Mitochondria are the primary target in the induction of apoptosis by chiral ruthenium(II) polypyridyl complexes in cancer cells

Article abstract of DOI:10.1007/s00775-013-1069-2

A series of novel chiral ruthenium(II) polypyridyl complexes (Δ-Ru1, Λ-Ru1, Δ-Ru2, Λ-Ru2, Δ-Ru3, Λ-Ru3) were synthesized and evaluated to determine their antiproliferative activities. Colocalization, inductively coupled plasma mass spectrometry, and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay studies showed that these ruthenium(II) complexes accumulated preferentially in the mitochondria and exhibited cytotoxicity against various cancer cells in vitro. The complex Δ-Ru1 is of particular interest because it was found to have half-maximal inhibitory concentrations comparable to those of cisplatin and better activity than cisplatin against a cisplatin-resistant cell line, A549-CP/R. Δ-Ru1 induced alterations in the mitochondrial membrane potential and triggered intrinsic mitochondria-mediated apoptosis in HeLa cells, which involved the regulation of Bcl-2 family members and the activation of caspases. Taken together, these data suggest that Δ-Ru1 may be a novel mitochondria-targeting anticancer agent. SBIC 2013.

Full text of DOI:10.1007/s00775-013-1069-2

J Biol Inorg Chem (2014) 19:335–348
DOI 10.1007/s00775-013-1069-2
ORIGINAL PAPER
Mitochondria are the primary target in the induction of apoptosis
by chiral ruthenium(II) polypyridyl complexes in cancer cells
•
Jin-Quan Wang Ping-Yu Zhang Chen Qian
•
•
•
Xiao-Juan Hou Liang-Nian Ji Hui Chao
•
Received: 11 September 2013 / Accepted: 12 November 2013 / Published online: 28 November 2013
Ó SBIC 2013
Abstract A series of novel chiral ruthenium(II) poly-
pyridyl complexes (D-Ru1, K-Ru1, D-Ru2, K-Ru2,
D-Ru3, K-Ru3) were synthesized and evaluated to deter-
mine their antiproliferative activities. Colocalization,
inductively coupled plasma mass spectrometry, and
3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bro-
mide assay studies showed that these ruthenium(II) com-
plexes accumulated preferentially in the mitochondria and
exhibited cytotoxicity against various cancer cells in vitro.
The complex D-Ru1 is of particular interest because it was
found to have half-maximal inhibitory concentrations
comparable to those of cisplatin and better activity than
cisplatin against a cisplatin-resistant cell line, A549-CP/R.
D-Ru1 induced alterations in the mitochondrial membrane
potential and triggered intrinsic mitochondria-mediated
apoptosis in HeLa cells, which involved the regulation of
Bcl-2 family members and the activation of caspases.
Taken together, these data suggest that D-Ru1 may be a
novel mitochondria-targeting anticancer agent.
Keywords Ruthenium(II) complexes ꢀ
Mitochondria targeting ꢀ Cytotoxicity ꢀ Apoptosis
Introduction
Although cisplatin is active against a variety of cancers, its
use is often associated with severe dose-limiting side
effects, including nephrotoxicity, neurotoxicity, and gas-
trointestinal toxicity [1–4]. In addition, other factors, such
as low water solubility and drug resistance, have limited
the clinical applications of cisplatin [5, 6]. These limita-
tions have led to an urgent search for other metal-based
agents that can replace platinum-based drugs. In recent
years, ruthenium-based complexes have emerged as
promising antitumor and antimetastatic agents with equal
or even greater antitumor activity and lower toxicity [7, 8].
To date, two ruthenium-based complexes, NAMI-A and
KP1019, have entered human clinical trials [9, 10]. These
two ruthenium(III) complexes exhibited great antitumor
activity in preclinical studies. NAMI-A had inhibitory
effects against the formation of cancer metastases in a
variety of animal models but lacked direct cytotoxic effects
on the primary tumor [11]. In contrast, KP1019 exhibited
direct antitumor effects against a series of primary human
tumor explants [12]. In addition, several mechanisms have
been proposed to explain the anticancer effects of these
ruthenium complexes, including interaction with DNA
[13–16], modification of the cell membrane and cell
adhesion properties [17], blocking of the cell cycle [18],
and inhibition of various kinases [19] and topoisomerase
[20–22].
Electronic supplementary material The online version of this
article (doi:10.1007/s00775-013-1069-2) contains supplementary
material, which is available to authorized users.
J.-Q. Wang ꢀ P.-Y. Zhang ꢀ C. Qian ꢀ L.-N. Ji ꢀ H. Chao (&)
MOE Key Laboratory of Bioinorganic and Synthetic Chemistry,
State Key Laboratory of Optoelectronic
Materials and Technologies,
School of Chemistry and Chemical Engineering,
Sun Yat-Sen University, Guangzhou 510275,
People’s Republic of China
e-mail: ceschh@mail.sysu.edu.cn
J.-Q. Wang
Guangdong Pharmaceutical University,
Guangzhou 510006, People’s Republic of China
X.-J. Hou
Huaihua Medical College,
Huaihua 418000, People’s Republic of China
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Recently,
a
novel chemotherapeutic mechanism
It has been reported that some ruthenium(II) complexes,
especially those bearing 2-phenylimidazo[4,5-f][1,10]phe-
nanthroline (PIP) ligands, can induce mitochondria -medi-
ated apoptosis in cancer cells [27–36]. Motivated by these
findings, we synthesized six chiral ruthenium polypyridyl
complexes (D-Ru1, K-Ru1, D-Ru2, K-Ru2, D-Ru3, and K-
Ru3, as shown in Fig. 1). To explore the structure–activity
relationship of ruthenium(II) complexes, we functionalized
PIP by replacing one benzene ring with another heterocycle
with a hydroxyl group, in which an ortho hydroxyl group that
may form an intramolecular hydrogen bond with the nitro-
gen atom of the imidazole ring was introduced into the
ligands to extend the planarity. Inductively coupled plasma
mass spectrometry (ICP-MS) showed that these six com-
plexes accumulated primarily in the mitochondria. When
comparing these six complexes, we found that the cytotox-
icity was correlated with the cellular uptake properties, and
D-Ru1 exhibited the highest cytotoxicity. Further investi-
gations, including live-cell imaging, transmission electron
microscopy, flow cytometry, and Western blot analysis,
demonstrated that the mitochondria are the primary target of
D-Ru1 in the induction of apoptosis in cancer cells.
involving the disruption of mitochondria has been inves-
tigated because this mechanism circumvents upstream
apoptotic pathways that may be mutated or lacking in
cancer cells [23]. Cancer cell mitochondria are structurally
and functionally different from their normal counterparts.
