028(3)−2.057(3) Å (3),
which bracket the Ru−N distance in [Ru(bpy)3]2+ [2.056(6)
Å].38 The shortest Ru−Nbpy distance is to the nitrogen atom
trans to the σ-donating phenolate, which is consistent with
improved Ru(dπ) → bpy(π*) back-bonding due to the
increased electron density at the metal center. The Ru−N1
[1, 2.067(3) Å; 2, 2.066(5) Å; 3, 2.065(3) Å] bond length
(Nimine atom of L1−L3) is almost similar to that of Ru−N3 of
the cis-coordinated bpy moiety, indicating that the Nimine atom
is more strongly bound similar to the nitrogen atom of bpy but
longer than the Ru−N imine distance found in [Ru(bpy)2(HNCHC6H4O)]+, 2.039(4) Å.39 The ruthenium
phenolate moiety is essentially planar. Ru−Ophenolate is
observed to be 2.074(2) Å (1), 2.083(4) Å (2), or 2.074(2)
Å (3), which is also longer than 2.060(3) Å observed in
[Ru(bpy)2(HNCHC6H4O)]+.39 The longer bond distances in
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Scheme 2. Ligand Orientation and Proton Numbering Pattern for 1−3
= 8.7 and 11.4 ppm, respectively.39 A downfield shift is
observed for both protons of all of the ligands relative to the
tetradentate ligand, which is ascribed to the effect of the ring
current associated with the conjugated system. It seems that
this invariability in the present investigation is caused by a
rather strong hydrogen-bonding interaction between the
hydrogen atom of the phenolic −OH and the nitrogen atom
of the imine group. Such hydrogen bonding is not present in
the aforementioned salicylaldimine-type tetradentate ligand.
The X-ray crystal structure41,43 of the free ligand H(L1) reveals
that the molecule is planar and that the dangling pyridine ring
has the same orientation for the rest of the molecule
(salicylaldimine moiety). For instance, the pyridine nitrogen
atom is always oriented cis to the hydrogen atom of the imine
group, which indicates a strong intramolecular hydrogenbonding interaction. The orders of the protons in increasing
field strength are H(L1) > H(L2) ≈ H(L3) (−CHN−) and
H(L3) > H(L2) > H(L1) (−OH). These observations reveal
(i) an increase in the van der Waals interaction between the
azomethine proton and the nitrogen lone pair of 2-pyridine,
H(L1), and (ii) a stronger intramolecular hydrogen bonding
between the hydrogen atom of the phenolic −OH and the
nitrogen atom of the imine group in H(L3) relative to H(L1)
and H(L2).
The 1H NMR spectra of the complexes display 24 (1), 23
(2), or 21 (3) nonequivalent aromatic signals (Figure 2) due
to the presence of an unsymmetric salicylaldimine moiety in
1−3, which makes all six aromatic rings nonequivalent. The
absence of a phenolic −OH proton of the free ligands H(L1)−
H(L3) in the spectra of the complexes suggests coordination
through Ophenolate of H(L1)−H(L3). Therefore, the equatorial
plane contains one pyridine ring from each of the two bpy
ligands (rings B and D: H3−H6), the oxygen atom of the
phenolate group (ring E: H3−H6), and the azomethine
nitrogen atom of H(L1)−H(L3). The axial positions are
occupied by the other pyridine rings of the bpy ligands (rings
A and C: H3−H6), while the uncoordinated pyridine ring
(ring F: 1, H3−H6; 2, H3−H5 and 6-CH3; 3, H3, H5, and 4and 6-CH3) of H(L1)−H(L3) is dangling. When coordinated
to ruthenium(II), the planar bpy rings are expected to form a
five-membered chelate ring with an envelope conformation
and the unsymmetric salicylaldimine moiety of H(L1)−H(L3)
forms a six-membered chelate ring with a twist boat
conformation, as in the X-ray crystal structures of 1−3.
unit support a description of significant delocalization of the
negative charge between the Nimine and Ophenolate atoms.
The X-ray crystal structure41 of the free ligand H(L1) reveals
that both molecules are planar and that the dangling pyridine
ring has the same orientation with respect to the rest of the
molecule (salicylaldimine moiety). However, in the complexes,
the increased interplanar angles observed at 78.3° (1), 73.6°
(2), and 84.3° (3) indicate that the dangling pyridine ring has
a perpendicular orientation with respect to the salicylaldimine
unit. It is worth mentioning that the lattice structures42 of 1−3
are further stabilized by π−π-stacking interactions because of
intermolecular interactions between 2,2′-bipyridine ligands
(Figure 1D−F) with Cg···Cg′ distances of 3.840 Å (1), 3.946
Å (2), and 3.853 Å (3) (Cg and Cg′ are the N3/C13−C17
and N4/C18−C22 ring centroids) and interplanar angles of
6.53° (1), 5.19° (2), and 6.09° (3). These π−π aromatic
interactions in 2 are further reinforced by C−H···π interactions
(Figure 1E) via atoms C2−H2 of the phenolate ring and C25
of one of the bpy rings with a distance of 3.328 Å.
1
H NMR Spectra. The chemical shifts of the free and
coordinated ligands (Scheme 2) are summarized in the
Experimental Section, along with coordination-induced shifts
(c.i.s. = δcomplex − δligand) and coupling constants. The J values
were compatible with the spectral assignments, which were
based on the COSY spectra of the complexes.
The 1H NMR spectra of free ligands (HL1−HL3) could be
mainly classified into two groups (I and II). Group I has five
resonances, and the chemical shifts are typical for salicylaldimine derivatives.43 The peaks correspond to the protons of the
phenol moieties [H(L1)−H(L3), H3−H6, and −OH]. This
suggests that the Schiff bases exist in the enolimine form in a
DMSO solution. The two H(L3), three H(L2), or four H(L1)
aromatic resonances and the one H(L2) or two H(L3) aliphatic
resonances found for group II are assigned to the protons of
the corresponding pyridine rings connected to the Nimine atom.
