← Back
Alteration of steric hindrance modulates glutathione resistance and cytotoxicity of three structurally related Ru(II)-p-cymene complexes.
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton
Transactions
View Article Online
PAPER
Cite this: Dalton Trans., 2016, 45,
8541
View Journal | View Issue
Alteration of steric hindrance modulates
glutathione resistance and cytotoxicity of three
structurally related RuII-p-cymene complexes†
Kallol Purkait,‡ Saptarshi Chatterjee,‡ Subhendu Karmakar and Arindam Mukherjee*
The effect of steric hindrance on reactivity towards biomolecules while designing RuII-η6-p-cymene
based anticancer agents seems to be an important parameter in improving the activity and inducing
resistance against glutathione (GSH) deactivation. Herein we present the structure, hydrolysis, anticancer
activity and the effect of steric hindrance on deactivation by glutathione for three complexes, [RuII(η6-pcym)(L1)(Cl)](PF6) (1), [RuII(η6-p-cym)(L2)(Cl)](PF6) (2) and [RuII(η6-p-cym)(L3)(Cl)](PF6) (3). The ligands
L1–L3 are Schiff bases which show increasing substitution in a benzene ring, such that two ortho hydrogens are replaced by -methyl in 2 and by -isopropyl in 3. The cytotoxicity results strongly suggest that
Received 8th December 2015,
Accepted 5th April 2016
controlling the rate of hydrolysis through tuning of steric hindrance may be a feasible pathway to derive
DOI: 10.1039/c5dt04781a
GSH resistant anticancer agents. The cellular studies show that all the three complexes show good blood
compatibility (haemolysis <3%) and induce cellular death through caspase activation via the mitochondrial
www.rsc.org/dalton
pathway. They have anti-angiogenic activity and prevent the healing of treated cells.
Introduction
Metallo drugs have found wide use as anticancer agents.
Improving the design of a metal based anticancer agent has
certain advantages viz. higher number of accessible coordination numbers, reversible mode of binding ability with
different kinds of electron donor species, accessible oxidation
states under normal physiological conditions1 and choice of
suitable ligands. All the above factors tune the properties of
the metal complexes.2,3 Platinum complexes are the predominant contributors among the metal based drugs4 despite
several major drawbacks in the form of severe side effects and
induced resistance to several cancer types. During the past few
decades researchers have focused on non-platinum complexes
among which ruthenium has emerged as a significant
contributor.5–8 Ruthenium complexes are octahedral or tetrahedral unlike the case of PtII complexes which are generally
square planar, and the tuning of electronic properties shows
variation since they bear differences in electronic configur-
Department of Chemical Sciences, Indian Institute of Science Education and
Research Kolkata, Mohanpur Campus, 741246, India.
E-mail: a.mukherjee@iiserkol.ac.in
† Electronic supplementary information (ESI) available: Table for crystal refinement parameters, NMR, and UV-visible spectra of hydrolysis, plots of cell viability, cell cycle, JC-1. CCDC 1431153 and 1431154. For ESI and crystallographic
data in CIF or other electronic format see DOI: 10.1039/c5dt04781a
‡ These authors contributed equally to this work.
This journal is © The Royal Society of Chemistry 2016
ation and geometries. Among the promising ruthenium complexes, certain RuIII-complexes, NAMI-A, NKP1339 have moved
up to clinical trial phase-II.9,10 Ru-complexes may be delivered
to cells aided by plasma proteins11 and they may get reduced
to RuII inside the cell in the presence of cellular reducing
agents viz. glutathione, cysteine, and ascorbate.12–16 However,
under hypoxic conditions of tumor17 these reducing agents
can simultaneously inhibit those drugs by forming stable
bonds with the metal centre18,19 although activation of the
drugs due to glutathione is also reported.20 The literature data
reveals that DNA is a potential site of action for Ru complexes21,22 while other possibilities like, inhibition of the
active site of an enzyme8,23 are found to be alternative modes
of action for RuII complexes. One reason for interest in Rucomplexes is their relatively higher stability towards aquation
which is supposed to decrease the loss of effective dosage due
to the formation of active species before reaching the
target16,24,25 and this may minimize side effects compared to
Pt-complexes. In addition Ru is also thought to be interfering
with iron metabolism.11,26
Several ruthenium(II) complexes are evaluated in vitro
against various cancerous cell lines, especially the [RuII(η6arene)(L)Cl] class of complexes (where L = bidentate chelating
ligand) which shows activity against primary tumour
cells.2,14,27,28 The library of work from Sadler et al. shows that
the activity of these complexes can be tuned by changing the
ligand (L) and arene.28,29 To cite an example the [RuII(η6-arene)
(diamine)Cl] family of complexes show that their anticancer
Dalton Trans., 2016, 45, 8541–8555 | 8541
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
activity is related to the structure, hydrolytic ability,30 DNA
binding31 and also deactivation by thiolate sulfur.32 A different
generation but related to the above type of complex is the
RAPTA series of complexes in which a phosphorus based
monodentate ligand developed by Dyson et al. shows antimetastatic activity.33 Recent findings show that these RuII-anticancer agents have strong affinity for the thiolate sulfur of
cysteine,34 and glutathione.32 Elevated levels of the glutathione
(GSH) synthetase and other thiol based enzymes viz. γ-glutamylcysteine ligase, and γ-glutamyl-transpeptidase are
observed in tumour cells35,36 which are rendered as major contributing factors for chemoresistance37 and their depletion
may increase the sensitivity of the drugs and oxidative stress.38
Since Ru has an affinity for sulfur hence Ru-complexes may be
deactivated by the above enzymes, GSH and other cellular
thiols.32,39,40
Hence complexes that are not deactivated by GSH or show
less affinity towards GSH and other cellular thiols would be of
high significance provided they are cytotoxic. We recently have
shown that complex 3 shows in vitro cytotoxicity in certain
cancer cell lines and there is little deactivation in the presence
of glutathione under hypoxic conditions.41 In addition no GSH
binding was observed by NMR studies. We have synthesized
two more isostructural complexes where the only difference
lies in the substitution at the vicinity of the metal centre so as
to correlate with the effect of steric hindrance on the hydrolysis, anticancer activity, glutathione binding and the pathways
of cytotoxicity which is strongly related to the breaking and
making of new bonds with the metal centre leading to the
pattern of reactivity.
Results
Syntheses and characterization
The Schiff base ligands (L1–L2) were synthesized using a
similar procedure as described by us earlier.41 The reaction of
6
one mole equivalent of [RuII
2 (η -p-cym)2Cl4] precursor with ca.
2.4 mole equivalents of the respective ligand in methanolic
solution led to the formation of respective complexes
(Scheme 1). The complexes were crystallized using 1 equivalent
of ammonium hexafluorophosphate due to their residual positive charges. All of the ligands and complexes were well characterised by 1H, 13C, HMQC and DEPT 135 NMR, FT-IR, UV-Vis
Scheme 1 Synthetic scheme and chemical structure of ligands (L1–L3)
and metal complexes (1–3). (a) Amine (1 mmol), aldehyde (1 mmol),
EtOH, HCO2H (0.5%), reflux, dark, 12 h. (b) (i) Ligand (1.2 mmol), [Ru2(η6p-cym)2Cl4] (0.5 mmol), MeOH, reflux, dark, 4 h. (ii) NH4PF6.
8542 | Dalton Trans., 2016, 45, 8541–8555
Dalton Transactions
spectra and single crystal X-ray structure. The bulk purity was
confirmed by elemental analysis.
X-ray crystallography
The single crystals of complexes 1–2 were obtained by layering
petroleum benzene over a dichloromethane solution of the
respective complex. They were structurally characterized by a
single crystal X-ray diffraction method.
Complex 1 crystallizes in the space group P21/n (ESI,
Table S1†). The crystal structure shows that the RuII is bound
with two nitrogens of L1, one chloride and a η6-bonded
p-cymene ring. In imidazole as usual the sp2 hybridized
double bonded N-atom is the donor. The structure displays
that the benzene and imidazole rings are not in the same
plane (Fig. 1). In 1, the methyl group of p-cymene is on the
same side as that of chlorine, the isopropyl group of p-cymene
seems to be moving away from the benzene ring (Fig. 1). An
octahedral [PF6]− is present outside the coordination sphere
and the Ru is in +2 oxidation state. The average Ru–N bond
distance is 2.097 ± 0.024 Å and the Ru–Cl distance is 2.416(9)
Å (ESI, Table S2†). The distances between the carbons of the
p-cymene and the RuII are not uniform as described in ESI,
Table S2.†
ˉ (ESI,
Complex 2 crystallizes with the space group P1
Table S1†). Similar to 1 the RuII in 2 is bound to two nitrogens
from L2, a chloride and the p-cymene (in a η6-fashion).
However, unlike 1 the isopropyl group of the p-cymene is
oriented towards the chloride rather than the methyl group.
The average Ru–N distance is 2.105 ± 0.039 Å and the Ru–Cl
bond shows a distance of 2.410(7) Å (ESI, Table S2†). The bond
distances between the carbon atoms of the p-cymene ring and
RuII are not uniform (ESI, Table S2†). The structure of complex
3 which has been described elsewhere showed similar Ru–N
and Ru–Cl distances however the orientation of the chloride
was like 1.41
Lipophilicity
To determine the distribution coefficient we used the standard
shake flask method.42 The log Do/w values as depicted in Fig. 2,
show that 3 has the highest lipophilicity (3.2 ± 0.1) whereas,
complexes 1 and 2 show log Do/w of 0.42 ± 0.12 and 0.66 ± 0.1
Fig. 1 Single crystal structures of complexes 1 and 2 with 50% probability level. All hydrogen atoms, counter anions and solvent molecules
are omitted for clarity.
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
Paper
Fig. 2 Representative diagram of lipophilicity of 1–3 represented by a
bar diagram showing comparative log Do/w values of complexes in the
octanol–aqueous phosphate buffer system, where error bars in the
graph indicate the standard deviation in measurement. For 3 data was
obtained from ref. 41.
respectively. Since among complexes 1–3, complex 1 shows
quicker hydrolysis, hence we use the term distribution coefficient as a standard practice.