Accordingly, tumor cells often use glycolysis for ATP
generation, which is normally performed primarily by
oxidative phosphorylation [24]. Moreover, cancer cells
have higher mitochondrial membrane potentials, rendering
them more susceptible to mitochondrial perturbations than
nonimmortalized cells. On the basis of these factors, doz-
ens of mitochondria-targeting agents have emerged as a
means to disrupt the mitochondrial membrane potential and
further permeabilize the mitochondrial outer membrane,
leading to the release of proapoptotic proteins and the
impairment of the bioenergetic functions of mitochondria
[25]. The activation of the cell death machinery in cancer
cells by inhibiting tumor-specific alterations to mitochon-
drial metabolism or by stimulating mitochondrial mem-
brane permeabilization could therefore be a promising
strategy in cancer therapy [26].
Fig. 1 Chemical structures of
the ruthenium(II) complexes
studied
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Materials and methods
Materials
337
(s, 1H), 6.61 (s, 1H), 6.06 (s, 2H). FAB-MS: m/z 357
(M ? 1).
2-(8-Hydroxyjulodinyl)imidazo[4,5-f][1,10]phenanthroline
All reagents and solvents were of analytical grade unless
otherwise noted. Cisplatin, 40,6-diamino-2-phenylindole
(DAPI), 20,70-dichlorofluorescein diacetate (DCFH-DA),
and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium
bromide (MTT) were obtained from Sigma-Aldrich. Stock
solutions of cisplatin (10 mM) and ruthenium(II) com-
plexes (1 mM) were prepared in dimethyl sulfoxide
(DMSO), and were stable for at least 48 h at room tem-
perature, as monitored by UV–visible spectroscopy. 1,10-
Phenanthroline-5,6-dione [37], D-[Ru(bpy)2(py)2][O,O0-
dibenzoyl-D-tartrate]ꢀ12H2O [38] (where bpy is 2,20-
bipyridyl and py is pyridyl), and K-[Ru(bpy)2(py)2] [O,O0-
dibenzoyl-D-tartrate]ꢀ12H2O [38] were prepared according
to methods described in the literature.
A
mixture of 1,10-phenanthroline-5,6-dione (0.21 g,
1 mmol), 9-formyl-8-hydroxyjulodine (0.217 g, 1 mmol),
ammonium acetate (1.94 g, 25.0 mmol), and glacial acetic
acid (15 mL) was refluxed with stirring for 4 h. The yel-
low-brown precipitate was removed by filtration while hot,
washed with H2O and diethyl ether, and then dried in
vacuo. The resulting crude solid was purified by chroma-
tography over 100-mesh SiO2 using ethanol as the eluent
and then dried in vacuo to afford 2-(8-hydroxyjulodi-
nyl)imidazo[4,5-f][1,10]phenanthroline (HQIP). Yield:
0.35 g (85.9 %). Anal. Calcd for C25H21N5O (%): C,
73.69; H, 5.19; N, 17.19. Found (%): C, 73.66; H, 5.21; N,
1
17.21. H NMR (400 MHz, DMSO-d6): d (ppm) 8.90 (dd,
J = 7.8 Hz, 2H), 8.24 (dd, J = 7.5 Hz, 2H), 7.48 (m, 3H),
3.37 (t, J = 7.3 Hz, 4H), 2.97 (t, J = 7.7 Hz, 2H), 2.79 (t,
J = 7.7 Hz, 2H), 1.96 (m, 4H). FAB-MS: m/z 408
(M ? 1).
Syntheses
2-(2-Hydroxyphenyl)imidazo[4,5-f][1,10]phenanthroline
D-Ru1
2-(2-Hydroxyphenyl)imidazo[4,5-f][1,10]phenanthroline
(HPIP) was prepared according to methods described in the
literature [15]. Anal. Calcd for C19H12N4O (%): C, 73.07;
H, 3.87; N, 17.94. Found (%): C, 73.05; H, 3.90; N, 17.91.
1H NMR (400 MHz, DMSO-d6): d (ppm) 8.84 (d,
J = 7.8 Hz, 2H), 8.24 (d, J = 7.6 Hz, 2H), 7.74 (dd,
J = 6.9 Hz, 1H), 7.49 (t, J = 6.9 Hz, 2H) 7.42 (t,
J = 6.9 Hz, 1H), 7.26 (t, J = 6.9 Hz, 1H), 7.20 (d,
J = 6.9 Hz, 1H). Fast atom bombardment mass spec-
trometry (FAB-MS): m/z 313 (M ? 1).
D-[Ru(bpy)2(HPIP)](ClO4)2 (D-Ru1) was prepared accord-
ingtomethodsdescribedintheliterature[15]. Anal. Calcdfor
C39H28N8ORu (%): C, 64.54; H, 3.89; N, 15.44. Found (%):
C, 64.51; H, 3.92; N, 15.43. 1H NMR (400 MHz, DMSO-d6):
d (ppm) 9.14 (d, J = 8.2 Hz, 2H), 8.86 (d, J = 8.2 Hz, 2H),
8.86 (d, J = 8.2 Hz, 2H), 8.20 (t, J = 8.4 Hz, 2H), 8.02–8.13
(m, 4H), 7.88–7.93 (m, 2H), 7.83 (d, J = 5.2 Hz, 2H), 7.58
(dd, J = 7.4 Hz, 4H), 7.42 (t, J = 8.2 Hz, 2H), 7.32 (t,
J = 8.0 Hz, 2H), 7.09 (t, J = 7.8 Hz, 2H). Electro-
spray mass spectrometry (ES-MS) (CH3CN): m/z 726
([M - 2ClO4 ? H]?), 363 ([M - 2ClO4]2?). Circular
dichroism (CD) (H2O): k/nm (De/M-1 cm-1) 472 (-10.7),
421 (9.7), 291 (-116.8), 274 (34.6), 240 (15.9).
2-(6-Hydroxy-3,4-methylenedioxyphenyl)imidazo[4,5-
f][1,10]-phenanthroline
A
mixture of 1,10-phenanthroline-5,6-dione (0.21 g,
1 mmol), 6-hydroxy-3,4-(methylenedioxy)benzaldehyde
(0.166 g, 1 mmol), ammonium acetate (1.94 g,
25.0 mmol), and glacial acetic acid (15 mL) was refluxed
with stirring for 4 h. The yellow-brown precipitate was
removed by filtration while hot, washed with H2O and
diethyl ether, and then dried in vacuo. The resulting crude
solid was purified by chromatography over 100-mesh SiO2
using ethanol as the eluent and then dried in vacuo to afford
2-(6-hydroxy-3,4-methylenedioxyphenyl)imidazo[4,5-f]
[1,10]-phenanthroline (HMIP). Yield: 0.30 g (84.2 %).