The most significant signal in the spectra is a singlet (δ = 9.4−
9.5 ppm), which is assigned to the azomethine (−CHN−)
proton. As expected, the resonances for the protons H3−H6 of
the phenol moiety are unaffected by the pyridine group
attached to the Nimine atom. Interestingly, the nature of the
substituent present in the pyridine ring connected to the Nimine
atom severely affects the chemical shift of the protons of the
−CHN− and −OH functions. Notably, the −CHN− and
−OH protons in the analogous tetradentate ligand appear at δ
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Figure 2. 1H NMR spectra of (a) 1, (b) 2, and (c) 3 in (CD3)2SO.
Thus, the observable singlet of the azomethine (−CHN−)
proton of the salicylaldimine moiety is shielded at 8.405 ppm
(1), 8.364 ppm (2), or 8.363 ppm (3) compared to free
ligands (9.422−9.485 ppm), resulting in negative c.i.s. values.
The azomethine proton is likely close to ruthenium(II) and
hence experiences electron density around it because of the
twist boat conformation imposed by the six-membered chelate
ring. The most upfield signals appearing for ring D protons
(H3−H6), which are trans to Ophenolate, are significantly
different from the other resonances because the ring
experiences a trans effect of the σ-donor phenolate function.
The order of the pyridine ring protons in increasing field
strength appears to be 6 > 3 > 4 > 5 for rings A and B, but for
rings C and D, it is 6 > 5 > 3 > 4. The protons of the phenolate
function follow the order 6 > 4 > 5 > 3 for 1, but for 2 and 3, it
is 6 > 5 > 3 > 4. The 6-CH3 (2.079 ppm) and the 4-CH3
(1.914 ppm) and 6-CH3 (2.038 ppm) upfield signals of 2 and
3, respectively, appear as singlets. Because coordination forces
the pyridine rings of bpy to be coplanar, the dangling pyridine
ring in H(L2)/H(L3) is obliged to rotate away from bpy to
minimize the steric interactions with bpy, and hence the 6-CH3
(2) and 4- and 6-CH3 (3) protons of a dangling pyridine ring
are exposed to the shielding magnetic anisotropy due to the
ring currents of the pyridine ring of bpy.
The large positive c.i.s. value for H6A (1-3) proton arise
from an σ-effect based on electron donation to Ru(II) via the
nitrogen lone pairs. A negative c.i.s. value observed for the
H6D (1-3) proton, adjacent to the coordinating nitrogen,
would result from interligand through-space ring-current
anisotropy effects and there is a lesser influence on H6B and
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tions ΔEp (67−68 mV) are larger than the Nernstian value of
59 mV for a one-electron transfer; they are typical for
complexes of this kind, owing to uncompensated solution
resistance.49 The substitution of one π-acidic bpy ligand from
the [Ru(bpy)3]2+ core by one σ-donating H(L′) in 1−3 results
in a decrease of the RuII/RuIII potential. This is due to a
reduction of the overall charge of the complex cation from 2+
in [Ru(bpy)3]2+ to 1+ in 1−3, which provides electrostatic
stabilization of the oxidized RuIII-L′ species. Other RuN5O
chromophoric systems, [RuII(bpy)2(L’)]+, exhibit the RuII/
RuIII couple at 0.52−0.77 V.39 The similarity of the RuII/RuIII
potential of 1−3 with that of the N-salicylaldimine system
further supports the close ligand-field strengths of these classes
of complexes. Complexes 1−3 (Epa, 1, 1.84; 2, 1.83; 3, 1.80)
also exhibit a second irreversible oxidation wave. The oneelectron nature is confirmed by differential pulse voltammetry
(DPV). It could be due to either the RuII/RuIII couple or
oxidation of the ligand. However, the free ligands H(L1)−
H(L3) exhibit an irreversible oxidation wave [Epa: H(L1), 1.31
V; H(L2), 1.21 V; H(L3), 1.22 V], and their Epa values are
substantially lower (500−600 mV) than the ruthenium(II)
complexes. In addition, the potential difference between the
two successive oxidation couples is ∼1.2 V, which agrees well
with the average potential difference between the redox
couples of the ruthenium center (RuII/III − RuIII/IV ∼ 1.0−
1.5 V) observed in other mononuclear complexes.50 Therefore,
the second irreversible oxidation wave corresponds to the
RuIII/RuIV couple. Further, two successive quasi-reversible
ligand-based reductions are also observed for all of the
complexes, the one-electron nature of which is established by
the magnitude of the peak currents, which may involve the
addition of electrons in the electrochemically accessible lowest
unoccupied molecular orbital (LUMO)51 of the diimine
(−NCCN−) fragment of bpy.
Resonance Raman Spectra. The electrochemical and
electronic spectral data presented above strongly suggest that
the LUMO of 1−3 is localized at the bpy ligand. To further
substantiate these assignments, we carried out resonance
Raman experiments at 532 nm, which is preresonant with 491
nm and postresonant with 571 nm absorption bands of these
complexes. At this excitation wavelength, the modes associated
with both the bpy and Schiff-base ligands, H(L1)−H(L3),
become enhanced. Inspection of the resonance Raman spectra
(Figure S1) indicates no change in the pattern with an increase
in the number of methyl groups at the dangling pyridine
moiety, which might be expected because of their similar
structures. The high-energy features in the spectra at 1602 and
1554 cm−1 [both ν(CC)], 1480 cm−1 (the most intense
one), 1312 cm−1 [both ν(CN)], and 1270 cm−1 [ν(CC)
inter-ring] and peaks with a shoulder at 1164/1154 cm−1
[δ(CCH) in plane] and 1020/1036 cm−1 (ring breathing) are
ascribable to the internal modes of the coordinated bpy
ligands,52 and this is confirmed by the spectral shifts (20−30
cm−1) induced in these bands upon deuteration of bpy. In the
low-frequency end of the spectra, there are bands at 664 cm−1
[δ(CCC) inter-ring, isotope sensitive] and 374 cm−1; the
former has been tentatively associated with a liganddeformation mode of bpy and the latter with a symmetric
Ru−Nbpy stretching mode.53 The band at 374 cm−1 shows only
a weak resonance enhancement because the MLCT transition
exerts little influence on the Ru−Nbpy bond. On the basis of
these results, we conclude that the broad absorption with a
maximum near 491 nm is assigned as a RuII(dπ) → bpy(π*)
H6C (1-3) protons. Similarly, the complexes show a negative
c.i.s. values for H3C, H4C, H3D, H4D, H3F, and H5F (1-3),
H4F (1,2), and H6F (1) protons due to the magnetic
anisotropy induced by proximate ring current. In coordination
with Ru(II), the H3E (1) and H3E-H6E (2,3) protons signal
is shifted upfield due to Ru(II) to ligand π-back-donation,
which has a lesser influence on the H4E and H5E (1) protons.