Hydrolysis study
To gain better insight into the properties of 1–3 we performed
hydrolysis studies under various conditions depicted in
Table 1 (a more elaborative table is present in the ESI,
Table S3†). The results show that under acidic conditions the
rate of hydrolysis increases, leading to decrease in solution
half-life and in the presence of higher chloride concentration
(40 and 110 mM) the half-life increases. At pH 6.7 the complexes show a higher rate of hydrolysis in 4 mM chloride containing buffer compared to that observed for water (Table 1,
ESI, Table S3 and Fig. S13–S15†). The time dependent 1H NMR
study in the 7 : 3 v/v D2O–DMSO-d6 mixture shows evidence of
formation of hydrolyzed species after 1 to 3 h (Fig. 3) for 1 and
2. The hydrolysis study using 1H NMR (7 : 3 v/v D2O–DMSO-d6,
110 mM chloride) shows excellent stabilities for up to 10 days
for all the complexes (ESI, Fig. S16†). In the NMR studies 1–3
do not show any peak corresponding to the dissociation of
p-cymene from the metal complexes up to 23 h (beyond which
we did not check anymore) even when the 7 : 3 v/v D2O–DMSOd6 mixture with no chloride was used.
Binding studies
CT-DNA binding. The DNA binding ability was probed using
UV visible spectroscopy (ESI, Fig. S17 and S18†). The com-
Table 1
Fig. 3 Hydrolysis study of complexes 1–2 in 3 : 7 v/v DMSO-d6 and D2O
mixtures measured by 1H NMR at 25 °C with time. * indicates the peak
corresponding to the hydrolysis product.
plexes were hydrolyzed for 12 h before addition of DNA so that
the spectral variations due to hydrolysis are minimum, and in
another case a fresh solution of complexes were used. We
observed similar results in both cases (ESI, Table S4†). Here,
we present the data which involves DNA binding after 12 h.
The results show that complex 2 binds more strongly than
1 or 3. The binding ability shows the order 2 > 1 > 3. The DNA
binding constants are 3.04 (0.1) × 104 and 4.11 (2.9) × 103 M−1
for 1 and 3 respectively but for 2 it is higher in magnitude
(1.11 (0.3) × 105 M−1) (ESI, Table S4, Fig. S17 and S18†).
Glutathione binding. The binding with reduced L-glutathione is monitored by 1H NMR (ESI, Fig. S19 and S20†) as
well as ESI mass spectrometry, for both complexes 1 and 2
(Fig. 4, ESI, Fig. S21 and S22†). 1H NMR study of 1 and 2 with
2 molar excess of GSH was monitored up to 8 hours and the
results show no binding for each case. The only new peaks
that are identified are of the autooxidation product leading to
the GSH dimer and hydrolyzed species (ESI, Fig. S19 and 20†).
But when we probed the concentration dependent binding
with excess GSH using ESI-MS, we found the mono-GSH
adduct of the complexes at a higher concentration of GSH (at
least 25 equivalents) for both 1 and 2. It appears that –SH
Half-lives (t1/2) of complexes 1–3 at pH 7.4 and 6.7 and at various chloride concentrations
t1/2 (h)
pH
Chloride conc. (mM)
7.4
4
40
4
40
—
110a
6.7
Water
—
a
1
2
3
2.79 ± 0.66
3.91 ± 0.91
1.18 ± 0.36
2.55 ± 0.18
2.01 ± 0.24
Stable for 10 days
5.67 ± 0.91
22.22 ± 2.13
4.37 ± 0.31
9.98 ± 2.90
7.56 ± 1.04
Stable for 10 days
17.75 ± 0.02
49.40 ± 1.90
8.07 ± 1.1
23.96 ± 2.98
11.70 ± 2.48
Stable for 10 days
Detected by 1H NMR.
This journal is © The Royal Society of Chemistry 2016
Dalton Trans., 2016, 45, 8541–8555 | 8543
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
Dalton Transactions
Fig. 4 ESI MS speciations of complexes 1 and 2 upon treatment with
excess glutathione showing the proposed speciation as per the m/z
peaks obtained. Complex 3 did not provide any GSH bound species,
hence not shown.
groups of GSH bind to the Ru-centre in the complexes rendering m/z of 714.12 and 742.23 for 1 and 2 respectively (Fig. 4,
ESI, Fig. S21†). Another GSH bound fragmented peak with loss
of p-cymene from 2, at m/z 542.13, was identified. The isotopic
distribution ratio confirms the speciation (ESI, Fig. S21 and
S22†). At this concentration it was not possible to obtain any
significant NMR data. The intensity of the mono-GSH adduct
peak was more for 1 compared to 2. In contrast even at the 100
equivalent concentration of GSH, no GSH bound peak was
obtained for 3 even with ESI-MS.
dependent on the conditions and cell lines. Although the
toxicity of 1–3 is relatively less in A549 but under hypoxic conditions a definite increase in activity is observed, except for the
case of 2 which exhibits deactivation for MCF-7 in hypoxia.
The deactivation is confirmed from the statistical significance
of the normoxia and hypoxia dataset. In addition, it is
encouraging to find that 1 and 2 are marginally less toxic to
the primary human foreskin fibroblast (HFF-1). Based on
hypoxia and normoxia data, 1 shows more cytotoxicity against
MiaPaCa-2 and HepG2 compared to HFF-1. However their
studies in the presence of GSH in hypoxia shows that 1 and 2
are not as effective when excess GSH is present, as the required
dosage increases by 1.5 to 2 fold. Complex C1 of Sadler et al.
showed less activity in the probed cell lines compared to 1–3,
which was reported earlier by us for comparison.41 C1 also
exhibits better activity in hypoxia although the activity is
decreased in the presence of excess GSH in hypoxia.41
Cell cycle analysis. The effect of complexes (1, 2) on the cell
cycle of MCF-7 was probed with two different concentrations
(4 and 6 μM) of each complex and the observation shows that
the percentage of cell population in the S and G2/M phases are
comparatively higher with respect to the control (Table 3, ESI,
Fig. S28 and S29†), whereas for 3, we found an increase in the
sub G1 population along with the G2/M phase arrest.41
DNA ladder assay. Genomic DNA fragmentation, known as
the hallmark of apoptosis,43 was observed in the ladder assay
Cellular studies
Cell viability assay. The cytotoxic activity of the synthesized
compounds (1, 2 and 3) against various cell lines are presented
in Table 2 in the form of IC50 values. All the complexes show a
potent cytotoxicity profile against the tested cell lines when
compared to the in vitro activity of cisplatin. Under normoxic
conditions, complexes 1 and 3 are more promising against
HepG2 while 2 has the best activity against MCF-7. Complexes
1–3 are found to be least effective against A549 and are in
general efficient towards MCF-7, MiaPaCa-2 or HepG2. To
probe their relative activity in hypoxia we used MCF-7 and
A549. We found that the toxicity profile of 1–3 shows variation
Table 2
Table 3
Cell cycle inhibition of MCF-7 using 1 and 2a
DMSO control
1, 4 μM
1, 6 μM
2, 4 μM
2, 6 μM
Sub G1
G0/G1
S
G2/M
5.19
4.07
1.66
2.42
3.62
57.8
50.6
50.09
46.71
43.39
21.62
26.33
24.96
25.41
28.88
15.96
19.69
24.02
26.03
24.91
a
Cells were treated for 24 h with 1 and 2. Cells were treated with
propidium iodide and analyzed by FACS. Cell populations were
analyzed and expressed as the percentage of cells in each phase. The
data presented is an average of two independent experiments.
Cytotoxicity of complexes 1–3 in comparison to [RuII(η6-p-cym)(en)Cl](PF6) (C1) and cisplatin
IC50 (µM) ± S.D.a
Hypoxiab
Normoxia
1
2
3
C1 f
Cisplatin
Hypoxia + glutathioned
MCF-7
A549
HFF-1
MIA PaCa-2
Hep G2
NIH 3T3
MCF-7
A549
MCF-7
A549
9±1
7±1
13.8 ± 1 f
43.9 ± 3.6
15 ± 1
21 ± 1
19 ± 1
23.2 ± 1 f
36.7 ± 2.3
24 ± 1
10.2 ± 1
10.5 ± 1
6±1
N.D.c
N.D.c
7±1
12 ± 1
10 ± 1
N.D.c
13.8 ± 5
6±1
10 ± 1
8±1
N.D.c
14.3 ± 1
10 ± 1
11 ± 1
9.2 ± 1
N.D.c
7±1
9±1
11 ± 1
9.1 ± 1 f
31.7 ± 1.7
19 ± 2
18 ± 1
17 ± 1
15.6 ± 1 f
31.2 ± 1.5
27 ± 1
16 ± 1
15 ± 1
10.8 ± 1 f
49.3 ± 1.8
29 ± 2e
37 ± 1
27 ± 1
16.7 ± 1 f
31.7 ± 2.6
40 ± 2e
a
IC50 values were calculated by non-linear curve fitting in dose response inhibition–variable slope model using Graph pad prism. S.D. = standard
deviation. The data presented are mean of three independent experiments, in a single experiment each concentration was assayed in triplicate.
The statistical significance (P) of the data is >0.001 to <0.05. b Hypoxia (1.5% O2). c Not determined. d With 1 mM of reduced L-glutathione.
e
20 molar equivalents of reduced L-glutathione used with respect to IC50 dosage of the respective cell line in hypoxia. f Data was obtained from
ref. 41.
8544 | Dalton Trans., 2016, 45, 8541–8555
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
Fig. 5 Agarose gel image of DNA ladder formation due to apoptosis
using the MCF-7 cell line. (A) 50 bp step ladder, (B) DMSO control, (C) 1
(8 μM), (D) 1 (10 μM), (E) 2 (8 μM) and (F) 2 (10 μM) treated for 24 h.
Paper
Fig. 7 Representative diagram for caspase-7 activation of MCF-7 after
24 h treatment of 1–3 with three different concentrations (4, 6 and
8 μM) of each. Activities were shown in terms of pNA released. Error
bars show standard deviations in data.
for all of the three complexes. Multiple bands (i.e., ladder)
were visible on agarose gel electrophoresis of genomic DNA
isolated from the MCF-7 cells treated with 1 and 2 (Fig. 5).
Optical microscopy imaging
Cell swelling and nuclear fragmentation were observed for
both the cases upon treatment of complexes 1 and 2 on MCF-7
(ESI, Fig. S33†). Our earlier reports on 3 suggest that the observation is similar for all the three complexes.
ICP-MS studies
To know the amount of ruthenium inside the cells we used the
ICP-MS study. We obtained the ruthenium concentration
inside the cell after 24 h incubation of the complex treated
MCF-7 cells (with the respective complex) and the results show
that the concentration of Ru inside the cells for complex 3 is
more than that for 1 and 2 (Fig. 6). Complex 3 also happens to
be the most lipophilic and slowest in hydrolysis among the
three.