Anal. Calcd for C20H12N4O3 (%): C, 67.42; H, 3.39; N,
K-Ru1
K-[Ru(bpy)2(HPIP)](ClO4)2 (K-Ru1) was prepared accord-
ing to methods described in the literature [15]. The elemental
analysis, 1H NMR, andmassspectral data forK-Ru1 werethe
same as those for D-Ru1, within experimental error. CD
(H2O): k/nm (De/M-1 cm-1) 472 (10.8), 421 (-9.8), 291
(117.0), 274 (-35.1), 240 (-15.8).
D-Ru2
1
15.72. Found (%): C, 67.45; H, 3.36; N, 15.74. H NMR
A
mixture of D-[Ru(bpy)2(py)2][O,O0-dibenzoyl-D-tar-
(400 MHz, DMSO-d6): d (ppm) 8.87 (d, J = 7.8 Hz, 2H),
8.14 (d, J = 7.5 Hz, 2H), 7.39 (dd, J = 8.0 Hz, 2H), 7.12
trate]ꢀ12H2O (0.2 g, 0.2 mmol), HMIP (0.071 g, 0.2 mmol),
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and ethylene glycol (10 mL) was refluxed under argon for
8 h, and the color of the solution changed from purple to
red. The solution was cooled to room temperature, and
20 mL of H2O was added. After filtration, a dark-red pre-
cipitate was obtained by the dropwise addition of an
aqueous NaClO4 solution. The crude product was purified
by column chromatography using neutral alumina with
CH3CN–toluene (4:1, v/v) as the eluent. The red-brown
band was collected. The solvent was removed
under reduced pressure, and D-[Ru(bpy)2(HMIP)](ClO4)2
(D-Ru2) was obtained as a red-brown powder. Yield:
0.17 g (87.6 %). Anal. Calcd for C40H28N8O3Ru (%): C,
62.41; H, 3.67; N, 14.56. Found (%): C, 62.43; H, 3.69; N,
J = 6.8 Hz, 4H), 7.83 (m, 3H), 7.61 (t, J = 7.6 Hz, 2H),
7.37 (t, J = 7.8 Hz, 2H), 3.02 (t, J = 7.6 Hz, 4H), 2.27 (t,
J = 7.8 Hz, 2H), 1.75 (t, J = 7.4 Hz, 2H), 1.27 (m, 4H).
ES-MS (CH3CN): m/z 821 ([M - 2ClO4 ? H]?), 411
([M - 2ClO4]2?). CD (H2O): k/nm (De/M-1 cm-1) 472
(-13.3), 420 (15.9), 292 (-196.8), 276 (73.3), 239 (17.6).
K-Ru3
K-[Ru(bpy)2(HQIP)](ClO4)2 (K-Ru3) was synthesized in a
manner identical to that described for D-[Ru(bpy)2(H-
QIP)](ClO4)2, but using K-[Ru(bpy)2(py)2][O,O0-diben-
zoyl-L-tartrate]ꢀ12H2O (0.20 g, 0.2 mmol) rather than
D-[Ru(bpy)2(py)2][O,O0-dibenzoyl-D-tartrate]ꢀ12H2O. Yield:
0.18 g (87.8 %). The elemental analysis, 1H NMR, and mass
spectral data for K-Ru3 were the same as those for D-Ru3,
within experimental error. CD (H2O): k/nm (De/M-1 cm-1
472 (12.9), 420 (-14.7), 292 (190.0), 276 (-70.5), 239
(-16.9).
1
14.55. H NMR (400 MHz, DMSO-d6): d (ppm) 9.08 (d,
J = 8.0 Hz, 2H), 8.84 (dd, J = 8.2 Hz, 4H), 8.20 (t,
J = 7.9 Hz, 2H), 8.08 (dd, J = 7.9 Hz, 4H), 7.91 (d,
J = 7.8 Hz, 2H), 7.82 (d, J = 8.3 Hz, 2H), 7.69 (s, 1H), 7.57
(m, 4H), 7.32 (t, J = 7.8 Hz, 2H), 6.74 (s, 1H), 6.09 (s, 2H).
ES-MS (CH3CN): m/z 770 ([M - 2ClO4 ? H]?), 384
([M - 2ClO4]2?). CD (H2O): k/nm (De/M-1 cm-1) 471
(-13.6), 420 (14.3), 291 (-169.8), 274 (59.4), 240 (18.8).
Physical measurements
K-Ru2
Microanalysis (C, H, and N) was performed using a Vario
1
EL elemental analyzer. H NMR spectra were recorded at
K-[Ru(bpy)2(HMIP)](ClO4)2 (K-Ru2) was synthesized in a
manner identical to that described for D-Ru2, but using K-
[Ru(bpy)2(py)2][O,O0-dibenzoyl-L-tartrate]ꢀ12H2O (0.20 g,
0.2 mmol) rather than D-[Ru(bpy)2(py)2][O,O0-dibenzoyl-
D-tartrate]ꢀ12H2O. Yield: 0.16 g (85.6 %). The elemental
analysis, 1H NMR, and mass spectral data for K-Ru2 were
the same as those for D-Ru2, within experimental error.
CD (H2O): [k/nm (De/M-1 cm-1) 471 (13.9), 420 (-14.5),
291 (171.0), 274 (-59.9), 240 (-19.0).
room temperature with a Bruker AVANCE AV 400-NMR
spectrometer using DMSO-d6 as the solvent and tetra-
methylsilane as the internal standard. Fast atom bombard-
ment mass spectra were acquired using a VG ZAB-HS
spectrometer in a 3-nitrobenzyl alcohol matrix. Electro-
spray mass spectra were recorded with an LCQ system
(Finnigan MAT, USA). The spray voltage, tube lens offset
voltage, capillary voltage, and capillary temperature were
set to 4.50 kV, 30.00 V, 23.00 V, and 200 °C, respec-
tively, and the quoted m/z values are for the major peaks in
the isotope distribution. The CD spectra were measured
with a JASCO J810 spectrometer.