The positive c.i.s. values observed for H3A and H3B (1-3)
protons are obviously due to van der Waals interaction.
Further, the positive c.i.s. values for H4A, H5A, H4B, H5B,
H5C and H5D (1-3) and H6E (1) protons suggest that the
ligand-to-metal σ-donation is more important (as this will
decrease the electron density at these sites leading to positive
c.i.s. values) than metal-to-ligand π-back-donation in the
ground state of these complexes.
Electronic Spectral Properties. The absorption spectra
of 2 and 3 are virtually identical with those obtained for
compound 1. These spectra are similar39 to that of [Ru(bpy)2(HNCH-C6H4O)]+ except for the presence of an
intense band at 342 nm, and the bands are slightly blue-shifted
with higher absorptivity. In the visible region, the complexes
exhibit two transitions at 491 and 371 nm. The lower-energy
transition (491 nm) is associated with a shoulder at lower
energy (571 nm). First, the two visible bands (491 and 371
nm) have been assigned based on the reported spectra of
[Ru(bpy)2]2+ complexes having other kinds of chelating
ligands.44,45 There are two different kinds of bipyridine π*acceptor orbitals involved in the dπ(RuII) → π*(bpy) metalto-ligand charge-transfer (MLCT) transitions, one symmetric
(χ) and one antisymmetric (ψ) concerning the C2 axis of the
ligand,44b,46 and the transition from metal-filled dπ orbitals to
these two π* orbitals results in the above-mentioned bands.
The lower-energy band at 491 nm is due to dπ(RuII) → π*(ψ)
and the higher-energy band at 371 nm to dπ(RuII) → π*(χ),
approximately 6580 cm−1 higher in energy than the former, as
expected. Second, an absorption maximum at 342 nm is
observed with nearly the same intensity (ε ≈ 10000 dm3 mol−1
cm−1) as that of the other two visible bands. This can be
interpreted as follows: (i) This may be due to a metal-centered
d−d transition similar to [Ru(bpy)3]2+ (340 nm); this
increased intensity is consistent with the lower symmetry
and is also likely to have a contribution from intensity
borrowing from other charge-transfer bands.47 (ii) This is
expected because transitions in the MLCT region are expected
to appear due to the dπ(RuII) → π*(imine) transition.48
Finally, the two intense bands observed at 245 and 295 nm are
characteristic of ligand-centered π−π* transitions and very
similar to the analogous absorptions of [Ru(bpy)3]2+. The
lowest-energy MLCT transition of [Ru(bpy)3]2+ appears at
450 nm;48 therefore, replacement of one bpy ligand by an
asymmetric ligand H(L1), H(L2), or H(L3) results in a red
shift of the same transition. The negative charge erected on the
metal by the strongly σ-donating phenolate moiety would be
predicted to destabilize the metal dπ orbital, reducing the dπ
→ π* MLCT band energy relative to [Ru(bpy)3]2+.
Redox Behavior. Complexes 1−3 display a redox wave
(E1/2: 1, 0.55 V; 2, 0.58 V; 3, 0.54 V versus Ag/AgCl) that
corresponds to the RuII/RuIII couple based on the magnitude
of the diffusion coefficients [D, (7.2−7.4) × 106 cm2 s−1]
calculated from cyclic voltammetry (CV) at different scan rates
(30−500 mV). The ipa versus ν1/2 plots, which pass through
the origin, are linear, indicating that the redox processes at the
electrode are diffusion-controlled. The peak potential separa2872
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Figure 3. Absorption spectra of 1 (A), 2 (B), and 3 (C) (3.0 × 10−5 M) in 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1 in the absence
(R = 0) and presence (R = 25) of increasing amounts of CT DNA.
Figure 4. Fluorescence quenching curves of EthBr bound to DNA in a 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1: (a) EthBr (1.25
μM); (b) EthBr + DNA (125 μM); (c−m) EthBr + DNA + 1 (A), 2 (B), and 3 (C) (0−10 μM).
DNA Binding Studies. Upon the incremental addition of
CT DNA to 1−3 (Table S1), the ligand-centered π → π*
absorption band (293 nm) of 1 shows a slight decrease in the
molar absorptivity (hypochromism: 1, ∼10%) with no red
shift, while 2 exhibits a slight increase and 3 displays a
significant increase in the molar absorptivity (hyperchromism:
2, ∼14%; 3, ∼42%) (Figure 3) at R = 25 (R = [DNA]/[Ru]),
suggesting a strong interaction between the ruthenium(II)
complexes and DNA. The spectral change might be interpreted
as due to groove binding of the adducts56 because 1−3
containing fused polyaromatic systems having coplanar atoms
(organic ligand) facilitates the formation of van der Waals
contacts or hydrogen bonds during interaction with the DNA
grooves. The intrinsic binding constant, Kb, values of 1 (6.97 ×
104 M−1), 2 (7.22 × 104 M−1), and 3 (7.55 × 104 M−1) are
almost the same (Figure S2), which is expected because of
their very similar molecular structures.57 These values are in
agreement with those of a well-established groove binding
rather than classical intercalation.58
Upon the addition of 1−3 (0−10 μM) to CT DNA
pretreated with EthBr ([EthBr]/[DNA] = 0.01) in a 2%
DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1 (Figure
4), the emission intensity at 595 nm of DNA-bound EthBr
decreases (% quenching: 1, ∼71%; 2, ∼68%; 3, ∼76%),
revealing that the ruthenium(II) complexes competitively
bound to CT DNA with EthBr. The observation of EthBr
fluorescence quenching due to the release of some EthBr
molecules from the EthBr/DNA system is supportive of the
interaction of 1−3 with CT DNA through groove binding.59
According to the Stern−Volmer equation, the relative binding
propensity (KSV: 1, 5.21 × 104 M−1; 2, 4.58 × 104 M−1; 3, 7.51
× 104 M−1) of the complex to CT DNA was determined from
MLCT transition. This was anticipated because the Schiff-base
ligands, H(L1)−H(L3), containing phenolate and imine
functionalities possess no empty low-lying energy levels.