Caspase-7 activation
The caspase-7 activation study of all the complexes (1, 2 and 3)
shows activation of caspase-7 in MCF-7 upon treatment of the
respective complexes (Fig. 7). The colorimetric data based on
Fig. 8 Representative diagram for change of mitochondrial membrane
potential upon treatment of complexes (1–3) on MCF-7 stained by JC-1.
the cleavage of the p-nitrophenol from the DEVD sequence
shows that 1 and 2 have almost similar activities and the
caspase-7 activity appears to be more in the case of complex 3
as the absorbance obtained is higher.
Mitochondrial membrane potential change
When MCF-7 cells were treated with four different concentrations of 1–3, we observed a significant change in mitochondrial transmembrane potential (MMP). The change is
maximum for complex 3 and minimum for 2 (Fig. 8, ESI,
Fig. S30–S32†) using the same concentration of each complex
although the dose dependence on IC50 is not the same for the
complexes, rather complex 3 requires higher dosage to achieve
IC50.
Wound assay (migration)
Fig. 6 Plot shows the amount of ruthenium present inside the cells (in
ppb) with respect to the control experiment of complexes 1–3. Each
data is the mean of three independent experiments.
This journal is © The Royal Society of Chemistry 2016
The effects of 1–3 on cancer cell migration in the MCF-7 cell
line were assessed using a scratch wound healing assay. From
Fig. 9 it can be seen that the untreated control cells rapidly
migrate over time, resulting in almost 45% healing in 12 h and
>90% healing at 24 h of incubation. While for 1–3 the wide
denuded area remained un-healed indicating the antiproliferative
Dalton Trans., 2016, 45, 8541–8555 | 8545
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
Dalton Transactions
Fig. 10 Ex ovo Chick Embryo Angiogenesis (CEA) assay depicting the
anti-angiogenic potential of complexes 1–3. The zoomed area is of the
dotted white box upon treatment after 4 h.
Fig. 9 Analysis of migration of MCF-7 after treatment with the corresponding complexes. The images were taken after 12 h and 24 h of drug
loading.
Discussion
nature of the cells, attained by the treatment with the respective complexes (1–3).
Haemolysis assay
Interaction of the drug with the blood components particularly
RBCs is an important and inevitable phenomenon, thus assessing the haemolysis becomes crucial in evaluating the blood
compatibility of drugs.44,45 The results (ESI, Fig. S34 and
Table S5†) show that compounds 1, 2 and 3 show good compatibility with RBCs. The haemolysis was found to be least for
complex 1 it was ca. 1% at a concentration of 15 μM. For
complex 2 at the same concentration the haemolysis was
ca. 1.5% whereas it was the highest for complex 3 but less than
3% (ESI, Fig. S34†).
Chick embryo angiogenesis assay (CEA)
The anti-angiogenic potential of the compounds was evaluated
by the chick embryo angiogenesis (CEA) assay (Fig. 10). The
results indicate the anti-angiogenic properties of 1–3 as degenerate and damaged blood vessels were observed on the chick
embryo (affected zone is marked with white box and damaged
blood vessels are marked with black arrows) after 4 h of
exposure to 1–3. No new blood vessels were formed during the
period of treatment. However the control shows healthy blood
vessels and new blood vessels were formed during the 4 h
incubation period.
8546 | Dalton Trans., 2016, 45, 8541–8555
Half sandwich RuII-complexes bearing the formulation
[RuII(η6-p-cym)(L1)(Cl)](PF6) (1) and [RuII(η6-p-cym)(L2)(Cl)]
(PF6) (2) along with some new studies on our previously
reported [RuII(η6-p-cym)(L3)(Cl)](PF6) (3) are discussed here. All
the complexes are well characterized by NMR and single
crystal X-ray structure. The X-ray structures of the complexes
show that all of them have a chloride and one η6-bonded
p-cymene ring. The geometric orientation rendered shows
differences in terms of steric bulkiness in the vicinity of the
metal centre (isopropyl to methyl to hydrogen). Complex 3 displays maximum steric hindrance and 1 shows the least. The
hydrolysis results show that the solution half-life has increased
with increase in steric hindrance (Table 1; t1/2 trend: 3 > 2 > 1)
rendering better stability and changes the cytotoxicity pattern
as discussed later (Table 2).
The hydrolysis is also affected by the pH and concentration
of the leaving anion (e.g. Cl−) as well as the overall ionic
strength of a solution (Table 1). Although, the complexes
hydrolyze faster in pH 6.7 but due to the steric hindrance the
trend remains similar as mentioned earlier. The effect of
chloride concentration change seems to be the least on the
half-life of 1 at both the pH values. The trend in hydrolysis
does not show uniform increase with increase in steric hindrance. However, the effect of better stabilization at higher
chloride concentration is prominent. It is encouraging to find
that at extracellular concentration of chloride (∼110 mM), all
the three complexes are stable for at least 10 d (we did not
probe beyond 10 d). In the case where there was no chloride
added viz. the hydrolysis performed in 7 : 3 (v/v) D2O–DMSO-d6,
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
by 1H NMR, showed that the complexes started to hydrolyze
after 1–3 h but none of them lose any other ligand than Cl− up
to 23 h (beyond which it was not probed) (Fig. 3). The ESI-MS
data also supported the loss of chloride since we could
obtain the peaks corresponding to the general formulation
[RuII(η6-p-cym)(L-H)]+ at m/z values of 407.09, 436.18 and
490.15 for 1–3 respectively which matches well with the simulated values of 407.09, 436.13 and 490.18 for 1–3 respectively.
The CT-DNA binding studies showed that the strongest interaction is exhibited by complex 2 (Kb = 1.11(3) × 105 M−1),
although 2 is not the least hindered complex. This suggests
that the presence of the ortho methyl groups on the benzene
ring of the L2 leads to enhanced complex DNA interaction,46
which increases the DNA binding ability. The similarity in the
DNA binding constants for the 12 h hydrolyzed sample and
the fresh solution of complexes may be attributed to the slow
hydrolysis, since the timescale of the DNA binding experiment
is smaller (1.5 h). The p-cymene complexes of RuII show
similar binding constants with DNA as reported by Garcia
et al.47–50
Aquation is thought to be a major step which may be
involved in activation of this type of metal complex. However,
extracellular aquation may prevent the internalization of the
complexes due to their di-positive charge and higher hydrophilicity. Since, the lipid bi-layered cell membrane has a hydrophobic interior and hydrophilic exterior surface, hence if a
complex has to enter the cell through the diffusion pathway
then the lipophilicity is an important parameter. The partition
co-efficient values of 1 and 2 (0.42 and 0.66 respectively) are
significantly less than 3 (log D = 3.2), which clearly tells us that
the complexes (1 and 2) may not very efficiently cross the cell
membrane by a simple diffusion process. However, the cytotoxicity studies described later shows that both 1 and 2 are in
general more cytotoxic than 3. The cellular internalization of 3
is the highest in the series as per the ICP-MS studies, which
signifies that the effective cytotoxic concentration is lower for 1
and 2. The results suggest that the steric hindrance may
also prevent 3 from reacting with the relevant biomolecules
resulting in relatively less cytotoxicity.
However, we have reported earlier that complex 3 is
very effective in hypoxia under an excess concentration of
GSH. The dose dependence in hypoxia is also better than the
normoxic IC50 dosage as per the in vitro studies. Cytotoxicity
data of all the three complexes show that 3 is the most
effective one under hypoxia in the presence of excess GSH.
Complexes 1 and 2 are very effective under normal circumstances without added excess GSH and also in hypoxia,
except in MCF-7 for 2 (Table 2). When excess GSH is present
in the surroundings, which may be thought of as a case
similar to many resistant forms of cancer, than the dose
dependence worsens for 1 and 2. The required dosage in the
presence of ca. 750–1000 equivalent excess GSH escalates to
almost double as compared to the general normoxia/hypoxia
IC50 dose.
It is well known that A549 expresses higher GSH levels than
normal lung fibroblasts.51 All the complexes also show poorest
This journal is © The Royal Society of Chemistry 2016
Paper
activity in A549 (Table 2). However, 3 requires the higher
dosage but does exhibit better activity in hypoxia in the
presence of excess extracellular GSH suggesting its resistance.
However, it should be noted that inside the cell there is
glutathione-S-transferase which catalyzes the conjugation of
GSH to similar complexes for detoxification hence the higher
dosage required. However, in hypoxia 3 seems to overcome
that and shows better IC50 in the entire family which is
encouraging.
Complex 1 may be considered to be the most effective
candidate against HepG2 and MIA PaCa-2 followed by MCF-7.
However, it is most susceptible to GSH deactivation. Complex
2 although showing a greater DNA binding constant is less
effective than 1 in most of the cell lines except for MCF-7,
where it shows the best activity in the series (Table 2). The
cytotoxicity data of the complexes suggests that by the change
of two ortho substitutions in L1–L3 (H in L1, methyl in L2 and
isopropyl in L3) the activity of the complexes gets affected in
terms of mechanism and pathways as described later. During
our previous work, we found that the effect of hypoxia
may also be applicable to other known potential RuII based
anticancer agents41 viz. the [(RuII(η6-p-cymene)(en)Cl)(PF6)]
designed by Sadler et al.52 [(RuII(η6-p-cymene)(en)Cl)(PF6)]
showed better activity in hypoxia against MCF-7 and A54941
but the required IC50 dosage is much higher for these cell
lines. However, the encouraging part here is that RuII may have
potential as a hypoxic active anticancer agent. Based on the
higher degree of aquation at acidic pH, it may be speculated
that complexes 1–3 have the potency to be more active inside
tumors due to their lower interstitial pH.53
The activity comparison of complexes 1–3 against primary
cell lines, primary human foreskin fibroblast (HFF-1) and
mouse embryonic fibroblast (NIH 3T3) shows that complex 1
is more active against HepG2 and MIA PaCa-2 than HFF-1 and
NIH 3T3 which is encouraging (Table 2). The clinical drug cisplatin is twice more cytotoxic to the primary cell line NIH 3T3
than any of the probed cancer cell lines in vitro. However, the
in vitro cytotoxicity of complex 3 against NIH 3T3 is comparable to its activity against HepG2 or MIA PaCa-2, which is still
better than the cytotoxicity pattern depicted by cisplatin in
these cell lines.