D-Ru3
A
mixture of D-[Ru(bpy)2(py)2][O,O0-dibenzoyl-D-tar-
trate]ꢀ12H2O (0.2 g, 0.2 mmol), HQIP (0.081 g,
0.2 mmol), and ethylene glycol (10 mL) was refluxed
under argon for 8 h, and the color of the solution changed
from purple to red. The solution was cooled to room
temperature, and 20 mL of H2O was added. After filtration,
a dark-red precipitate was obtained by the dropwise addi-
tion of an aqueous NaClO4 solution. The crude product was
purified by column chromatography using neutral alumina
with CH3CN–toluene (4:1, v/v) as the eluent. The red-
brown band was collected. The solvent was removed
under reduced pressure, and D-[Ru(bpy)2(HQIP)](ClO4)2
(D-Ru3) was obtained as a red-brown powder. Yield:
0.17 g (83.8 %). Anal. Calcd for C45H37N9ORu (%): C,
65.84; H, 4.54; N, 15.36. Found (%): C, 65.83; H, 4.58; N,
Cell lines and cell culture
Human cancer cell lines, including the HepG2 hepatocel-
lular carcinoma cell line, the HeLa cervical carcinoma cell
line, and the MCF-7 breast carcinoma cell line, were pur-
chased from American Type Culture Collection (Manassas,
VA, USA). The A549 human lung carcinoma cell line, the
cisplatin-resistant cell line A549-CP/R, the MG63 osteo-
sarcoma cell line, the BEL-7402 hepatocellular carcinoma
cell line, and LO2 normal hepatic cells were obtained from
the cell bank of the Cell Institute of Sinica Academia
Shanghai (Shanghai, China). All cell lines were cultured in
25-cm2 culture flasks in either RPMI-1640 medium
(Invitrogen, Carlsbad, CA, USA) or Dulbecco’s modified
Eagle’s medium (Gibco, Gaithersburg, MD, USA) sup-
plemented with fetal bovine serum (10 %), penicillin
1
15.35. H NMR (400 MHz, DMSO-d6): d (ppm) 10.06 (d,
J = 8.2 Hz, 2H), 8.89 (m, 6H), 8.27 (m, 4H), 8.10 (t,
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339
(100 U/mL), and streptomycin (50 U/mL) at 37 °C in a
CO2 incubator (95 % relative humidity, 5 % CO2). The
cells were cultured until they reached the logarithmic
growth phase unless otherwise specified.
the samples were observed under an inverted fluorescence
microscope (Zeiss Axio Observer D1) [41].
To evaluate the intracellular localization of the ruthe-
nium(II) complexes, we examined the extent of color
merging using the endoplasmic-reticulum-specific probe
ER-Tracker Green and the mitochondria-specific probe
Mito-Tracker Green FM [42, 43]. HeLa cells were treated
as described earlier, and then, Mito-Tracker Green FM or
ER-Tracker Green was added to the dishes. After incuba-
tion for 15 min, the dishes were washed with 1 mL of PBS
and the samples were observed under an inverted fluores-
cence microscope (Zeiss Axio Observer D1).
Cytotoxicity assay
The standard MTT assay was used to assess the cytotox-
icity as previously described [39]. Cells were adjusted to
the proper concentration, seeded in 96-well microplates,
and incubated overnight at 37 °C in a 5 % CO2 incubator.
After serial dilutions of the ruthenium(II) complexes had
been added, the cells were incubated for an additional 48 h.
The negative control cells were treated with the vehicle,
and cisplatin was used as a positive control agent. Wells
containing culture medium without cells were used as
blanks. On completion of the incubation, MTT solution
[MTT working solution, 5 mg/mL phosphate-buffered
saline (PBS), 20 lL per well] was added, and the cells
were incubated for 4 h. Then, the medium was aspirated
and replaced with 150 lL of 2-propanol per well to dis-
solve the formazan. The optical density of each well was
measured using a microplate reader at a wavelength of
570 nm. The half-maximal inhibitory concentration (IC50)
was determined by a plot of the viability versus the dose
used to treat the cells. All data were from at least three
independent experiments and are expressed as the
mean the standard deviation.
Annexin V staining to detect early-phase apoptosis
Early-phase apoptosis was detected by annexin V staining.
HeLa cells were adjusted to the proper concentration,
seeded in six-well culture plates, and incubated overnight
at 37 °C in a 5 % CO2 incubator. Then, the ruthenium(II)
complex (15 lM) was added, and the cells were incubated
for 0–36 h. The cells were trypsinized, washed twice with
cold PBS, and then resuspended in 100 lL of binding
buffer [50 mmol/L N-(2-hydroxyethyl)piperazine-N0-eth-
anesulfonic acid/NaOH, pH 7.4, 700 mmol/L NaCl,
12.5 mmol/L CaCl2] containing 5 lL of annexin V stock
solution (Invitrogen, Paisley, UK). After the cells had been
incubated for 15 min at room temperature in a light-pro-
tected area, 400 lL of binding buffer was added, and the
specimens were quantified by flow cytometry (FACSCan-
to II, BD Biosciences, San Jose, CA, USA).
Inductively coupled plasma mass spectrometry
ICP-MS (Thermo Elemental system) was used to quantify
the levels of ruthenium in different subcellular compart-
ments, as described in our previous report [40]. HeLa cells
were cultured in 25-cm2 culture plates (Corning) and
incubated with the six ruthenium(II) complexes at the same
concentration of 20 lM for 48 h. The cytoplasmic, nuclear,
and mitochondrial fractions were obtained. Aliquots were
removed and used to determine the protein concentration
using bicinchoninic acid (Pierce, Rockford, IL, USA). The
results are reported as nanograms of ruthenium per milli-
gram of cellular protein. The data are reported as the
mean the standard deviation (n = 3).
DAPI staining to detect late-phase apoptosis
HeLa cells were prepared as described earlier. At different
time points, cells were stained with DAPI to analyze
nuclear fragmentation. The cells were fixed with 4 %
formaldehyde in PBS for 15 min, washed twice with PBS,
and then permeabilized with 0.2 % Triton X-100 in PBS
for 5 min. Next, the cells were washed with cold PBS
before incubation with DAPI (5 lg/mL) for 10 min at
room temperature, and then the DAPI-stained cells were
washed twice with PBS. The samples were observed under
an inverted fluorescence microscope [44].
Cellular localization analysis
Measurement of intracellular reactive oxygen species
HeLa cells were adjusted to the proper concentration and
seeded on 35-mm glass-bottomed culture dishes. When the
cells reached more than 60 % confluency, the ruthe-
nium(II) complexes were added at a concentration of
10 lM, and the cells were incubated for 48 h. Then, DAPI
(5 lg/mL) was added to the dishes, which were incubated
for 5 min. The cells were washed with 1 mL of PBS before
Accumulation of reactive oxygen species (ROS) was
quantified using the DCFH-DA assay. DCFH-DA is a
nonfluorescent compound that becomes the highly fluo-
rescent 20,70-dichlorofluorescein on oxidation by intracel-
lular ROS. HeLa cells were treated with ruthenium(II)
complexes or H2O2 at different concentrations for 6 h, and
untreated cells were used as a control. Then, the cells were
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incubated with 10 lM DCFH-DA (Sigma-Aldrich) for
30 min at 37 °C. The intracellular fluorescence intensity of
the cells was measured using a flow cytometer (FACSC-
anto II, BD Biosciences, San Jose, CA, USA) and a
were taken and scanned using the program Eversmart Jazz
(Scitex).