These results indicate that the first reduction potential is
bpy-based and the first oxidation potential is metal-based,
which is in agreement with the electrochemical measurements.
Further, the high-energy shoulders observed at 1484 and
1316 cm−1 are assigned to ν(CN) stretching vibrations due
to the imine functionality of H(L1)−H(L3), and a weak lowfrequency feature at 428 cm−1 might best be considered as the
Ru−Nimine stretching mode. The weak feature at 1526 cm−1
and shoulders at approximately 1474 and 1606 cm−1, obscured
by the bpy features at 1602 and 1554 cm−1, are attributed to
the phenolate functionality because they are insensitive to
deuteration of the bpy moieties.54 Furthermore, bands
observed at 1422, 1388, and 1264 cm−1 and features at 622,
598, 524, and 478 cm−1 (1), 618, 572, 528, and 476 cm−1 (2),
and 622, 570, 530, and 470 cm−1 (3) are associated with the
phenolate functionality of H(L1)−H(L3). On the basis of a
comparison with other oxygen-bound complexes,55 the band in
the range 570−598 cm−1 is assigned to Ru−O stretching
vibrations. These features are distinctly enhanced by the
transition at 571 nm. These observations, in conjunction with
the simultaneous presence of bpy-centered modes, suggest that
the longest-wavelength absorbance in these complexes is
associated with an H(L)(π) → bpy(π*) interligand chargetransfer (ILCT) transition due to the loss of electron density
on the oxygen because the electron is transferred from the
oxygen lone pair of phenolate in H(L1)−H(L3) to π* of the
bpy units. The interpretation of these resonance Raman effects
would require separate normal-mode and molecular orbital
calculations, which have not yet been attempted.
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Table 3. Electrochemical Dataa for the Ruthenium(II) Complexes in the Presence and Absence of CT DNA
E1/2 (V)
complex
R
Epc (V)
Epa (V)
CV
DPVb
ΔEp (mV)
ipc/ipa
D (×106 cm2 s−1)
K2+/K3+
1
0
5
0
5
0
5
0.373
0.371
0.358
0.358
0.360
0.360
0.431
0.431
0.432
0.436
0.423
0.428
0.402
0.401
0.395
0.397
0.391
0.394
0.376
0.376
0.369
0.370
0.368
0.369
59
60
73
77
63
67
0.73
1.08
1.02
1.13
0.82
1.35
5.8
5.9
4.9
5.1
4.9
4.7
0.969
2
3
0.962
0.925
Measured versus standard calomel electrode. Scan rate: 50 mV s−1. Supporting electrolyte: 5 mM Tris-HCl/50 mM NaCl. Complex
concentration: 2.5 × 10−4 M. bDPV. Scan rate: 2 mV s−1. Pulse height: 50 mV.
a
Figure 5. Cyclic voltammograms of 1 (A), 2 (B), and 3 (C) (0.5 mM) in the absence (a) and presence (b) of CT DNA (R = 5) at 25.0 ± 0.2 °C
and a 50 mV s−1 scan rate in a 2% DMF/5 mM Tris-HCl/50 mM NaCl buffer at pH 7.1.
The redox potentials of the RuII/RuIII couple (E0′ or
voltammetric E1/2) for 1−3 (1, 0.402 V; 2, 0.395 V; 3, 0.391
V) are consistent with the general trends in E1/2 values in a
MeCN solution (cf. above). Upon the addition of CT DNA to
1−3 (R = 5), a drop in the peak currents of both anodic and
cathodic waves in the CV (Figures 5 and S5) and DPV
responses (Figure S6) is observed. The intensity of the peak
current (1, ∼14%; 2, ∼10%; 3, ∼21%) decreases with the
addition of DNA, and the decrease is higher for 3 than for
others, which is consistent with the above spectral studies. This
indicates the slower mass transfer of 1−3 bound to DNA
fragments,63 which leads to a decrease in the concentration of
the unbound redox-active species in solution. It can be
observed that 1−3 exhibit more or less the same electrochemical behavior in both the DNA-free and -bound
complexes, such as the formal potentials (E1/2) of the RuII/
RuIII couple (DNA-free/DNA-bound: 1, 0.402/0.401 V; 2,
0.395/0.397 V; 3, 0.391/0.394 V), peak potential separation
ΔEp (DNA-free/DNA-bound: 1, 59/60 mV; 2, 73/77 mV; 3,
63/67 mV), and the diffusion coefficient D (DNA-free/DNAbound: 1, 5.8/5.9 × 106 cm2 s−1; 2, 4.9/5.1 × 106 cm2 s−1; 3,
4.9/4.7 × 106 cm2 s−1), revealing that 1−3 interact with CT
DNA in a groove binding fashion. The values of K2+/K3+ were
determined using the Nernst equation and found to be in the
range of 0.925−0.969 for ruthenium(II) complexes, whose
potential shift is 1−3 mV. Interestingly, a K2+/K3+ value of
near-unity for 1−3 shows that they are involved in DNA
interaction, favoring both of the oxidation states equally.64
BSA Interaction. The BSA solution has a strong
fluorescence emission peak at 340 nm when excited at 280
nm. Therefore, the emission spectra of BSA (1 × 10−6 M) in
the presence of increasing concentrations of 1−3 [(1−4.5) ×
10−6 M] were recorded at 300 and 310 K (Table 4). The
fluorescence intensity of BSA decreased regularly (Figure 6),
the slope of a straight line (Figure S3) obtained from the plot
of the fluorescence intensity versus the complex concentration.60 The fluorescence quenching curve of EthBr-bound
CT DNA by 1−3 showed that quenching of the EthBr/DNA
system by 1−3 is in good agreement with the linear Stern−
Volmer equation, which also indicates that the complexes bind
to DNA. According to the equation K EthBr[EthBr] =
Kapp[complex], where KEthBr is 4.94 × 105 M−1,61 the
concentration of EthBr is 1.25 × 10−6 M and the concentration
of the complex is that used to obtain a 50% reduction of
fluorescence intensity of EthBr. The Kapp values are calculated
to be 8.82 × 104 M−1 (1), 8.23 × 104 M−1 (2), and 9.50 × 104
M−1 (3), and the apparent binding constant decreases in the
order 2 < 1 < 3, which is less than the binding constant of the
classical intercalators and metallointercalators (107 M−1). The
Kapp value is consistent with the Kb value obtained by the UV−
vis absorption spectral study, suggesting that the interaction of
1−3 with DNA is a groove binding mode.62
The circular dichroism (CD) spectrum of CT DNA was
monitored (Table S2) in the presence of 1−3 at a 1/R (=[Ru
complex]/[DNA]) value of 3; the positive band showed little
increase in the molar ellipticity (1, ∼14%; 2, ∼10%; 3, ∼18%),
and the negative band displayed a small decrease in the molar
ellipticity (1, ∼18%; 2, ∼12%; 3, ∼12%) with no red or blue
shift of the band maxima (Figure S4, curve b). These
observations are supportive of the groove binding mode of
interaction of 1−3, which is consistent with the results
obtained from the UV−vis absorption and fluorescence
spectral studies.