The NMR studies to probe the binding of GSH by 1–3 due
to the known inhibitory properties of GSH32 showed that
under NMR conditions, where less excess of GSH is used (to
obtain a good signal to noise ratio for both bound and
unbound species), there is no binding of GSH by any of the
complexes. However when we probed the GSH adduct formation by ESI-MS using more excess of GSH (around 25–100
equivalent excess) we found that with ca. 25 equivalents of
excess GSH, there is GSH adduct formation by complexes 1
and 2. Under the same conditions of experimentation, the
signal intensity of the GSH adduct is much higher for 1 as
compared to 2. However using up to 100 equivalents of GSH
we did not obtain any ESI-MS signal of the GSH adduct with 3
supporting our earlier reported NMR studies and the cytotoxicity data.41
Dalton Trans., 2016, 45, 8541–8555 | 8547
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
The bulkiness of the ligands is in the order of 3 > 2 > 1
which corroborates well with the glutathione mediated deactivation under hypoxic conditions (1 > 2 > 3). Complex 3
having isopropyl, the most bulky group, appears to exhibit
higher resistance to GSH. The slowest rate of hydrolysis may
be the reason behind such activity. As a trend, complex 3 is
least deactivated by GSH under hypoxia, followed by complexes
2 and 1. Complex 1 showed 77% and 100% increase in IC50
values in MCF7 and A549 respectively as a result of GSH
mediated deactivation under hypoxia. The deactivation by
extracellular glutathione is encouraging and suggests that
steric hindrance exhibits a significant role against deactivation
by GSH. The steric hindrance also affects the partition coefficient of the complexes rendering 3 more lipophilic hence
ICP-MS studies show more internalization of 3 inside cells
(Fig. 6) but yet 1 is more effective emphasizing that 1 may be
a better anticancer agent for low GSH expressing hypoxic
tumors.
The cell cycle inhibition studies show that there may be
variation in the mechanistic pathway of action since, 1 and 2
arrest MCF-7 cells at the G2/M and S phases whereas 3 arrests
in the G2/M and sub G1 phases. The arrest in the sub G1
phase is indicative of the apoptotic pathway for cytotoxicity by
3.54,55 However, it should be noted that apoptosis does not
always render arrest of the sub G1 phase.56,57 The DNA ladder
assay which provides a qualitative indication to apoptosis confirms that all the complexes follow the apoptotic pathway. In
addition, the microscopy images also suggest apoptosis since
there is membrane blebbing, cell swelling as well as nuclear
fragmentation for all the complexes (ESI, Fig. S33†). Apoptotic
cell killing involves extrinsic and intrinsic pathways. The
intrinsic pathways lead to greater change in mitochondrial
membrane potential. Complex 3 in spite of a poorer IC50
shows the highest change in MMP followed by complex 1 and
the least change in MMP is observed with 2. Hence, the mitochondria mediated intrinsic apoptotic pathway may be least
feasible in 2. In addition, the caspase activation trend by 2 and
1 although less than 3 is still high and suggests that perhaps
extrinsic pathways of apoptosis are active for them. The overall
trend of caspase-7 activation and MMP change shows that, it
cannot be ruled out that there are multiple pathways active,
especially in the case of 1 and 2.
Inhibition of migration of cancer cells is taken as a positive
property to prohibit invasion and metastasis.58 The effects of
1–3 on cancer cell migration in the MCF-7 cell line showed
that for complexes 1–3 the wide denuded area remained unhealed after 24 h of exposure indicating the antiproliferative
nature of the cells, attained by the treatment of the
respective drugs. Whereas, the untreated control cells rapidly
migrated over time due to the metastatic property resulting
in >90% healing. The chick embryo angiogenesis (CEA) assay
is a well known angiogenesis assay.59,60 The results showed
degenerate blood vessels with 1–3 treated embryos compared
to the normal development of blood vessels in the control,
which is indicative of the anti-angiogenic potential of the
complexes.
8548 | Dalton Trans., 2016, 45, 8541–8555
Dalton Transactions
Experimental section
Materials and methods
The solvents used for this work were dried and distilled as per
the literature mentioned procedure.61 Aniline (Merck), 2,6-dimethylaniline (Sigma-Aldrich), imidazole-2-carboxaldehyde
(Sigma-Aldrich), formic acid (Merck), ruthenium(III) chloride
(Precious Metal Online, Australia), ammonium hexafluorophosphate (Sigma-Aldrich), ethidium bromide (SRL, India),
agarose (molecular biology grade) (SRL, India), MTT [(3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide)] (USB)
and other cell growth media and their supplements (Gibco),
reduced L-glutathione (GSH) (Aldrich) were used without
6
further purification. [RuII
2 (η -p-cym)2(Cl)4] was synthesized
62
using a known procedure. The spectroscopy grade solvents
were used for spectroscopy and lipophilicity, and the NMR solvents were purchased from Cambridge Isotope Laboratories,
Inc. (CIL). Fertilizable eggs were purchased from a recognized
local poultry farm and incubated at 37 °C at the 60% humidity
level to grow the embryo.
SECOR India melting point apparatus was used to detect
the melting point and the average value of three data are
reported without standard deviation. A Perkin Elmer Lambda
35 spectrophotometer and a cary 300 UV-Visible spectrophotometer were used for UV-Visible experiments. FT-IR spectra
were recorded on a Perkin-Elmer Spectrum RX I spectrometer
in the solid state using KBr pellets. Either a JEOL ECS
400 MHz or a Bruker Avance III 500 MHz spectrometer was
used to obtain 1H, proton decoupled 13C, HMQC and DEPT
135 NMR spectra, each spectrum was recorded at 25 °C and
the values are reported in ppm. A Perkin-Elmer 2400 series II
CHNS/O analyzer was used for elemental analysis. Electrospray ionization mass spectra were recorded in +ve mode electrospray ionization using a Q-Tof micro™ (Waters) mass
spectrometer. The recrystallization yields of isolated products
are only reported. The synthesized ligands and complexes were
dried in a vacuum and stored in desiccators in the dark under
a nitrogen atmosphere. The synthetic procedure of 3 was
recently reported by our group, hence not described here.41
The well known [RuII(η6-p-cym)(en)Cl](PF6) (C1) reported by
Sadler et al. was synthesized according to the reported procedure and the analytically pure complex41 was used to study
the cytotoxicity in normoxia and hypoxia due to its known
toxicity profile.52 Human blood was collected from a volunteer
and all the work was performed by following the institute
ethical guidelines.
Synthesis
General procedure for synthesis of L1 and L2. 1 mmol of
imidazole-2-carboxaldehyde was dissolved in methanol at
30 °C followed by addition of 1 mmol of the respective amine
to the reaction mixture and refluxed. After 12 h the reaction
mixture was cooled at room temperature and evaporated to
dryness, an off white coloured solid mass of product was
formed. Finally the purification was done by washing the
obtained mass several times with diethyl ether. This procedure
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
produced ligands in a satisfactorily pure form which was used
for complex syntheses. The analytical data of L1 and L2 are
given below. L3 has been reported earlier by us.41
Synthesis of N-((1H-imidazol-2-yl)methylene)aniline (L1). L1
was synthesized using the above mentioned general procedure.
Yield: 0.164 g (95%), Anal. Calc. for C10H9N3: C, 70.16; H, 5.30;
N, 24.54; Found: C, 70.05; H, 5.33; N, 24.50%, mp: 126 °C,
1
H NMR (400 MHz, CDCl3, 25 °C): δ 8.54 (s, 1H, CHvN), 7.43
(t, J = 6.44 Hz, 2H, Ar–H), 7.33 (t, J = 5.36 Hz, 1H, Ar–H), 7.27
(d, J = 12.96 Hz, 2H, Ar–H), 7.25 (s, 2H, Imi-H) ppm (ESI,
Fig. S1†); 13C NMR (100 MHz, CDCl3, 25 °C): δ 149.9 (imine-C),
149.5 (Imi-C), 145.2 (Ar–C), 129.5 (Imi-CH), 127.1 (Ar–CH),
121.1 (Ar–CH), 115.4 (Ar–CH) ppm (ESI, Fig. S2†); ESI-HRMS
(methanol) m/z (calc.): 194.07 (194.07) [C10H9N3·Na+], FT-IR
(KBr pellets, cm−1): 2914, 1625, 1440; UV-Vis: [CH3CN, λmax,
nm (ε/dm3 mol−1 cm−1)]: 220 (6967), 296 (11 262), 317 (11 669).
Synthesis of N-((1H-imidazol-2-yl)methylene)-2,6-dimethylaniline (L2). L2 was synthesized using the above mentioned
general procedure. Yield: 0.180 g (90%), Anal. Calc. for
C12H13N3: C, 72.33; H, 6.58; N, 21.09; Found: C, 72.25; H, 6.62;
N, 21.02%, mp: 121 °C, 1H NMR (400 MHz, CDCl3, 25 °C):
δ 8.24 (s, 1H, CHvN), 7.13 (s, 2H, Imi-H), 7.08 (d, J = 6 Hz, 2H,
Ar–H), 7.02 (t, J = 5.2 Hz, 1H, Ar–H), 2.13 (s, 6H, CH3) ppm
(ESI, Fig. S3†); 13C NMR (100 MHz, CDCl3, 25 °C): δ 153.2
(imine-C), 135.8 (Imi-C), 128.6 (Ar–C), 128.4 (Imi-CH), 127.9
(Ar–CH), 125.3 (Ar–CH), 118.2 (Ar–CH), 18.4 (CH3) ppm (ESI,
Fig. S4†); ESI-HRMS (methanol) m/z (calc.): 222.10 (222.10)
[C12H13N3·Na+], FT-IR (KBr pellets, cm−1): 2910, 1638, 1442;
UV-Vis: [CH3CN, λmax, nm (ε/dm3 mol−1 cm−1)]: 282 (18 744),
323 (3871).
General procedure for synthesis of complexes 1 and 2
The complexes were synthesized according to a procedure
similar to that reported by us earlier.41 Precisely, 2.4 mmol
solution of the respective ligand in methanol (MeOH) was
6
added to the methanolic solution of [RuII
2 (η -p-cym)2(Cl)4]
(0.612 g, 1 mmol) in the dark under a nitrogen atmosphere at
room temperature. Then the reaction mixture was stirred for
10 h. NH4PF6 (0.326 g, 2 mmol) was added to the resultant
reaction mixture and stirred for another 2 h. Evaporation of
the reaction mixture led to an orange coloured mass which
was washed twice with diethyl ether. The crude product was
purified by crystallization from a dichloromethane solution
layered with petroleum benzene.
[RuII(η6-p-cym)(L1)(Cl)](PF6) (1). Yield: 0.352 g (60%), Anal.