Western blot analysis
¨
microplate analyzer (Infinite M200, Tecan, Mannedorf,
Switzerland). When necessary, N-acetylcysteine (NAC;
10 mM) was applied 1 h before the treatment with the
ruthenium(II) complexes or H2O2 and was kept in the
medium during treatment until the cells were analyzed. The
excitation wavelength was 488 nm, and the fluorescence
was measured at 530 nm. The acquisition and analysis of
the flow cytometry data were performed using FlowJo.
Exponentially growing HeLa cells were treated with the
ruthenium(II) complexes (15 lM) for 6–48 h (1–24 h for
detection of cytochrome c). For the preparation of whole-
cell or cytoplasmic protein extracts [46], at each time point,
cells were washed twice with ice-cold PBS and lysed
in radioimmunoprecipitation assay buffer {50 mM
tris(hydroxymethyl)aminomethane–HCl, 150 mM NaCl,
0.1 % sodium dodecyl sulfate, 1 % NP-40, 0.5 % sodium
deoxycholate, 1 mM phenylmethanesulfonyl fluoride,
100 mM leupeptin, and 2 mg/mL aprotinin, pH 8.0] to
extract the total cellular proteins. The protein extracts were
resolved using 10 % sodium dodecyl sulfate–polyacryl-
amide gels loaded with equal amounts of the proteins per
lane. Equal amounts of protein (20–50 lg) were electro-
phoresed on 10 % or 15 % sodium dodecyl sulfate–poly-
acrylamide gels. Next, the proteins were electroblotted
onto Immobilon-P poly(vinylidene difluoride) transfer
membranes (Millipore, Bedford, MA, USA), which were
then blocked with 5 % nonfat milk in a buffer consisting of
50 mM tris(hydroxymethyl)aminomethane, pH 7.6,
150 mM NaCl, and 0.1 % Tween 20 on a shaker at room
temperature for 1 h. Then, the membranes were probed
with primary antibodies (Epitomics, Burlingame, CA,
USA) at 1:1,000 dilution in 5 % nonfat milk overnight at
4 °C and then with secondary antibodies (goat anti-rat,
Santa Cruz Biotechnology, Santa Cruz, CA, USA) conju-
gated with horseradish peroxidase at 1:2,000 dilution for
1 h at room temperature. To ensure the presence of com-
parable amounts of proteins in each lane, the membranes
were stripped and probed for glyceraldehyde 3-phosphate
dehydrogenase (Santa Cruz Biotechnology, Santa Cruz,
CA, USA). The protein bands were visualized using elec-
trochemiluminescence detection reagents according to the
instructions for the SuperSignal West Pico kit (Pierce
Biotechnology, Rockford, IL, USA).
Evaluation of the mitochondrial membrane potential
Mitochondrial dysfunction was detected by measuring
changes in the mitochondrial membrane potential using an
inverted fluorescence microscope and flow cytometry after
live cells had been stained with 5,50,6,60-tetrachloro-
1,10,3,30-tetraethylbenzimidazolylcarbocyanine iodide (JC-1)
[45]. For microscopic observation, HeLa cells were adjusted
to the proper concentration and seeded in six-well plates.
When the cells reached more than 60 % confluency, the
ruthenium(II) complexes were added at a concentration of
15 lM, and the cells were incubated for 0–12 h. Then, the
cells were incubated for 30 min in complete medium con-
taining5 mg/mL JC-1. Aftertheyhadbeenwashedtwicewith
PBS, the stained cells were analyzed using an inverted fluo-
rescence microscope.
For flow cytometry analysis, HeLa cells were prepared
as described earlier and incubated with the ruthenium(II)
complexes at concentrations of 5, 10, and 20 lM for 6 h.
After exposure to the complexes, the cells were trypsinized
and resuspended in 0.5 mL of PBS containing 5 mg/mL
JC-1. After the cells had been incubated for 30 min in the
dark, the supernatant was immediately removed by cen-
trifugation. Next, the cells were washed and resuspended in
PBS, and the stained cells were immediately analyzed by
flow cytometry. The relative intensities of the green fluo-
rescence signals from JC-1 monomers were used to show
the cells that lost mitochondrial membrane potential.
Transmission electron microscopy
Results and discussion
HeLa cells (5 9 105) were treated with the ruthenium(II)
complexes (15 lM) for 24 h. The cells were washed twice
and fixed with 2 % glutaraldehyde at 4 °C for 1 h and
postfixed with 2 % osmium tetroxide. The cells were then
dehydrated with sequential washes in ethanol and embed-
ded in Spurr resin. The ultrathin sections obtained were
mounted on copper grids, counterstained with uranyl ace-
tate and lead citrate, and then visualized using an electron
microscope (JEM 100 CX, JEOL, Tokyo, Japan). Images
Synthesis and characterization
The synthesis of the ruthenium(II) complexes (D-Ru1,
K-Ru1, D-Ru2, K-Ru2, D-Ru3, and K-Ru3) is illustrated
in Scheme S1. These complexes were characterized by H
1
NMR spectroscopy, ES-MS, CD spectroscopy, and ele-
mental analysis. [M - 2ClO4 - H]? and [M - 2ClO4]2?
signals were observed in the ES-MS spectra of the ruthe-
nium(II) complexes. The enantiomers yielded identical,
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341
1
well-defined H NMR spectra, which permitted unambig-
study had showed the cellular uptake of the ruthenium(II)
complex was important for the cytotoxicity in vitro [22,
40]. So we used live-cell imaging and ICP-MS to deter-
mine the distribution of the ruthenium(II) complexes in the
HeLa cells to address this question. Live-cell imaging
showed that most of these complexes were distributed in the
cytoplasmwhencellswereincubatedwith thesecomplexes at
a concentration half the IC50 value for 48 h (Fig. S2). Fur-
thermore, ICP-MS showed that after exposed to the same
concentration of these six ruthenium (II) complexes in the
same time, the quantitative order of ruthenium in the whole-
cell lysates was as follows: D-Ru1 & K-Ru1 [
D-Ru2 & K-Ru2 [ D-Ru3 & K-Ru3 (Fig. 2). These data
suggested that there was a positive relationship between the
cellular uptakeandtheinvitrocytotoxicityfortheHeLa cells.
The highest level of ruthenium in the whole-cell lysates
(125 11 ng of ruthenium per milligram of protein) was
found for D-Ru1, and this complex was the most active
anticancer agent (IC50 = 9.3 2.1 lM for HeLa). K-Ru1
displayed slightly lower toxicity than D-Ru1, which also
seems to be attributed to the slightly less cellular uptake of
ruthenium; in HeLa cells, the same relationship was found for
D-Ru2 and K-Ru2, but the opposite relationship was found
for D-Ru3 and K-Ru3.
uous identification and the assessment of purity. The
enantiomeric purity of these complexes was assayed by CD
spectroscopy. D-Ru1, D-Ru2, and D-Ru3 exhibited two
primary CD signals, with a positive peak at approximately
275 nm and a negative peak at approximately 291 nm. As
expected, the opposite patterns were observed for the
enantiomers K-Ru1, K-Ru2, and K-Ru3 (Fig. S1).