The CV and DPV responses were obtained for 1−3 on a
glassy carbon electrode in a 2% DMF/5 mM Tris-HCl/50 mM
NaCl buffer at pH 7.1 in the presence and absence of DNA,
and the well-behaved CV and DPV responses are used to
monitor the interaction of the complexes with DNA (Table 3).
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initiated by a dynamic mechanism but originates from the
formation of a ground-state complex, resulting from the static
quenching mechanism.68
Alternatively, the UV−vis absorption spectra of BSA in the
absence and presence of 1−3 were recorded (Figure 7) to
explore the structural changes of BSA and to investigate
formation of the BSA−complex system (a static quenching
process). The UV−vis spectrum of BSA has two main
absorption peaks. The strong absorption peak around 210
nm reflects absorption of the backbone of BSA, and the weak
absorption peak around 280 nm is due to the aromatic acid
residues.69 With the addition of 1−3, the intensity of the peak
at 210 nm decreased with an ∼6 nm (1), ∼8 nm (2), or ∼13
nm (3) red shift and the intensity of the peak at 280 nm
increased slightly. The results indicate that the interaction
between 1−3 and BSA leads to the loosening and unfolding of
the BSA backbone and an increase in the hydrophobicity of the
microenvironment of BSA. It is well-known that dynamic
quenching does not change the absorption spectrum, but
formation of the nonfluorescence ground-state complex can
change it,66 and thus the interaction between 1−3 and BSA
was mainly a static quenching process.
Therefore, the quenching data were analyzed according to
the modified Stern−Volmer equation.70 The modified Stern−
Volmer plots (Figure S8) reveal a good linear relationship, and
the corresponding effective quenching constant (Ka) increases
with rising temperature (Table 4), following the dependence of
KSV on the temperature as mentioned above. The results show
that the binding constants between the ruthenium(II)
complexes and BSA are moderate. It is noted that the binding
constant of 104−106 M−1 is acceptable for drug-carrier
complexes.71 Thus, the binding constants of 1−3 show that
BSA can be considered to be a good carrier for transfer of these
ruthenium(II) complexes in vivo.
For the static quenching interaction, if it is assumed that
there are similar and independent binding sites in the
biomolecule, the number of complex bound per protein (n)
and binding constant (Kb) can be calculated using a doublelogarithmic equation.72 As shown in Figure S9, the doublelogarithmic plot is a straight line. The values of Kb and n can be
obtained from the intercept and slope of the plot, respectively,
at two different temperatures (Table 4). The results reveal that
the values of the binding constants with BSA increase with
rising temperature, which is due to an enhancement in the
stability of the BSA−complex system and shows that the
binding process is an endothermic reaction.73 All of the
binding constants are medium and indicate that the interaction
between BSA and the ruthenium(II) complexes is moderate.
The values of n at the experimental temperatures are
approximately equal to 1, which indicates that there is just a
single binding site74 in BSA for 1−3. In addition, BSA has two
tryptophan residues that have intrinsic fluorescence. Trp-212 is
located within a hydrophobic binding pocket, and Trp-134 is
located on the surface of the molecule. A linear Stern−Volmer
plot is generally indicative of a single class of fluorophore, all
equally accessible to the quencher. These molecules do not
rapidly penetrate the hydrophobic interior of proteins, and
only those tryptophan residues on the surface of the protein
are quenched. Accordingly, three similar complexes most likely
bind to Trp-134 in BSA.75
The calculated values of ΔG°, ΔH°, and ΔS° at two different
temperatures are given in Table 4. The value of ΔG° is
negative, so the binding process is spontaneous. The values of
Table 4. Quenching, Association, Binding, and
Thermodynamic Parameters of the Interaction of 1−3 with
BSA at Different Temperatures
parameters
KSV (104 M−1) ±
SD
kq (1012
M−1 s−1)
Ka (104 M−1) ±
SD
Kb (104 M−1) ±
SD
n ± SD
ΔH° (kJ mol−1)
ΔS°
(J mol−1 K−1)
ΔG° (kJ mol−1)
KSV (104 M−1) ±
SD
kq (1012
M−1 s−1)
Ka (104 M−1) ±
SD
Kb (104 M−1) ±
SD
n ± SD
ΔH° (kJ mol−1)
ΔS°
(J mol−1 K−1)
ΔG° (kJ mol−1)
KSV (104 M−1) ±
SD
kq (1012
M−1 s−1)
Ka (104 M−1) ±
SD
Kb (104 M−1) ±
SD
n ± SD
ΔH° (kJ mol−1)
ΔS°
(J mol−1 K−1)
ΔG° (kJ mol−1)
300 K
R
310 K
[Ru(bpy)2(L1)]+ (1)
5.264 ± 0.002
0.9992
5.751 ± 0.002
5.264
R
0.9989
5.751
5.053 ± 0.005
0.9992
5.521 ± 0.004
0.9995
5.063 ± 0.098
0.9969
5.575 ± 0.100
0.9968
0.987 ± 0.017
77.951
90.303
0.994 ± 0.017
91.032
−27.013
−28.142
[Ru(bpy)2(L2)]+ (2)
4.018 ± 0.004
0.9988
4.739 ± 0.004
4.018
0.9998
4.739
4.619 ± 0.003
0.9996
4.899 ± 0.002
0.9998
4.203 ± 0.120
0.9952
4.431 ± 0.112
0.9961
0.982 ± 0.02
77.742
89.556
0.992 ± 0.01
90.037
−26.789
−27.833
[Ru(bpy)2(L3)]+ (3)
6.347 ± 0.005
0.9988
6.952 ± 0.001
6.347
0.9995
6.952
5.809 ± 0.004
0.9997
6.247 ± 0.010
0.9984
6.553 ± 0.077
0.9982
6.873 ± 0.082
0.9979
0.996 ± 0.014
77.830
91.462
92.058
−27.361
−28.460
Article
0.996 ± 0.014
up to 35% (1), 25% (2), and 49% (3) at 300 K and 38% (1),
30% (2), and 50% (3) at 310 K, accompanied by a blue shift of
7−9 nm. All of the Stern−Volmer plots65 (Figure S7)