Calc. for C20H25ClF6N3OPRu: C, 39.71; H, 4.17; N, 6.95; Found:
C, 39.67; H, 4.12; N, 7.02%, mp: 206 °C, 1H NMR (500 MHz,
DMSO-d6, 25 °C): δ 8.46 (s, 1H, imine-H), 8.15 (s, 1H, Imi-H),
7.83 (d, 1H, J = 0.5 Hz, Imi-H), 7.75 (d, 2H, J = 7 Hz, Ar–H),
7.59 (t, 2H, J = 7.5 Hz, Ar–H), 7.52 (t, 1H, J = 7.5 Hz, Ar–H),
6.00 (d, 1H, J = 6 Hz, p-cym-H), 5.65 (d, 1H, J = 6 Hz, p-cym-H),
5.58 (d, 1H, J = 6 Hz, p-cym-H), 5.47 (d, 1H, J = 6 Hz, p-cym-H),
2.46 (m, 1H, J = 7 Hz, iPr-H), 2.09 (s, 3H, p-cym-CH3), 0.99 (d,
3H, J = 7 Hz, iPr-CH3), 0.83 (d, 3H, J = 7 Hz, iPr-CH3) ppm (ESI,
Fig. S5†); 13C NMR (125 MHz, DMSO-d6, 25 °C): δ 154.8
(imine-C), 151.9 (Imi-C), 146.2 (Ar–C), 133.7 (Imi-CH), 129.3
This journal is © The Royal Society of Chemistry 2016
Paper
(Ar–CH), 129.0 (Ar–CH), 123.9 (Imi-CH), 122.4 (Ar–CH), 103.7
( p-cym-C–CH3), 101.3 ( p-cym-C-iPr), 85.2 ( p-cym-CH), 84.6
( p-cym-CH), 83.7 ( p-cym-CH), 83.0 ( p-cym-CH), 30.4 (iPr-CH),
21.9 (iPr-CH3), 21.3 (iPr-CH3), 18.2 ( p-cym-CH3) ppm (ESI,
Fig. S6†); ESI-HRMS (methanol) m/z (calc.): 442.06(442.06)
[C20H23ClN3Ru+], 406.10(406.10) [C20H22N3Ru+]; FT-IR (KBr
pellets, cm−1): 3436, 2972, 2929, 1627, 1439, 841. UV-vis:
[CH3CN, λmax, nm (ε/dm3 mol−1 cm−1)]: 318 (12 301), 385
(2962).
[RuII(η6-p-cym)(L2)(Cl)](PF6) (2). Yield: 0.340 g (55%), Anal.
Calc. for C22H27ClF6N3PRu: C, 42.97; H, 4.43; N, 6.43; Found:
C, 42.85; H, 4.38; N, 6.49%; mp: >250 °C, 1H NMR (500 MHz,
DMSO-d6, 25 °C): δ 8.51 (s, 1H, imine-H), 8.17 (d, 1H, J = 1 Hz,
Imi-H), 7.84 (d, 1H, J = 1.5 Hz, Imi-H), 7.31 (d, 2H, J = 5.5 Hz,
Ar–H), 7.27 (t, 1H, J = 5.5 Hz, Ar–H), 5.73 (d, 1H, J = 6.5 Hz,
p-cym-H), 5.52 (d, 2H, J = 7 Hz, p-cym-H), 5.27 (d, 1H, J = 6 Hz,
p-cym-H), 2.64 (m, 1H, J = 3.5 Hz, iPr-H), 2.31 (s, 3H, p-cymCH3), 2.17 (s, 3H, Ar–CH3), 2.06 (s, 3H, Ar–CH3), 1.06 (d, 3H,
J = 5 Hz, iPr-CH3), 1.04 (d, 3H, J = 4.5 Hz, iPr-CH3) ppm (ESI,
Fig. S9†); 13C NMR (125 MHz, DMSO-d6, 25 °C): δ 160.0
(imine-C), 150.7 (Imi-C), 145.9 (Ar–C–N), 133.4 (Imi-CH), 131.1
(Ar–C–CH3), 129.0 (Ar–C–CH3), 128.8 (Ar–CH), 128.5 (Ar–CH),
127.6 (Ar–CH), 124.4 (Imi-CH), 105.4 ( p-cym-C–CH3), 99.0
( p-cym-C-iPr), 85.7 ( p-cym-CH), 84.7 ( p-cym-CH), 83.4 ( p-cymCH), 83.4 ( p-cym-CH), 30.5 (iPr-CH), 22.2 (Ar–CH3), 21.4 (Ar–
CH3), 19.7 ( p-cym-CH3), 18.2 (iPr-CH3), 17.9 (iPr-CH3) ppm
(ESI, Fig. S10†); ESI-HRMS (methanol) m/z (calc.): 470.09
(470.09) [C22H27ClN3Ru+], 434.12 (434.12) [C22H26N3Ru+],
FT-IR (KBr pellets, cm−1): 3430, 2971, 2927, 1634, 1436, 846.
UV-vis: [CH3CN, λmax, nm (ε/dm3 mol−1 cm−1)]: 276 (6006), 303
(7635), 342 (3871).
X-ray crystallography
Single crystals of 1 and 2 suitable for diffraction were coated
with Fomblin oil and fixed on a loop and mounted on the
goniometer under flow of nitrogen. The data were collected in
a SuperNova, Dual, Cu at zero, Eos diffractometer, Agilent
using a monochromatic Mo Kα source (λ = 0.71073 Å) using a
detector distance of 75 mm at 100 K with exposure of
6 s. CrysAlisPro, Agilent Technologies, Version 1.171.37.31
(release 14-01-2014 CrysAlis171.NET) software was used for
data collection and data reduction. All the structures were
solved with Superflip63 structure solution programme Charge
Flipping and refined with the ShelXL64 refinement package
using least squares minimisation, using Olex2.65 All the hydrogen atoms were calculated and fixed using ShelXL after hybridization of all non-hydrogen atoms.66 Crystals data were
deposited at https://deposit.ccdc.cam.ac.uk. The CCDC
numbers are 1431153 for 1 and 1431154 for 2.
The ORTEP diagrams were processed through the POV-ray
with 50% probability level. All the non-hydrogen atoms were
refined with anisotropic displacement parameters.
Lipophilicity
Distribution coefficient (log D) was determined using the standard shake-flask method42 for all ligands and complexes in
Dalton Trans., 2016, 45, 8541–8555 | 8549
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
the octanol–aqueous phosphate buffer (20 mM, pH 7.4)
system. After overnight pre-equilibration of octanol and
aqueous phosphate buffer (equal volume, 3 mL each)67 the
solid samples (1.2 mg each) were added to the solvent mixture
and shaken constantly for 10 hours at 37 °C on a shaking incubator to obtain full distribution between the two layers.
Followed by this, all the tubes were centrifuged and left undisturbed for an hour. Aliquots of the aqueous and octanol
layers were pipetted out separately and the absorbances were
measured with a UV-Vis spectrophotometer using proper
dilution. Each set was performed in triplicate, concentration
of the substances in each layer was calculated using the
respective molar extinction coefficients and the distribution
coefficient values (log D) were obtained from the ratio.
Aquation studies: dependence on pH and chloride
concentration
The pH dependent aquation of 1, 2 and 3 was measured by
UV-Vis absorption spectroscopy at pH 7.4 and 6.7 (using 1%
methanol–20 mM phosphate buffer in 40 mM or 4 mM NaCl)
for 12 h. The hydrolysis of the complexes follows the pseudofirst order kinetics with respect to water and the half-life as
well as the rate were obtained from monoexponential decay
fitting at a constant wavelength with time.
Hydrolysis of respective complexes was also monitored by
1
H NMR by obtaining spectra at different intervals of time in
DMSO-d6 and D2O mixtures at 25 °C (3 : 7 v/v).
Stability in saline solution
The stability of each complex in extracellular chloride concentration (110 mM) was observed by 1H NMR at 25 °C in
D2O : DMSO-d6 (7 : 3 v/v), 110 mM saline solution. 2 mg of
each complex was dissolved in 640 µL of 7 : 3 (v/v) D2O/DMSOd6 solution containing 3.5 mg of NaCl and the spectra were
recorded up to ten days.
Interaction with CT DNA
Deoxyribonucleic acid sodium salt isolated from calf thymus
(CT DNA) was dissolved in 50 mM Tris-NaCl solution at pH 7.4
and kept for 10 h at 4 °C. The UV-Visible spectrum showed two
absorption bands at 260 nm and 280 nm and their absorption
ratio was 1.9, which indicate that the DNA was apparently free
from protein.68 The concentration of the DNA was determined
from the absorption value at 260 nm (molar extinction coefficient is 6600 dm3 mol−1 cm−1)69 and the average value of the
concentration was taken after three independent measurements of the same stock solution. The stock solution of the
DNA was stored at 4 °C and used within four days.
Interaction of 1 and 2 with CT DNA was measured with the
help of UV-Vis spectroscopy in Tris-HCl buffer (50 mM,
pH 7.4)–acetonitrile (99 : 1 v/v) media. Acetonitrile was used for
the preparation of the stock solution of complexes. Spectroscopic titrations were carried out at room temperature (25 °C).
The concentration of 1 and 2 for the binding experiments was
fixed to 1 × 10−4 M and incubated for 12 h at 25 °C for prehydrolysis. In another set of experiments the concentrations
8550 | Dalton Trans., 2016, 45, 8541–8555
Dalton Transactions
were fixed to 5 × 105 and 4.4 × 105 M for 1 and 2 respectively
and binding titration started instantaneously after preparing
the complex solution. The change in absorbance at 311
and 300 nm for 1 and 2 respectively was monitored with
subsequent addition of an aliquot of 5 μL (concentration of
1 × 10−2 M and 5.3 × 10−3 M for 12 h and 0 h of hydrolysis
respectively) of CT DNA into the sample and also into the
reference cuvette after 5 min of equilibration. The titration was
continued until there was no significant change in absorbance
for at least three successive additions.
Binding studies with reduced L-glutathione (GSH)
The binding studies of 1 and 2 with reduced L-glutathione
were monitored by 1H NMR. The samples were prepared in a
degassed D2O/DMSO-d6 (7 : 3 v/v) mixture at 25 °C under a
nitrogen atmosphere to minimize the autooxidation of glutathione. Each experiment involved 0.002 mmol of complex and
0.004 mmol (2 equiv.) of GSH dissolved in the solution.
The binding was studied up to 29 hours at different intervals
of time although after 8 h, the total autooxidation process was
completed.