Cytotoxicity study
We examined the in vitro cytotoxic activity of these
ruthenium(II) complexes against six selected human cancer
cell lines (HeLa, A549, HepG2, MCF-7, BEL-7402, and
MG-63), one cisplatin-resistant cell line (A549-CP/R), and
a normal human liver cell line (LO2) using the MTT assay.
Cisplatin was used as a positive control. As shown in
Table 1, D-Ru1 exhibited cytotoxicity similar to that of
cisplatin against the six cancer cell lines. Notably, this
complex could inhibit the growth of the cisplatin-resistant
cell line A549-CP/R, suggesting that the anticancer
mechanism of D-Ru1 differs from that of cisplatin and that
this complex could help overcome the challenges associ-
ated with the emergence of cisplatin resistance in cancer
therapy. In addition, D-Ru1 was found to have potent
toxicity against tumor cells and was much less toxic than
cisplatin to the normal human liver cell line (LO2), indi-
cating that D-Ru1 most likely has a better therapeutic
profile than cisplatin. Because HeLa cells exhibited the
highest sensitivity to D-Ru1, we used this cell line as the
cell model for further investigation.
Most of D-Ru1 and K-Ru1 accumulated in the mito-
chondrial fraction of HeLa cells and they could hardly be
detected in the nucleus. In contrast, D-Ru2, K-Ru2,
D-Ru3, and K-Ru3 were found in the mitochondria,
nucleus, and cytoplasm. This significant difference in
subcellular distribution is probably another factor that is
responsible for the disparity in cytotoxicity among these
ruthenium(II) complexes.
Cellular uptake and subcellular distribution
Cellular localization analysis
The six ruthenium(II) complexes, with similar ligands,
displayed very different cytotoxicity in vitro. Our previous
We used fluorescence microscopy to obtain further informa-
tion on the exact cellular localization of D-Ru1 and K-Ru1,
Table 1 Half-maximal inhibitory concentrations (IC50) of the ruthenium(II) complexes and cisplatin toward different tumor cell lines
Cell line
IC50 (lM)
D-Ru1
K-Ru1
D-Ru2
K-Ru2
D-Ru3
K-Ru3
Cisplatin
HeLa (ovary)
9.3 2.1
13.6 2.2
17.6 3.2
12.7 1.4
10.6 1.3
16.2 3.2
26.9 4.5
88.9 5.6
17.1 3.5
21.3 4.5
22.9 3.6
18.7 3.5
22.1 7.8
34.8 6.3
30.3 5.6
90.3 5.6
20.7 5.6
151.8 10.3
153.7 12.6
47.9 5.6
34.6 3.5
179.2 12.5
161.3 13.5
36.0 3.8
69.5 7.8
76.5 8.9
61.9 5.6
57.9 5.3
130.8 9.1
105.9 9.6
251.7 26.2
87.1 4.2
86.5 5.8
76.5 8.8
10.5 1.8
14.6 2.4
19.7 3.2
7.5 1.6
9.5 1.2
18.2 3.5
60.5 5.7
14.5 5.7
HepG2 (liver)
BEL-7402 (liver)
MG-63 (osteosarcoma)
MCF-7 (breast)
A549 (lung)
156.2 14.5
126.1 12.8
255.6 22.4
112.2 14.5
156.5 12.4
162 13.4
242.2 21.3
182.4 14.7
246.2 12.5
212 12.5
227.6 21.4
170.6 18.7
220.4 11.3
190.4 11.3
A549-CP/R (lung)
LO2 (liver)
See Fig. 1 for the structures of the ruthenium(II) complexes. The data are presented as the mean the standard deviation, and cell viability was
assessed after 48 h of incubation. A549-CP/R is a cisplatin-resistant cell line and LO2 is a human normal liver cell line
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D-Ru2 and K-Ru2, and D-Ru3 and K-Ru3. Live-cell imag-
ing showed that D-Ru1 and K-Ru1 were limited to the
cytoplasm and were not present in the nucleus of HeLa cells
stained with DAPI (Figs. 3a–d, 4a–d). In previous studies, it
was found that ruthenium(II) complexes can induce cancer
cell apoptosis by activating the intrinsic mitochondrial path-
ways or endoplasmic reticulum pathways [47, 48]. Therefore,
in this study, colocalization experiments were used to
determine if D-Ru1 and K-Ru1 were localized to the mito-
chondria or endoplasmic reticulum. As shown in Figs. 3h and
4h, excellent superimposition, indicated by the yellow color,
between the signals for D-Ru1 or K-Ru1 (red) and Mito-
Tracker Green FM (green) was observed; however, there was
no overlap between ER-Tracker Green (green) and D-Ru1 or
K-Ru1 (Figs. 3l, 4l). Results were also obtained for D-Ru2
and K-Ru2 and for D-Ru3 and K-Ru3 (Figs. S3–S6). Among
these six ruthenium(II) complexes, the best colocalization
with the mitochondria was for D-Ru1. Taken together, the
ICP-MS and live-cell imaging results indicated that D-Ru1
primarily accumulated in the mitochondria, and thus mito-
chondria may be its primary target. On the basis of its having
the best anticancer effect among these six complexes and its
targeting of mitochondria, the rest of the study focused on the
biological properties of D-Ru1 for further investigation.
Apoptosis assays
To determine whether D-Ru1 causes cell death by apop-
tosis or necrosis, annexin V and DAPI staining assays were
performed. The externalization of phosphatidylserine on
the outer surface of the plasma membrane is a known
hallmark of early-phase apoptosis, and therefore the bind-
ing of annexin V to the cell surface is an indication of
early-phase apoptosis [49, 50]. Flow cytometry revealed
Fig. 2 Subcellular accumulation of ruthenium(II) complexes deter-
mined in HeLa cells after 48 h of exposure to the same concentration
of 20 lM
Fig. 3 Cell localization imaging of D-Ru1 (10 lM). a, e, i Bright
field images. b, f, j Luminescence images of D-Ru1 (red). c Lumi-
nescence image of 40,6-diamino-2-phenylindole (DAPI; blue).
g Luminescence image of MitoTracker Green FM (green). k Lumi-
nescence image of ER-Tracker Green (green). d, h, l Merged pictures.
Scale bar 20 lm
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343
Fig. 4 Cell localization imaging of K-Ru1 (10 lM). a, e, i Bright
field images. b, f, j Luminescence images of K-Ru1 (red). c Lumi-
MitoTracker Green FM (green). k Luminescence image of ER-
Tracker Green (green). d, h, l Merged pictures. Scale bar 20 lm
nescence image of DAPI (blue).
g Luminescence image of
that early-phase apoptosis was induced in HeLa cells by
3 h of treatment with D-Ru1 (15 lM), and as the exposure
time increased, the percentage of annexin V-positive cells
increased (Fig. 5). Nuclear condensation and fragmentation
are considered indicative of late-phase apoptosis [51, 52].