represent a good linear relationship. As is known, linear
Stern−Volmer plots represent a single quenching mechanism,
either static (the ground-state complex formation between a
quencher and a fluorophore) or dynamic (a collisional
process).66 The value of the Stern−Volmer quenching
constant (KSV) is obtained from the slope of the plot F0/F
versus [Q] at different temperatures. The value of the
quenching rate constant (kq) is obtained from the equation
KSV = kqτ0. The result shows that KSV increases with rising
temperature (Table 4), indicating that the fluorescence
quenching of BSA by ruthenium(II) complexes is likely to
occur via a dynamic quenching mechanism. The maximum
value of kq of various quenchers with biological macromolecules is 1010 M−1 s−1.67 In the present study, the kq values
of the quenching processes (∼1012 M−1 s−1) are greater than
1010 M−1 s−1. These results suggest that the quenching is not
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Figure 6. Changes in the fluorescence spectra of BSA through titration with complexes 1−3 at 300 K (left, A) and 310 K (right, B). The
concentration of BSA is 1 × 10−6 mol L−1, and the concentration of 1−3 was varied from (a) 0.0 to (i) 4.5 × 10−6 mol L−1, with pH 7.4 and λex =
280 nm.
Figure 7. UV−vis absorption spectra of BSA in the absence and presence of 1 (A), 2 (B), and 3 (C): (a) Absorption spectrum of BSA. (b)
Absorption spectrum of BSA in the presence of 1−3 at the same concentration, [BSA] = [Cu complex] = 3.5 × 10−6 mol L−1. The absorbance of
1−3 is negligible in the spectral region shown.
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Figure 8. Agarose gel showing cleavage of 20 μM ΦX174 RF DNA incubated with 2.5 mM 1 (A), 2 (B), and 3 (C) in a 0.1 M phosphate buffer at
37 °C for 4 h. Lane 1: DNA control. Lanes 2−6: DNA + 1 (A), 2 (B), or 3 (C). (pH 8.0, 7.5, 7.0, 6.5, and 6.0, respectively.) Forms I and II are the
SC and NC forms of DNA, respectively.
Scheme 3. Protonation of the Dangling Pyridine Moiety of Schiff-Base Ligands (1−3), Postulated as Being Responsible for
pH-Dependent Damage to DNA
ΔH° and ΔS° are positive. The positive value of ΔH° shows
that the binding process is mainly an endothermic reaction.
The greater negative values of TΔS° compared to ΔH° reveal
that the formation of a complex between 1−3 and BSA is
mainly an entropy-driven reaction and TΔS° governs the
spontaneity of the reaction. The positive ΔH° and ΔS° values
indicate that the hydrophobic interaction is the main force in
the binding of 1−3 to BSA.
pH-Dependent DNA Cleavage. To assess the DNA
cleavage ability of the complexes, supercoiled plasmid ΦX174
RF DNA (20 μM) was incubated for 4 h with 1−3 (2.5 mM)
in the absence of an activator in a 0.1 M phosphate buffer, pH
6.0−8.0 in 0.5 step increments. When plasmid DNA is nicked
in its supercoiled form (form I), an open circular relaxed form
(form II) is generated, and with further cleavage, a linear form
(form III) is produced. When electrophoresis is performed on
the reaction mixture, the compact form I migrates relatively
faster, while the nicked form II migrates more slowly and the
linearized form (form III) migrates between forms I and II.
The ΦX174 RF DNA substrate used in agarose gel
electrophoresis is 95% supercoiled (SC), 5% nicked circular
(NC), and 0% linear circular, and their positions can be
distinguished on the gel (Figure 8, lane 1). The results show
that when DNA is incubated with 1−3 at pH 7.5 or 8.0 (Figure
8, lanes 2 and 3), the DNA migrates similarly to the DNA
substrate (SC, 95%; NC, 5%). However, at pH 7.0 (Figure 8,
lane 4), the SC form of DNA incubated with 1−3 is slightly
retarded (SC, 90%; NC, 10%) compared to the substrate DNA
(SC, 95%; NC, 5%). Interestingly, when the pH is reduced to
6.5 or 6.0 (Figure 8, lanes 5 and 6), all of the complexes show a
sudden increase in retardation because of the stronger cleavage
properties [pH 6.5/6.0: 1, SC (15/8), NC (85/90), LC (0/2);
2, SC (10/5), NC (90/90), LC (0/5); 3, SC (5/3), NC (90/
87), LC (5/10)]. This indicates that 1−3 convert form I into
form II and then into form III at pH 6.0, while at pH 6.5, 3
cleaves form I into form II and then into form III but 1 and 2
convert form I into form II alone. It was proposed that for the
dangling pyridine moiety in 1−3, which contains one nitrogen
atom with a pKa value of 6.84 (1, 2-aminopyridine), 7.60 (2, 2amino-6-picoline), or 9.50 (3, 2-amino-4,6-dimethylpyridine),
the nitrogen atom with a high pKa value could capture H+
strongly and enhance the interaction with negatively charged
DNA. It was found that at a pH >7 almost no damage to the
DNA is observed, whereas below pH 7, DNA damage is
prevalent. Because healthy cells grow at pH >7, typically pH
7.2, and (hypoxic) cancer cells have characteristically lower pH
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Figure 9. Agarose gel showing cleavage of 20 μM ΦX174 RF DNA incubated with 2.5 mM 1−3 in a 0.1 M phosphate buffer at 37 °C for 24 h (pH
6.5). Lane 1: DNA control. Lane 2: DNA + 1. Lane 3: DNA + 1 + NaN3 (500 μM). Lane 4: DNA + 1 + KI (500 μM). Lane 5: DNA + 1 + DMSO
(20 μM). Lane 6: DNA + 2. Lane 7: DNA + 2 + NaN3 (500 μM). Lane 8: DNA + 2 + KI (500 μM). Lane 9: DNA + 2 + DMSO (20 μM). Lane
10: DNA + 3. Lane 11: DNA + 3 + NaN3 (500 μM). Lane 12: DNA + 3 + KI (500 μM). Lane 13: DNA + 3 + DMSO (20 μM).