Binding studies were also carried out using ESI mass spectrometry. Lower equivalents of GSH viz. up to 10 equiv. did not
produce any m/z peak corresponding to the adduct with the
respective complexes. Hence, higher amounts viz. at least
25 equiv. and up to 100 equivalents of GSH with the complex
was incubated for half an hour at 37 °C under stirring conditions, diluted with water and ESI mass spectrometry was
performed.
Cell lines and culture
The human breast adenocarcinoma (MCF-7), human lung
adenocarcinoma (A549) and mouse embryonic fibroblast (NIH
3T3) were kindly donated from the Department of Biological
Sciences, IISER Kolkata (originally purchased from ATCC);
hepatocarcinoma (Hep G2) and human pancreatic carcinoma
(MIA PaCa-2) were obtained from NCCS, Pune, and Primary
Human Foreskin Fibroblast (HFF-1) was donated to us by
Dr Rupak Dutta, Department of Biological Sciences, IISER
Kolkata. Cell lines (MCF-7, A549, MIA PaCa-2) were maintained
in the logarithmic phase in Dulbecco’s Modified Eagle
Medium (DMEM) while HFF-1 and Hep G2 were grown in
Minimum Essential Medium (MEM). Both the media were supplemented with 10% fetal bovine serum (GIBCO) and antibiotics (100 units per ml penicillin and 100 mg per ml
streptomycin). The cell culture conditions used was 95%
humidity and 5% CO2 at a temperature of 37 °C. For the
hypoxic cell culture the oxygen level was maintained at 1.5%
and other aforementioned parameters were kept unaltered.
Cell viability assay
MTT assay was used to determine the cytotoxicity of the compounds against the cancer cells based on cell viability as an
indicator for the sensitivity of the cells to the individual compound. Briefly, exponentially grown cancer cells were seeded
in a 96-well microplate (Nunc) at a density of 6 × 103 viable
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
Paper
cells per well. Cells were incubated at 37 °C (5% CO2 atmosphere) for 48 h to resume exponential growth followed by
removal and replacement of fresh media containing the
desired concentration of test compounds. Each concentration
was loaded in triplicate. The stock solutions of compounds
were prepared in DMSO such that the concentration of DMSO
in the well does not exceed 0.2%. After 48 h of subsequent
incubation, compounds containing media were removed, followed by addition of 200 µL of fresh medium and 20 µL of
1 mg per ml MTT in PBS ( pH 7.2) into each well. On incubation of the plates for 3 h at 37 °C under a humidified 5%
CO2 atmosphere, MTT was allowed to form formazan crystals
in metabolically active cells. Finally the media were removed
after incubation and replaced with 200 µL of DMSO in
each well that would solubilize the formazan crystals. The
absorbance for each well was recorded at 515 nm using a
BIOTEK ELx800 plate reader.
IC50 values represent the drug concentration at which 50%
cells are inhibited compared to control, which were calculated
by fitting nonlinear curves in GraphPad Prism 5, constructed
by plotting cell viability (%) versus log of drug concentration in
µM or in nM. Each independent experiment was carried out in
triplicate.
For the experiment under hypoxic conditions the oxygen
percentage of the CO2 incubator (ESCO cell culture CO2
incubator, model: CCL-170T-8-UV) was maintained at 1.5%.
Compounds were loaded in 96 well microplates in a level-II bio
safety cabinet under atmospheric conditions which took
ca. 10 minutes and then the drug loaded 96 well microplate
was placed in the incubator programmed to attain 1.5%
oxygen concentration. The incubator takes ca. 30–40 minutes
to reach the 1.5% oxygen level.
were seeded in a 90 mm dia petri-dish in 12 mL DMEM and
incubated for 48 h at 37 °C under a 5% carbon dioxide atmosphere. Subsequently the media were replaced with fresh
media containing the required concentration of test compounds and incubated at 37 °C under a 5% carbon dioxide
atmosphere for 24 h. Cells were then harvested by trypsinization followed by centrifugation at 2000 rpm for 5 minutes.
Cells were subjected to PBS ( pH = 7.2), washed and resuspended in 500 μL lysis buffer [20 mM Tris-HCl ( pH 7.4),
0.4 mM EDTA, 0.25% Triton-X 100] and incubated for
15 minutes at room temperature. Lysed cells were centrifuged
at 14 000 rpm for 10 minutes and the supernatant was collected. This was mixed with a 1 : 1 (v/v) phenol–chloroform
mixture (1 mL) and the aqueous layer was carefully separated
to which 55 μL of 5 M NaCl and 500 μL isopropanol were
added respectively following incubation overnight at −20 °C.
After that the solution was centrifuged at 14 000 rpm and
washed with 70% ice cold ethanol and dried in air which was
again resuspended in 1× TE solution [10 mM Tris-HCl
( pH 8.0), 1 mM EDTA] containing RNase (150 μg mL−1) and
finally it was incubated at 37 °C for 20 minutes. The supernatant, containing isolated DNA was mixed with bromophenol
blue dye and loaded in 1.6% agarose gel containing
EtBr (1.0 μg mL−1) and run at 60 V for around 3 h in 1× TBE
(Tris-borate-EDTA) buffer. Untreated cells were used as controls whereas a 50 bp DNA ladder was used to track
the migration of bands and fragmentation sizes on the
agarose gel.
Visible bands on the gel were observed and a picture taken
on exposure to UV light through the gel documentation system
of Bio-Rad.
Cell cycle analysis
MCF-7 cells were seeded at 15 × 103 per well in a 6-well plate
and were grown with 3 mL of DMEM. Existing media were
replaced after 48 h of incubation and the cells were further
incubated with fresh medium containing the required concentrations of the metal complex for 24 h. After removal of the
drug containing media, the cells were treated using 4% paraformaldehyde solution in 1× PBS ( pH 7.2) and incubated at
4 °C for half an hour. The cells were then washed two times
with 1× PBS ( pH 7.2) and stained with DAPI (1 μg mL−1) for a
few seconds. The cells were washed several times using 1× PBS
( pH 7.2). The optical microscopy images of MCF-7 cells were
acquired using an OLYMPUS IX 81 epifluorescence inverted
microscope at 60× magnification. Both DIC and fluorescence
microscopy images were taken and processed using OLYMPUS
Cell P software. Merged images were produced for better
understanding.
5
MCF-7 cells (5 × 10 per plate) were seeded in a 90 mm dia
petri-dish in 12 mL DMEM and incubated for 48 h at 37 °C
under a 5% carbon dioxide atmosphere. After 48 h, media
were removed, followed by addition of fresh media containing
appropriate concentrations of test compounds and incubated
for another 24 h under the same conditions as above. After
24 h of incubation with test compounds, cells were harvested
by trypsinization and washed twice with cold 1× PBS ( pH 7.2).
Cells were fixed using 70% ethanol and stored at 4 °C for 12 h.
In order to stain DNA for determining the cell cycle distribution, cell pellets were transferred and resuspended in PBS
solution containing RNase (100 μg mL−1) and propidium
iodide (55 μg mL−1). The resulting solution was then incubated
for 30 min at 37 °C in the dark. Finally, homogenized cell
samples were analyzed by flow-cytometry using a fluorescenceactivated cell sorter (FACS) (FACS Calibur, Becton Dickinson,
CA). The resulting DNA histograms were quantified using the
CellQuestPro software (BD).
DNA ladder assay for apoptosis detection
DNA ladder assay was used to detect the cellular apoptosis
induced by the test compound. MCF-7 cells (5 × 105 per plate)
This journal is © The Royal Society of Chemistry 2016
Optical microscopy imaging
Ruthenium accumulation study in cancer cells
Briefly, 1 × 106 MCF-7 cells were seeded in each 90 mm dia
petri-dish and incubated for 48 h. After incubation, equimolar
concentrations (3 µM) of each complex were added and
a further 24 h of drug treatment was allowed. Following
this, the cells were washed with PBS ( pH 7.2), treated with
Dalton Trans., 2016, 45, 8541–8555 | 8551
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
trypsin-EDTA, counted and 1 × 106 number of cells were
collected as cell pellets using centrifugation. Cell pellets were
digested overnight in concentrated nitric acid (70% v/v) at
65 °C; the resulting solutions were diluted using doubledistilled water to give a final concentration of 5% HNO3 and
the amount of Ru taken up by the cells was determined by
ICP-MS (Thermo Scientific XSERIES2). The standard solutions
of ruthenium was freshly prepared and analyzed while analyses for the samples were carried out in triplicates and the
standard deviations were calculated.
Caspase-7 activation
A quantitative estimation of caspase activity was studied using
MCF-7 cells by using a colorimetric Caspase Assay kit from
Sigma. The technique principally detects the free chromophore ( p-nitroaniline, pNA), following its cleavage from the
Ac-DEVD-pNA substrate. Though the substrate Ac-DEVD-pNA is
cleaved by both caspase-3 and caspase-7, yet this experiment
would indicate the presence of caspase-7 70–74 since MCF-7
cells are already known not to have caspase-3.75 Briefly, 1 × 106
MCF-7 cells were seeded in a 90 mm dia petri-dish and incubated for 48 h followed by treatment with test compounds
with an additional incubation of 24 h. Then the cells were harvested, lysed and treated as per the kit manufacturer’s (Sigma)
standard protocol. All the samples were tested in triplicate
including the standard curve of pNA and data were recorded
after 24 h of incubation with cell lysate at 405 nm using an
ELISA plate reader.
The data is graphically represented as the absolute concentration of pNA released (nmol min−1 ml−1) in the y-axis and
concentration of the treated drugs in the x-axis. Each concentration has been performed in triplicate.
Detection of mitochondrial membrane potential
1 × 106 MCF-7 cells were seeded in each 90 mm dia petri-dish
and incubated for 48 h. Media were then replaced with fresh
media containing the desired sub IC50 concentrations (4, 6
and 8 μM) of each compound. After further incubation of 24 h,
the cells were harvested by removing the media and subsequently washed with 1× PBS solution. The entire washings
were collected in falcon tubes and centrifuged at 2500 rpm for
10 minutes. Supernatants were discarded and the pellets were
washed with 1× PBS solution. Washed cells were suspended in
the 1× PBS buffer containing 10 µg mL−1 of JC-1 and supplemented with 10% FBS followed by incubation for
30 minutes at 37 °C under dark conditions. Finally the supernatant was removed by centrifugation and the cell pellet was
further re-suspended in 1× PBS buffer. The stained cells were
analyzed using a BD Biosciences FACS Calibur flow cytometer
measuring the red and green fluorescence intensities.