These characteristics can be detected by DAPI, which is a
DNA-binding dye that has been used extensively for
staining cell nuclei [53]. For this purpose, HeLa cells were
treated with D-Ru1 (15 lM) for various time periods, and
then the cell nuclei were stained with DAPI. As expected,
the cell nuclei shrank after exposure for 6 h, and these
effects were severer with increasing exposure time.
Nuclear condensation and fragmentation were obvious
after 48 h (Fig. 6). On the basis of the results of the two
apoptosis assays, we concluded that the cytotoxic effects of
D-Ru1 on HeLa cells are primarily caused by apoptosis.
[55]. It has been reported that ROS have an important role in
the induction of apoptosis by some ruthenium(II) com-
plexes [27, 40, 56]. However, another study showed that
ROS were not produced during treatment with ruthe-
nium(II) complexes [28]. We therefore investigated whe-
ther D-Ru1 could increase the ROS level in HeLa cells.
Flow cytometry showed a slight concentration-dependent
increase in the amount of ROS compared with the amount in
the vehicle-treated cells (Fig. 7a). To further examine the
role of ROS generation in ruthenium(II)-induced cytotox-
icity, we used H2O2 as a positive control and NAC as the
ROS scavenger. The results showed that pretreatment of the
cells with NAC could abrogate both H2O2-induced and
ruthenium(II)-induced ROS generation (Fig. 7b). Further-
more, the presence of NAC significantly suppressed H2O2-
induced cytotoxicity, but interestingly, the presence of
NAC could not reduce the cytotoxicity of D-Ru1 toward
HeLa cells (Fig. 7c). Taken together, these results indi-
cate that D-Ru1 can induce the production of a small
amount of ROS, but ROS production is not the cru-
cial factor in the apoptosis of HeLa cells induced by D-Ru1.
Intracellular ROS study
ROS include the superoxide radical (O2ꢀ-), hydrogen per-
oxide (H2O2), and the hydroxyl radical (OHꢀ). Oxidative
stress generated by an imbalance between ROS and anti-
oxidants contributes to the pathogenesis of cancer, respi-
ratory diseases, and some toxic diseases [54]. NAC is a
source of sulfhydryl groups in cells and is a scavenger of
free radicals as it interacts with ROS such as OHꢀ and H2O2
Induction of mitochondrial dysfunction
Mitochondria play a crucial role in apoptosis triggered by
chemical agents. Chemical-induced apoptosis mediated by
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Fig. 5 Early-phase apoptosis of HeLa cells induced by D-Ru1 was examined by the annexin V assay. HeLa cells were treated with D-Ru1
(15 lM) for 0–36 h and then analyzed by flow cytometry
Fig. 6 Late-phase apoptosis of HeLa cells induced by D-Ru1 was examined by analyzing nuclear fragmentation with DAPI staining. HeLa cells
were treated with D-Ru1 (15 lM) for 0–48 h. Scale bar 20 lm
the mitochondria/apoptotic cascades is often associated
with the collapse of the mitochondrial membrane potential
as a result of the leakage of proapoptotic factors [57]. Very
recently, it has been reported that ruthenium(II) complexes
can induce cell death by interfering with the membrane
potential, finally leading to apoptosis [27, 28, 34, 35]. To
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345
Fig. 7 a Analysis of production of reactive oxygen species (ROS)
after HeLa cells had been treated with D-Ru1 for 6 h. The
intracellular ROS level was detected by flow cytometry (FL-1
channel; excitation at 488 nm and emission at 525 nm). b The ROS
level was expressed as the relative fluorescence intensity measured
with a microplate reader after 6 h of treatment with the ruthenium(II)
complex and H2O2 with or without antioxidants (N-acetylcysteine,
NAC) at the concentrations indicated. The data are presented as
mean the standard deviation and were obtained in three indepen-
dent experiments. c Effects of NAC (10 mM) on D-Ru1-induced or
H2O2-induced cytotoxicity. HeLa cells were exposed to different
doses of D-Ru1 or H2O2 with or without NAC for 48 h. Cell viability
was assessed by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazo-
lium bromide assay. Con control, DCF 20,70-dichlorofluorescein
Apoptosis induced through the intrinsic mitochondrial
pathway
investigate the role of this mechanism, the lipophilic cat-
ionic molecular probe JC-1 was used to determine whether
D-Ru1 can induce alterations in the mitochondrial mem-
brane potential. JC-1 exhibits potential-dependent accu-
mulation in the mitochondria, and the loss of mitochondrial
membrane potential can be visualized by a fluorescence
shift from red (590 nm) to green (525 nm) [58]. After cells
had been treated with D-Ru1 for 6 h, there was a signifi-
cant dose-dependent decrease in the ratio of red to green
fluorescence in HeLa cells (Fig. 8a). In addition, when
HeLa cells were incubated with D-Ru1 at a concentration
of 15 lM, a rapid depolarization of the mitochondrial
membrane potential was detected starting at 0.5 h, reaching
a maximum at 12 h (Fig. 8b). These results are in accor-
dance with the time course of apoptosis observed in HeLa
cells (as showed in Figs. 5, 6) and suggest that the collapse
of the mitochondrial membrane potential is an early event
in D-Ru1-induced apoptosis. As mentioned earlier, cancer
cells have higher mitochondrial membrane potential than
their normal counterparts and are more sensitive to damage
by mitochondria-targeting agents [25, 26], which may
partly account for the toxicity of D-Ru1 toward tumor cells
over nonmalignant cells.
There are two primary signaling pathways involved in
apoptosis: the extrinsic (receptor-mediated) and intrinsic
(mitochondria-mediated) pathways [59]. The crucial step in
the intrinsic pathway is the loss of mitochondrial mem-
brane potential and release of cytochrome c, which occur
as early initiating events before caspase-mediated activa-
tion of apoptotic cascades [60]. The mitochondrial
response to the disruption of the mitochondrial membrane
potential includes the release of cytochrome c into the
cytosol. In the cytosol, cytochrome c binds to Apaf-1,
allowing the recruitment of caspase-9 and the formation of
an apoptosome complex, resulting in caspase-3 activation
and the execution of cell death [61–63]. To assess whether
D-Ru1 initiates apoptosis through this signaling pathway,
we determined whether this complex can trigger the release
of cytochrome c from the mitochondria. Western blot
analysis showed that the cytochrome c level in the whole
cell did not obviously change, whereas the release of
cytochrome c was detected at 1 h in the cytosol after
treatment. This ruthenium(II) complex induced an obvious
time-dependent increase in the cytochrome c level in the
cytosol (Fig. 9a). These results clearly showed that the
induction of apoptosis by D-Ru1 began with the collapse of
the mitochondrial membrane potential, which resulted in
the permeabilization of the mitochondrial outer membrane.