values, typically pH 6.8, we propose that 1−3 might selectively
target cancer cells. At the same time, we propose that the
dangling pyridine moiety in 1−3 could be protonated at lower
pH and in this form cause damage to DNA, as illustrated in
Scheme 3. The ability of 1−3 to exhibit pH-dependent
cleavage of DNA follows the order 3 > 2 > 1. Upon
replacement of the dangling pyridyl group in 1 by the 2methylpyridyl group as in 2 and the 4,6-dimethylpyridyl group
as in 3, there is an enormous increase in the electron density
on the dangling pyridyl nitrogen atom because of the presence
of electron-donating methyl group(s) in 2 and 3. This reveals
the importance of the electronic effect of a substituted methyl
group(s) in enriching the electron density on the dangling
pyridyl nitrogen atom of 2 and 3, thereby rendering the
abstraction of H+ more facile. On the other hand, the ability of
hydrophobic interaction of the complex depends on the
incorporation of a number of methyl groups in the dangling
pyridyl group. As a consequence, the increase in hydrophobic
interaction from 1 to 3 at a pH <6.8 can promote stronger
binding to DNA and raise the cleavage activity.
For the ruthenium(II) complexes 1−3, which show efficient
pH-dependent DNA cleavage, mechanistic studies were
performed at pH 6.5. DNA cleavage generally proceeds via
two major pathways: one is hydrolytic cleavage, and the other
is oxidative cleavage. The oxidative processes generally form
reactive singlet oxygen or hydroxyl radical species involving a
photoactive or a redox-active metal center, causing damage to
the sugar and/or base and resulting in the formation of
fragmented species that cannot be religated. Hydrolytic
cleavage in the absence of any external additives does not
suffer from such drawbacks because the cleaved products can
be religated enzymatically. To investigate the role of radicals in
DNA damage by 1−3, cleavage reactions were carried out
under aerobic conditions in the absence of an external reagent
by incubating 1−3 with DNA for 4 h in the presence of a
variety of radical scavengers like DMSO (hydroxyl radical),
NaN3 (singlet oxygen), and KI (superoxide). The results show
that DNA cleavage by 1−3 is not inhibited by any of the
classical radical scavengers (Figure 9). To ascertain the
mechanism of the DNA cleavage reaction by 1−3, form II
(NC) obtained from the cleavage of SC DNA has been
isolated, treated with a T4 ligase enzyme, and subjected to gel
electrophoresis.76 We have observed ∼73% (1), ∼78% (2), or
∼84% (3) conversion of form II to its original form I,
indicating that a hydrolytic mechanism dominates over other
mechanisms (Figure 10).
Cytotoxicity of Ruthenium(II) Complexes. The free
ligand H(L1)−H(L3) and complexes 1−3 have been screened
against cell lines of different cancer origins, viz., renal cancer
(A498), breast cancer (EVSA-T and MCF-7), nonsmall cell
lung cancer (H226), ovarian cancer (IGROV), melanoma
Figure 10. Analysis of the capacity of T4 DNA ligase to religate DNA
cleaved by 1−3. Lane 1, DNA control. Lane 2: products (NC ΦX174
RF) obtained from the reaction with complex 1. Lane 3: lane 2 + T4
ligase. Lane 4: products (NC ΦX174 RF) obtained from the reaction
with complex 2. Lane 5: lane 4 + T4 ligase. Lane 6: products (NC
ΦX174 RF) obtained from the reaction with complex 3. Lane 7: lane
6 + T4 ligase.
(M19), and colon cancer (WIDR). For comparison, the
cytotoxicities of six known anticancer drugs, viz., doxorubicin
(DOX), cisplatin (CPT), 5-fluorouracil (5-FU), methotrexate
(MTX), etoposide (ETO), and taxol (TAX), have also been
screened against all of the above cell lines (Table 5). Indeed, a
ligand or complex with an IC50 value higher than 10 μM is
considered to be inactive.77 The test results indicate that the
free ligands H(L1)−H(L3) are inactive (IC50, 95 to >100 μM)
to all of the cancer lines. However, chelation of free ligands
with ruthenium(II) increases the cytotoxicities of the cancer
cell lines. The cytotoxicities of 1−3 are insignificant for A498
and H226. Also, 1 shows low or no cytotoxicity at all in all of
the cell lines except EVSA-T. The IC50 values obtained with 2
and 3 are higher compared to those achieved with CPT in
IGROV, M19, MCF-7, and WIDR and with 5-FU in IGROV,
M19, and WIDR. In all of the cell lines, the cytotoxicities of 2
and 3 are even higher than those reached with DOX, MTX,
ETO, or TAX. Interestingly, 1−3 exhibit a similar potency
against EVSA-T, which is comparable to CPT. The ability of
the complexes to exhibit cytotoxicity follows the order 3 > 2 >
1 in all of the tested cell lines except EVSA-T. Earlier
investigations have shown that the lower cationic charge of the
complex and the hydrophobicity of the ligands confer greater
lipophilicity, which promotes complex permeation across the
cell membrane to exhibit cytotoxicity.78 The monocationic
charge of the complexes (1−3) and the hydrophobicity of the
dangling pyridine rings of the coordinated ligands [H(L1)−
H(L3)] might thus be attributed to the efficient cytotoxicity of
complexes, in addition to their DNA and protein binding
affinity. Further, the only structural difference among 1−3 is
the substitution of electron-donating methyl group(s) in the 6
position [H(L2) and 2] and 4 and 6 positions [H(L3) and 3]
of the dangling pyridine moiety of the Schiff-base ligand, and
the increasing hydrophobicity follows the order dangling
simple pyridine (1) < 6-methylpyridine (2) < 4,6-dimethylpyridine (3) (cf. above). A molecule with an elevated hydrophobicity also has an elevated lipophilicity (the lipophilicity is
directly proportional to the hydrophobicity), which means it
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Table 5. In Vitro Cytotoxicity Assays for 1−3, DOX, CPT, 5-FU, MTX, ETO, and TAX Screened against Seven Human Tumor
Cell Lines of Different Origins, viz., Renal Cancer (A498), Breast Cancer (EVSA-T and MCF7), Nonsmall Cell Lung Cancer
(H226), Ovarian Cancer (IGROV), Melanoma (M19), and Colon Cancer (WIDR)
druga (IC50/μM Using SRB as the Cell Viability Test)
cell line
1
2
3
DOX
CPT
5-FU
MTX
ETO
TAX
A498
EVSA-T
H226
IGROV
M19
MCF7
WIDR
50.811
2.913
32.400
16.601
10.020
10.905
14.523
48.510
2.514
19.913
8.699
5.599
5.701
8.020
42.520
1.810
15.823
7.010
4.110
4.412
5.701
0.166
0.015
0.366
0.110
0.029
0.018
0.020
7.509
1.406
10.895
0.563
1.860
2.330
3.223
1.099
3.651
2.613
2.283
3.397
5.764
1.729
0.081
0.011
5.033
0.015
0.051
0.040
<0.007
1.803
0.435
5.397
0.780
0.693
3.551
0.206
<0.004
<0.004
<0.004
<0.004
<0.004
<0.004
<0.004
a
IC50 = concentration of the drug required to inhibit the growth of 50% of the cancer cells.