Wound assay (migration)
To evaluate the effect of drugs against cancer cell migration,
the wound healing assay was used as a tool.76,77 The migration
capacities of drug treated cells were compared to the control in
MCF-7 cell lines. Briefly, 1 × 106 cells were seeded in each well
8552 | Dalton Trans., 2016, 45, 8541–8555
Dalton Transactions
of a 6 well plate and incubated at 37 °C in a humidified incubator with 5% CO2. Sufficient time was provided for the formation of a uniform monolayer in the plate. The monolayer of
the cells was then carefully scratched using a sterile microtip
to form a denuded area across the diameter of each well. Each
well was carefully washed with 1× PBS followed by replenishment of fresh medium and addition of a sub IC50 concentration (6 μM) of 1–3. The experiments were carried out in
duplicate. The images were captured on a Phase contrast
microscope (OLYMPUS IX 81 epifluorescence inverted microscope) at variable time points (0 h i.e. initial, 12 h and 24 h).
The wound area was quantified using ImageJ software.
Haemolysis assay
The haemolysis assay was performed following the standard
method.44 Briefly, blood was collected in EDTA containing
vials, followed by centrifugation at 3000 rpm for 10 min.
Erythrocytes were subsequently washed with cold PBS ( pH 7.4)
3 times by centrifugation at 3000 rpm for 10 min. The cells
were re-suspended in the same buffer to a final concentration
of 20% (v/v) and stored at 4 °C. Various concentrations of 1, 2
and 3 were prepared in DMSO and diluted in PBS such that
the final concentration of compounds remains 5, 10 and
15 μM in the reaction vial, while the final DMSO concentration
is kept at 0.2%. The reaction vial contains 2% (final concentration) cell suspension and the required concentrations of
compounds. The vials were incubated at 37 °C for 1 h in a
shaking water bath. After incubation the vials were centrifuged
at 3000 rpm for 10 min and the supernatants were collected.
The release of haemoglobin in the supernatant was assessed
spectrophotometrically at 540 nm. Positive control in which
complete haemolysis takes place was achieved using 0.2%
(final concentration) Triton X-100. Appropriate DMSO containing PBS was used as a negative control. The experiments were
run in triplicate and the percentages of haemolysis were calculated as follows:
Haemolysis percentage ð%Þ ¼ ½ðAs An Þ=ðAp An Þ 100
where, As, An, and Ap are the absorbance of sample, negative
control and positive control respectively at 540 nm.
Chick embryo angiogenesis assay (CEA)
The chick embryo angiogenesis assay (CEA) was used to investigate the anti-angiogenic activity of 1–3 using a reported protocol.76 To briefly mention the used procedure, specific
pathogen free (SPF) fertile chicken eggs were obtained from a
recognized local poultry farm which was incubated at 37 °C
under a humid atmosphere. After the fourth day of incubation,
the shells of the eggs were cautiously broken using forceps
and placed on a sterile petri-dish. Precautions were taken to
prevent puncture of any of the blood vessels while transferring.
Stock solutions of complexes (1–3) were prepared in 10%
DMSO–PBS (v/v) solution and further diluted with PBS to
achieve a complex concentration of 10 µM (final DMSO concentration 0.2%). Sterile filter paper discs (6 mm dia) were
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
soaked in a solution of the respective complexes and placed in
3 different positions (10 µM; 10 µL) over the generating blood
vessels. Sterile filter paper discs (6 mm dia) soaked with 10 µL
of 0.2% DMSO in PBS were also used in triplicate as control.
Images of blood vessels were captured using a camera attached
to a stereomicroscope (Leica M80) at initial (i.e. 0 h) and after
4 h of incubation.
Statistical analysis
All the IC50 data are expressed as mean ± standard deviation of
three independent experiments carried out in each cell line
and in each experiment each concentration was assayed in
triplicate. The statistical analyses were performed using Graph
pad prism® software 5.0 with student’s t-test. Differences with
*P < 0.05 were considered statistically significant.
Conclusions
In summary, we have shown that design of RuII-p-cymene complexes (1–3) by variation of the steric hindrance in the ligands
(L1–L3) may result in the change of cytotoxicity and mechanism of action towards various cancer cells. Faster hydrolysis
does lead to higher cytotoxicity but it appears that the pathway
of action also changes as found from the cell cycle arrest,
MMP change and caspase activation data of the complexes. All
our complexes are stable against 110 mM chloride concentration. None of the complexes lose the arene moiety up to
24 h or more. All the complexes arrest the cell cycle in the G2/
M phase. However, 1 and 2 also exhibit arrest in the S-phase
whereas 3 shows significant arrest in the sub G1 phase.
Although the lipophilicity of complexes 1 and 2 is low, yet they
exhibit higher cytotoxicity in spite of lower accumulation
inside cells. The cellular toxicity data suggests that higher
steric hindrance can prohibit deactivation by GSH and also
alter the pathway of action. Hence by proper tuning of the
steric effect it may be possible to avoid direct inhibition by cellular thiols, in RuII–arene complexes, in those forms of
cancers where GSH is overexpressed. It is encouraging to see
that none of the complexes exhibit binding to GSH until they
are incubated with at least 25 equivalent excess GSH and 3
shows no GSH adduct even up to 100 equivalents of GSH.
Complex 3 shows the maximum change in the MMP and
highest caspase-7 activation, suggesting an intrinsic pathway
of apoptosis involving the mitochondria may be active.
Whereas in the case of 1 and 2, other extrinsic pathways may
be playing a significant role in cell killing, especially in
complex 2 which has the lowest MMP change but higher activation of caspase than 1. In addition complexes 1–3 exhibit
potential anti-metastatic and anti-angiogenic activity.
Abbreviations
GSH
MMP
ppm
Glutathione
Mitochondrial membrane potential
Parts per million
This journal is © The Royal Society of Chemistry 2016
Paper
Acknowledgements
We sincerely acknowledge DST for financial support vide
project no SB/S1/IC-02/2014. We also thank IISER Kolkata for
the annual research funding and infra-structural support,
including NMR, single crystal X-ray, microscopy and FACS
facilities. K. P. and S. K. thank UGC, S. C. thanks IISER
Kolkata for providing the research fellowship. We sincerely
thank Prof. Subhransu Pan, West Bengal University of Animal
and Fishery Sciences for useful discussions on the CEA assay.
Notes and references
1 E. Meggers, Curr. Opin. Chem. Biol., 2007, 11, 287–292.
2 C. G. Hartinger and P. J. Dyson, Chem. Soc. Rev., 2009, 38,
391–401.
3 G. Suess-Fink, Dalton Trans., 2010, 39, 1673–1688.
4 M. A. Jakupec, M. Galanski, V. B. Arion, C. G. Hartinger
and B. K. Keppler, Dalton Trans., 2008, 183–194.
5 V. Brabec and O. Novakova, Drug Resist. Updates, 2006, 9,
111–122.
6 A. Levina, A. Mitra and P. A. Lay, Metallomics, 2009, 1, 458–
470.
7 E. S. Antonarakis and A. Emadi, Cancer Chemother. Pharmacol., 2010, 66, 1–9.
8 A. Bergamo, C. Gaiddon, J. H. M. Schellens, J. H. Beijnen
and G. Sava, J. Inorg. Biochem., 2012, 106, 90–99.
9 J. M. Rademaker-Lakhai, D. Van Den Bongard, D. Pluim,
J. H. Beijnen and J. H. M. Schellens, Clin. Cancer Res.,
2004, 10, 3717–3727.
10 C. G. Hartinger, S. Zorbas-Seifried, M. A. Jakupec,
B. Kynast, H. Zorbas and B. K. Keppler, J. Inorg. Biochem.,
2006, 100, 891–904.
11 M. Pongratz, P. Schluga, M. A. Jakupec, V. B. Arion,
C. G. Hartinger, G. Allmaier and B. K. Keppler, J. Anal. At.
Spectrom., 2004, 19, 46–51.
12 P. Schluga, G. Hartinger Christian, A. Egger, E. Reisner,
M. Galanski, A. Jakupec Michael and K. Keppler Bernhard,
Dalton Trans., 2006, 1796–1802.
13 M. Matczuk, M. Przadka, S. S. Aleksenko, Z. Czarnocki,
K. Pawlak, A. R. Timerbaev and M. Jarosz, Metallomics,
2014, 6, 147–153.
14 M. J. Clarke, Coord. Chem. Rev., 2003, 236, 209–233.
15 A. D. Kelman, M. J. Clarke, S. D. Edmonds and
H. J. Peresie, J. Clin. Hematol. Oncol., 1977, 7, 274–288.
16 R. Trondl, P. Heffeter, C. R. Kowol, M. A. Jakupec,
W. Berger and B. K. Keppler, Chem. Sci., 2014, 5, 2925–
2932.
17 M. J. Clarke, S. Bitler, D. Rennert, M. Buchbinder and
A. D. Kelman, J. Inorg. Biochem., 1980, 12, 79–87.
18 Y. Han, Q. Luo, X. Hao, X. Li, F. Wang, W. Hu, K. Wu,
L. Shuang and P. J. Sadler, Dalton Trans., 2011, 40, 11519–
11529.
19 J. Reedijk, Chem. Rev., 1999, 99, 2499–2510.
Dalton Trans., 2016, 45, 8541–8555 | 8553
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Paper
20 K. Mitra, S. Patil, P. Kondaiah and A. R. Chakravarty, Inorg.
Chem., 2015, 54, 253–264.
21 M. Groessl, Y. O. Tsybin, C. G. Hartinger, B. K. Keppler and
P. J. Dyson, JBIC, J. Biol. Inorg. Chem., 2010, 15, 677–
688.
22 R. Fernandez, M. Melchart, A. Habtemariam, S. Parsons
and P. J. Sadler, Chem. – Eur. J., 2004, 10, 5173–5179.
23 S. M. Smalley Keiran, R. Contractor, K. Haass Nikolas,
N. Kulp Angela, G. E. Atilla-Gokcumen, S. Williams
Douglas, H. Bregman, T. Flaherty Keith, S. Soengas Maria,
E. Meggers and M. Herlyn, Cancer Res., 2007, 67, 209–217.
24 J. Reedijk, Platinum Met. Rev., 2008, 52, 2–11.
25 W.-H. Ang, A. Casini, G. Sava and P. J. Dyson, J. Organomet.
Chem., 2011, 696, 989–998.
26 C. A. Smith, A. J. Sutherland-Smith, B. K. Keppler, F. Kratz
and E. N. Baker, JBIC, J. Biol. Inorg. Chem., 1996, 1, 424–
431.
