Jakub Cervinka1,2, Alberto Gobbo3,4, Lorenzo Biancalana3, Lenka Markova1, Vojtech Novohradsky1, Massimo Guelfi3, Stefano Zacchini4, Jana Kasparkova1,5, Viktor Brabec1, Fabio Marchetti3. 1. Czech Academy of Sciences, Institute of Biophysics, Kralovopolska 135, CZ-61265 Brno, Czech Republic. 2. Faculty of Science, Department of Biochemistry, Masaryk University, Kamenice 5, CZ-62500 Brno, Czech Republic. 3. Department of Chemistry and Industrial Chemistry, University of Pisa, Via G. Moruzzi 13, I-56124 Pisa, Italy. 4. Department of Industrial Chemistry "Toso Montanari", University of Bologna, Viale Risorgimento 4, I-40136 Bologna, Italy. 5. Faculty of Science, Department of Biophysics, Palacky University in Olomouc, Slechtitelu 27, CZ-78371 Olomouc, Czech Republic.
Abstract
While ruthenium arene complexes have been widely investigated for their medicinal potential, studies on homologous compounds containing a tridentate tris(1-pyrazolyl)methane ligand are almost absent in the literature. Ruthenium(II) complex 1 was obtained by a modified reported procedure; then, the reactions with a series of organic molecules (L) in boiling alcohol afforded novel complexes 2-9 in 77-99% yields. Products 2-9 were fully structurally characterized. They are appreciably soluble in water, where they undergo partial chloride/water exchange. The antiproliferative activity was determined using a panel of human cancer cell lines and a noncancerous one, evidencing promising potency of 1, 7, and 8 and significant selectivity toward cancer cells. The tested compounds effectively accumulate in cancer cells, and mitochondria represent a significant target of biological action. Most notably, data provide convincing evidence that the mechanism of biological action is mediated by the inhibiting of mitochondrial calcium intake.
While ruthenium arene complexes have been widely investigated for their medicinal potential, studies on homologous compounds containing a tridentate tris(1-pyrazolyl)methane ligand are almost absent in the literature. Ruthenium(II) complex 1 was obtained by a modified reported procedure; then, the reactions with a series of organic molecules (L) in boiling alcohol afforded novel complexes 2-9 in 77-99% yields. Products 2-9 were fully structurally characterized. They are appreciably soluble in water, where they undergo partial chloride/water exchange. The antiproliferative activity was determined using a panel of human cancer cell lines and a noncancerous one, evidencing promising potency of 1, 7, and 8 and significant selectivity toward cancer cells. The tested compounds effectively accumulate in cancer cells, and mitochondria represent a significant target of biological action. Most notably, data provide convincing evidence that the mechanism of biological action is mediated by the inhibiting of mitochondrial calcium intake.
Complexes of d-block metals possess unique
properties otherwise
not available to organic compounds and thus offer significant medicinal
potential.[1−3] In particular, platinum compounds have been used
in clinical treatments against various types of cancer;[4−6] however, despite their undisputed efficacy, they exhibit serious
drawbacks, such as negative side effects, phenomena of intrinsic or
acquired resistance, a limited number of treatable tumors, and the
necessity of hospitalization for the intravenous administration.[7−10] These facts have stimulated research to develop new drugs based
on other transition-metal elements.[11−13] Specifically, a variety
of ruthenium complexes have shown great promise.[14,15] Besides the prototypal NAMI-A, KP1019, and related ruthenium(III)
salts that entered clinical trials,[14,16,17] half-sandwich organometallic complexes based on the
RuII–arene scaffold have attracted considerable
attention. In particular, RAPTA compounds, featured by the amphiphilic
1,3,5-triaza-7-phosphaadamantane ligand (PTA), have emerged as prominent
and are currently pointing to clinical trials.[18,19] The popularity of RAPTAs and the easy accessibility of related structures
have steered the way to the exploration of a considerable number of
derivatives with a diversity of arenes and coligands (Figure A,B).[20−22] However, a
suitable combination of electronic factors should be formulated to
avoid the removal of the arene moiety and the consequent disaggregation
of the complexes in aqueous media, which is a disliking characteristic
for a drug candidate.[23−27]
Figure 1
(A)
structure of RAPTA-C, leading compound of the RAPTA family;
(B) general structure of ruthenium(II)–arene complexes investigated
as anticancer drugs (R = alkyl/aryl; X = neutral or ionic ligand;
L,L′ = pair of neutral ligands of bidentate neutral/ionic ligand);
(C, D) structures of ruthenium–tpm compounds assessed for their
cytotoxicity (L = MeCN, DMSO, PMePh2; n = 2–4).