can permeate the cell membrane more easily. Therefore, the
incorporation of methyl group(s) in the dangling pyridyl group
enhances the cytotoxicity of the complex from 1 to 3, and a
subtle change in the ligand framework could contribute
significantly to the cytotoxicity of the complex.
grams, normal and modified Stern−Volmer plots of
BSA, double-logarithmic plot of the quenching effect,
absorption and fluorescence spectral properties, CD
spectral parameters, and experimental details (PDF)
■
Accession Codes
CONCLUSIONS
A new series of ruthenium(II) complexes of salicylaldiminetype ligands containing a dangling pyridine have been
synthesized and characterized. The newly synthesized complexes (1−3) are obtained in the crystalline state, and thus the
structures of the complexes [Ru(bpy)2(L1/L2/L3)]+ possess a
distorted octahedral geometry and exist as a dimer due to π−πstacking and C−H···π interactions. They are involved in
noncovalent interaction through the groove binding mode with
DNA. DNA binding studies such as absorption, emission, CD
spectral, and electrochemical studies suggest that 3 exhibits the
highest binding affinity among the present complexes. It is
noteworthy that these complexes show a higher propensity for
binding to BSA protein in the hydrophobic region, which will
be helpful to understand the drug−protein interaction.
Remarkably, the ruthenium(II) complexes hydrolytically cleave
SC DNA into NC and linear forms in the absence of any
external reagent. Interestingly, they also show pH-dependent
DNA damage, indicating that DNA is destroyed at the pH
typical of hypoxic tumor cells, while little or no damage is
identified at the pH distinctive of healthy cells. Such behavior
is ascribed to the fact that (i) the dangling pyridine moiety can
be protonated at low pH and the protonated form is the active
agent or (ii) higher hydrophobicity encourages DNA binding
and enhances the cleavage activity. All of the ruthenium(II)
complexes involved in noncovalent DNA binding are cytotoxic
to the human EVSA-T breast cancer cell line. Their
cytotoxicity is comparable to CPT, which is presently used
to treat breast cancer. Thus, the ruthenium(II) complexes of
salicylaldimine-type ligands containing a dangling pyridine are
new and improved DNA- and BSA-binding and DNA-cleaving
agents and have the capability of being developed as effective
cytotoxic drugs for the treatment of human breast cancer.
■
CCDC 1567540, 1567542, and 1567544 contain the
supplementary crystallographic data for this paper. These
data can be obtained free of charge via www.ccdc.cam.ac.uk/
data_request/cif, or by emailing data_request@ccdc.cam.ac.
uk, or by contacting The Cambridge Crystallographic Data
Centre, 12 Union Road, Cambridge CB2 1EZ, UK; fax: +44
1223 336033.
■
AUTHOR INFORMATION
Corresponding Author
Mariappan Murali − Coordination and Bioinorganic
Chemistry Research Laboratory, Department of Chemistry,
National College (Autonomous), Tiruchirappalli 620001
Tamil Nadu, India; orcid.org/0000-0001-8669-5939;
Phone: +91-431-2482995; Email: murali@nct.ac.in,
ma66mu@gmail.com; Fax: +91-431-2481997
Author
Somasundaram Sangeetha − Coordination and Bioinorganic
Chemistry Research Laboratory, Department of Chemistry,
National College (Autonomous), Tiruchirappalli 620001
Tamil Nadu, India
Complete contact information is available at:
https://pubs.acs.org/10.1021/acs.inorgchem.1c03399
Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS
We are grateful for the DST-FIST initiative of the National
College (Autonomous), Tiruchirappalli, India. Thanks are due
to STIC, Cochin University of Science and Technology, for the
X-ray crystal structure data, and Dr. Babu Varghese, SAIF,
Indian Institute of Technology Madras, for structure solution
and refinement. The authors thank Professor A. Ramu, School
of Chemistry, Madurai Kamaraj University, for CD spectral
measurements. This paper is dedicated to Professor G.
Mugesh, Department of Inorganic and Physical Chemistry,
Indian Institute of Science, Bangalore, India for his magnificent
contributions in the field of Chemical Biology, Bioinorganic
and Medicinal Chemistry.
ASSOCIATED CONTENT
sı Supporting Information
*
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acs.inorgchem.1c03399.
Resonance Raman spectra, plots of [DNA] versus
[DNA]/(εa − εf), I0/I versus [complex]/106, and ipc
versus ν1/2, CD spectra, differential pulse voltammo2879
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Article
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■
NOTE ADDED AFTER ASAP PUBLICATION
This paper was published on January 31, 2022. Due to
production error, an incorrect version of Figure 4 was included.
The corrected version was reposted on February 2, 2022.
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