27 S. H. van Rijt, A. J. Hebden, T. Amaresekera, R. J. Deeth,
G. J. Clarkson, S. Parsons, P. C. McGowan and P. J. Sadler,
J. Med. Chem., 2009, 52, 7753–7764.
28 T. Bugarcic, O. Novakova, A. Halamikova, L. Zerzankova,
O. Vrana, J. Kasparkova, A. Habtemariam, S. Parsons,
P. J. Sadler and V. Brabec, J. Med. Chem., 2008, 51, 5310–
5319.
29 A. Habtemariam, M. Melchart, R. Fernandez, S. Parsons,
I. D. H. Oswald, A. Parkin, F. P. A. Fabbiani, J. E. Davidson,
A. Dawson, R. E. Aird, D. I. Jodrell and P. J. Sadler, J. Med.
Chem., 2006, 49, 6858–6868.
30 F. Wang, A. Habtemariam, E. P. L. van der Geer,
R. Fernandez, M. Melchart, R. J. Deeth, R. Aird,
S. Guichard, F. P. A. Fabbiani, P. Lozano-Casal,
I. D. H. Oswald, D. I. Jodrell, S. Parsons and P. J. Sadler,
Proc. Natl. Acad. Sci. U. S. A., 2005, 102, 18269–18274.
31 H. Chen, J. A. Parkinson, S. Parsons, R. A. Coxall,
R. O. Gould and P. J. Sadler, J. Am. Chem. Soc., 2002, 124,
3064–3082.
32 F. Wang, J. Xu, A. Habtemariam, J. Bella and P. J. Sadler,
J. Am. Chem. Soc., 2005, 127, 17734–17743.
33 S. Chatterjee, S. Kundu, A. Bhattacharyya, C. G. Hartinger
and P. J. Dyson, JBIC, J. Biol. Inorg. Chem., 2008, 13, 1149–
1155.
34 F. Wang, H. Chen, J. A. Parkinson, P. d. S. Murdoch and
P. J. Sadler, Inorg. Chem., 2002, 41, 4509–4523.
35 J. Estrela, A. Ortega and E. Obrador, Crit. Rev. Clin. Lab.
Sci., 2006, 43, 143–181.
36 M. L. O’Brien and K. D. Tew, Eur. J. Cancer, Part A, 1996,
32A, 967–978.
37 N. Traverso, R. Ricciarelli, M. Nitti, B. Marengo,
A. L. Furfaro, M. A. Pronzato, U. M. Marinari and
C. Domenicotti, Oxid. Med. Cell. Longevity, 2013, 972913.
38 M. Benlloch, A. Ortega, P. Ferrer, R. Segarra, E. Obrador,
M. Asensi, J. Carretero and J. M. Estrela, J. Biol. Chem.,
2005, 280, 6950–6959.
39 Y. Lin, Y. Huang, W. Zheng, F. Wang, A. Habtemariam,
Q. Luo, X. Li, K. Wu, P. J. Sadler and S. Xiong, J. Inorg.
Biochem., 2013, 128, 77–84.
8554 | Dalton Trans., 2016, 45, 8541–8555
Dalton Transactions
40 F. Wang, J. Xu, K. Wu, S. K. Weidt, C. L. MacKay,
P. R. R. Langridge-Smith and P. J. Sadler, Dalton Trans.,
2013, 42, 3188–3195.
41 K. Purkait, S. Karmakar, S. Bhattacharyya, S. Chatterjee,
S. K. Dey and A. Mukherjee, Dalton Trans., 2015, 44, 5969–
5973.
42 C. Zhang, Y. Wang and F. Wang, Bull. Korean Chem. Soc.,
2007, 28, 1183–1186.
43 M. M. Compton, Cancer Metastasis Rev., 1992, 11, 105–
119.
44 H.-W. An, S.-L. Qiao, C.-Y. Hou, Y.-X. Lin, L.-L. Li, H.-Y. Xie,
Y. Wang, L. Wang and H. Wang, Chem. Commun., 2015, 51,
13488–13491.
45 B. Naeye, H. Deschout, M. Roding, M. Rudemo,
J. Delanghe, K. Devreese, J. Demeester, K. Braeckmans,
S. C. De Smedt and K. Raemdonck, Biomaterials, 2011, 32,
9120–9127.
46 S. Ramakrishnan, E. Suresh, A. Riyasdeen, M. A. Akbarsha
and M. Palaniandavar, Dalton Trans., 2011, 40, 3245–3256.
47 N. Busto, J. Valladolid, C. Aliende, F. A. Jalon,
B. R. Manzano, A. M. Rodriguez, J. F. Gaspar, C. Martins,
T. Biver, G. Espino, J. M. Leal and B. Garcia, Chem. – Asian
J., 2012, 7, 788–801.
48 J. Valladolid, C. Hortigueela, N. Busto, G. Espino,
A. M. Rodriguez, J. M. Leal, F. A. Jalon, B. R. Manzano,
A. Carbayo and B. Garcia, Dalton Trans., 2014, 43, 2629–
2645.
49 M. Martinez-Alonso, N. Busto, F. A. Jalon, B. R. Manzano,
J. M. Leal, A. M. Rodriguez, B. Garcia and G. Espino, Inorg.
Chem., 2014, 53, 11274–11288.
50 N. Busto, M. Martinez-Alonso, J. M. Leal, A. M. Rodriguez,
F. Dominguez, M. I. Acuna, G. Espino and B. Garcia,
Organometallics, 2015, 34, 319–327.
51 A. Russo, W. DeGraff, N. Friedman and J. B. Mitchell,
Cancer Res., 1986, 46, 2845–2848.
52 R. E. Morris, R. E. Aird, P. d. S. Murdoch, H. Chen,
J. Cummings, N. D. Hughes, S. Parsons, A. Parkin, G. Boyd,
D. I. Jodrell and P. J. Sadler, J. Med. Chem., 2001, 44, 3616–
3621.
53 B. A. Webb, M. Chimenti, M. P. Jacobson and D. L. Barber,
Nat. Rev. Cancer, 2011, 11, 671–677.
54 W. Xia, S. Spector, L. Hardy, S. Zhao, A. Saluk, L. Alemane
and N. L. Spector, Proc. Natl. Acad. Sci. U. S. A., 2000, 97,
7494–7499.
55 M. Kajstura, H. D. Halicka, J. Pryjma and Z. Darzynkiewicz,
Cytometry, Part A, 2007, 71A, 125–131.
56 I. Romero-Canelon, L. Salassa and P. J. Sadler, J. Med.
Chem., 2013, 56, 1291–1300.
57 R. Pettinari, C. Pettinari, F. Marchetti, B. W. Skelton,
A. H. White, L. Bonfili, M. Cuccioloni, M. Mozzicafreddo,
V. Cecarini, M. Angeletti, M. Nabissi and A. M. Eleuteri,
J. Med. Chem., 2014, 57, 4532–4542.
58 P. Friedl and K. Wolf, Nat. Rev. Cancer, 2003, 3, 362–374.
59 P. Nowak-Sliwinska, C. M. Clavel, E. Paunescu, M. T. te
Winkel, A. W. Griffioen and P. J. Dyson, Mol. Pharmaceutics,
2015, 12, 3089–3096.
This journal is © The Royal Society of Chemistry 2016
View Article Online
Open Access Article. Published on 07 April 2016. Downloaded on 5/2/2026 2:33:48 AM.
This article is licensed under a Creative Commons Attribution-NonCommercial 3.0 Unported Licence.
Dalton Transactions
60 A. K. Barui, V. Veeriah, S. Mukherjee, J. Manna, A. K. Patel,
S. Patra, K. Pal, S. Murali, R. K. Rana, S. Chatterjee and
C. R. Patra, Nanoscale, 2012, 4, 7861–7869.
61 D. D. Perrin and W. L. F. Armarego, Purification of Laboratory Chemicals, 3rd edn, 1988.
62 M. A. Bennett, T. N. Huang, T. W. Matheson and
A. K. Smith, Inorg. Synth., 1982, 21, 74–78.
63 L. Palatinus and G. Chapuis, J. Appl. Crystallogr., 2007, 40,
786–790.
64 G. M. Sheldrick, Acta Crystallogr., Sect. A: Found. Crystallogr., 2008, 64, 112–122.
65 O. V. Dolomanov, L. J. Bourhis, R. J. Gildea, J. A. K. Howard
and H. Puschmann, J. Appl. Crystallogr., 2009, 42, 339–
341.
66 G. M. Sheldrick, Int. Union Crystallogr., Crystallogr. Symp.,
1991, 5, 145–157.
67 J. Sangster and A. D. Pelton, J. Phys. Chem. Ref. Data, 1987,
16, 509–561.
68 J. Marmur, J. Mol. Biol., 1961, 3, 208–218.
69 M. E. Reichmann, C. A. Rice, C. A. Thomas and P. Doty,
J. Am. Chem. Soc., 1954, 76, 3047–3053.
This journal is © The Royal Society of Chemistry 2016
Paper
70 X.-H. Yang, T. L. Sladek, X. Liu, B. R. Butler, C. J. Froelich
and A. D. Thor, Cancer Res., 2001, 61, 348–354.
71 U. Janicke Reiner, Breast Cancer Res. Treat., 2009, 117, 219–
221.
72 R. U. Janicke, M. L. Sprengart, M. R. Wati and A. G. Porter,
J. Biol. Chem., 1998, 273, 9357–9360.
73 H. Kurokawa, K. Nishio, H. Fukumoto, A. Tomonari,
T. Suzuki and N. Saijo, Oncol. Rep., 1999, 6, 33–37.
74 D. Twiddy, G. M. Cohen, M. MacFarlane and K. Cain,
J. Biol. Chem., 2006, 281, 3876–3888.
75 N. A. Thornberry, T. A. Rano, E. P. Peterson, D. M. Rasper,
T. Timkey, M. Garcia-Calvo, V. M. Houtzager,
P. A. Nordstrom, S. Roy, J. P. Vaillancourt, K. T. Chapman
and D. W. Nicholson, J. Biol. Chem., 1997, 272, 17907–
17911.
76 P. Nagababu, A. K. Barui, B. Thulasiram, C. S. Devi,
S. Satyanarayana, C. R. Patra and B. Sreedhar, J. Med.
Chem., 2015, 58, 5226–5241.
77 G. V. M. Sharma, A. Ramesh, A. Singh, G. Srikanth,
V. Jayaram, D. Duscharla, J. H. Jun, R. Ummanni and
S. V. Malhotra, MedChemComm, 2014, 5, 1751–1760.
Dalton Trans., 2016, 45, 8541–8555 | 8555