This permeabilization in turn led to the release of cyto-
chrome c from the mitochondria, inducing apoptosis.
Apoptosis is executed by the concerted activation of
caspases, including initiator (caspase-9) and executioner
(caspase-3) caspases. To determine further the involvement
To analyze further the mitochondrial morphological
alterations caused by treatment with the ruthenium com-
plex, transmission electron microscopy studies were per-
formed using HeLa cells that had been treated with D-Ru1
(15 lM) for 24 h. The control cells were characterized by a
normal morphology for the cytoplasm, nucleus, and mito-
chondria (Fig. 8c, images a and b). In contrast, the treated
cells had an apoptotic morphology and abnormally swollen
mitochondria (Fig. 8c, images c and d), indicative of the
loss of mitochondrial membrane potential [56].
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Fig. 8 a Depletion of mitochondrial membrane potential in HeLa
cells after treatment with D-Ru1 for 6 h and analyzed by flow
cytometry. The data are the mean of three independent experiments.
b Fluorescence images of 5,50,6,60-tetrachloro-1,10,3,30-tetraethylben-
zimidazolylcarbocyanine iodide-labeled HeLa cells treated with
D-Ru1 (15 lM) for 0–12 h taken with an inverted fluorescence
microscope. Scale bar 20 lm. c Impact of D-Ru1 on mitochondrial
morphology observed by transmission electron microscopy: control
(a); magnified normal mitochondria (b); HeLa cells treated with
D-Ru1 (15 lM) for 24 h (c); magnified abnormal (swollen) mito-
chondria (d). Scale bar 1 lm for a and c, and 0.1 lm for b and d
of caspase-9 and caspase-3 in D-Ru1-induced apoptosis, we
determined the levels of the uncleaved and cleaved (active)
forms of caspase-9 and caspase-3 in cell lysates. As shown in
Fig. 9b, the level of uncleaved caspase-9 (47 kDa) was
reduced in parallel with a time-dependent increase in the
level of its active subunit (35 kDa) after 6 h of treatment
with D-Ru1. Subsequently, the active cleaved form of cas-
pase-3 (17 kDa) was detected after caspase-9 activation at
24 h, consistent with the fact that caspase-3 is activated by
caspase-9 during apoptosis. As an irreversible hallmark of
apoptosis [64], the cleavage of caspase-3 (17 kDa) indicates
that HeLa cells treated with D-Ru1 completed the apoptosis
process. All of these results together indicate that D-Ru1-
induced apoptotic death is mediated by mitochondria-
dependent signaling pathways. Prosurvival factors such as
Bcl-2 and Bcl-xl can maintain the integrity of the mito-
chondrial outer membrane, whereas proapoptotic factors,
such as Bax, can disrupt its integrity, causing the release of
proapoptotic factors into the cytosol. Increasing the Bax to
Bcl-2 or Bax to Bcl-xl ratio could induce the permeabiliza-
tion of mitochondria [65, 66]. Many studies have reported
that Bcl-2 family members are important key regulators of
mitochondrial function during apoptosis induced by ruthe-
nium(II) complexes [27, 34–36, 40]. Although most studies
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347
D-Ru1 treatment greatly downregulated the expression level
of the prosurvival protein Bcl-xl after 6 h, and the Bcl-2
expression level was decreased from 12 h. In contrast, the
expression level of the proapoptotic protein Bax was
increased very slightly until 48 h (Fig. 9c). Hence, as a result
of these changes, the Bax to Bcl-2 and Bax to Bcl-xl ratios
increased dramatically, reducing the integrity of the mito-
chondrial outer membrane. However, these events occurred
after the loss of mitochondrial membrane potential, and
combined with the results above, this timing indicates that
although Bcl-2 family members play a crucial role in
apoptosis induced by ruthenium(II) complexes, they appear
to be synergistic factors rather than initiating factors in the
process of apoptosis.
Conclusions
Six chiral ruthenium(II) complexes containing PIP deriva-
tives were synthesized and characterized. Live-cell imaging
and quantitative analysis by ICP-MS demonstrated that the
ruthenium(II) complexes tended to accumulate preferen-
tially in the mitochondria of HeLa cells. These ruthenium(II)
complexes exhibited different inhibitory effects on cancer
cells. Among these complexes, D-Ru1 was found to have
IC50 values relatively close to those of the well-known drug
cisplatin for the cancer cell lines, and this complex was more
cytotoxic to tumor cells than to normal cells. In addition, the
cisplatin-resistant cell line A549-CP/R exhibited higher
sensitivity to D-Ru1 than to cisplatin. Further analysis
showed that D-Ru1 exerts its toxicity through the intrinsic
mitochondria-mediated apoptotic pathway, which is
accompanied by the regulation of Bcl-2 family members and
the activation of caspases. Major challenges in cancer ther-
apy are identifying and synthesizing drugs that kill tumor
cells while preserving normal cells. Thus, D-Ru1 may be a
more potent and selective chemotherapeutic agent.
Fig. 9 Effect of D-Ru1 on the apoptotic pathway in HeLa cells.
a Release of cytochrome c. HeLa cells were treated with D-Ru1
(15 lM) for 0–24 h. Cells were homogenized, and lysates (cytosolic
fraction and the whole cells) were assayed for cytochrome c levels
(C-Ctoc-c is the cytochrome c level in the cytosol, and W-Ctoc-c is
the cytochrome c level in the whole cell) using Western blot analysis.
b, c Effect of D-Ru1 on the expression of apoptosis-related proteins.
HeLa cells were treated with D-Ru1 (15 lM) for 0–48 h. Whole-cell
lysate was prepared and subjected to Western blotting for cyto-
chrome c, caspase-3, cleaved caspase-3 (C-caspase-3), caspase-9,
Bax, Bcl-2, and Bcl-xl. GAPDH glyceraldehyde 3-phosphate
dehydrogenase
Acknowledgments This work was supported by the National Sci-
ence of Foundation of China (nos. 21071155, 21172273, 21171177),
the National High Technology Research and Development Program
of China (863 Program, 2012AA020305), the National Science of
Foundation of Guangdong Province (9351027501000003), the
Research Fund for the Doctoral Program of Higher Education
(20110171110013), the State Key Laboratory of Optoelectronic
Materials and Technologies (2010-ZY-4-5), and Sun Yat-Sen Uni-
versity. X.H. thanks the Science and Technology Department of
Hunan Province for support (2010FJ4090).
have concentrated on the dose–response relationship, in this
study, we assessed the time dependence of the effects on Bcl-
2 family members to elucidate the detailed process by which
the ruthenium(II) complex induces apoptosis. We found that
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