(A)
structure of RAPTA-C, leading compound of the RAPTA family;
(B) general structure of ruthenium(II)–arene complexes investigated
as anticancer drugs (R = alkyl/aryl; X = neutral or ionic ligand;
L,L′ = pair of neutral ligands of bidentate neutral/ionic ligand);
(C, D) structures of ruthenium–tpm compounds assessed for their
cytotoxicity (L = MeCN, DMSO, PMePh2; n = 2–4).Tris(pyrazolyl)methane (tpm) and ring-substituted
derivatives are
homologous to arenes in that both types of compounds are neutral and
may behave as six-electron ligands towards transition-metal centers.
However, while the arene–metal bond possesses a π-backbonding
component, tpm is essentially a strong donor and provides substantial
stability to the resulting complexes.[28,29] Moreover,
tripodal coordination with three nitrogen atoms (κ3) is usually observed, although alternative modes are possible,[30,31] and the interchange between tri- and bidentate coordination might
play some key role in metal-catalyzed organic transformations.[32−34] In sharp contrast with the related arene systems and the fact that
NAMI-A and KP1019 contain a mono-pyrazolyl ring, ruthenium(II)–tpm
complexes remain almost unexplored for their medicinal potential heretofore.
Indeed, to the best of our knowledge, studies are limited to sparse
DNA binding experiments[35,36] and the assessment
of the in vitro cytotoxicity of complexes [RuCl(κ3-tpm)(L)2]PF6 (L = MeCN, DMSO, PMePh2) and [RuCl(κ3-tpm)(LL)]PF6 (LL=
diphosphine) against breast MCF-7 and cervical HeLa cancer cell lines
(Figure C,D).[37] In the latter case, while the MeCN and DMSO
adducts revealed inactive, the introduction of phosphine ligands resulted
in IC50 values falling in the low micromolar range. Presumably,
the lack of biological studies is also a consequence of the paucity
of straightforward routes to add diversity to the {RuII–tpm} scaffold.Here, we report a general synthetic
strategy to access a new family
of robust ruthenium(II)–tpm complexes and an extensive investigation
of their anticancer activity.
Results and Discussion
Synthesis and Structural Characterization of Ruthenium Complexes
The ruthenium–tpm complex 1, containing two
triphenylphosphine ligands, was obtained from commercial RuCl3·3H2O via a straightforward two-step procedure
that was optimized with respect to the literature (Scheme ).[38,39] In particular, toluene was found to be the optimal solvent for the
reaction of tpm with [RuCl2(PPh3)3], allowing to collect the desired product in 95% yield (gram scale).
Following a previous work reporting the thermal substitution of one
PPh3 with PTA (1,3,5-triaza-7-phosphaadamantane),[40] we investigated the possibility of modifying
the ruthenium coordination set in 1 by introducing different
types of ligands (i.e., different N-heterocyclic ligands; a phosphite
that should provide more hydrophilicity than PPh3; two
isocyanides with an aryl and alkyl substituent, respectively). Quantitative
PPh3/MeCN replacement was achieved by heating a solution
of 1 in acetonitrile at reflux, affording complex 2 in 95% yield (Scheme ). Note that nitrile ligands usually behave as labile ones
and are easily replaced by phosphines when coordinated to group 8[41−44] or other transition metals.[45,46] In the present case,
the reversed reaction is probably favored by steric factors arising
from two bulky triphenylphosphines occupying adjacent coordination
sites in 1.[38]
Scheme 1
Synthesis
of Ruthenium(II) Tris(pyrazolyl)methane Complexes Investigated
in This Work (1–9); Cy = C6H11
To enable other substitution reactions, we found
ethanol to be
the best solvent; the reactions of 1 with a slight excess
of various N-heterocyclic donors, trimethylphosphite and isocyanides,
were carried out in ethanol at reflux and proceeded straightforwardly
to afford novel complexes 3–9, which
were isolated in good to nearly quantitative yields (Scheme ).The IR spectra of 4 and 5 (solid state, Figures S1–S9 in the Supporting Information)
display the absorption attributed to the NH moiety at ca. 3450 cm–1, while the OH group belonging to 6 was
detected at 3668 cm–1. The intense band due to the
carbon–nitrogen triple bond in the isocyanide complexes occurs
at 2141 (CNCy, 8) and 2147 cm–1 (CNCH2P(O)(OEt)2, 9). These values are almost
coincident with those of the respective free isocyanides, indicating
a scarce metal to isocyanide backdonation.[47−51]The NMR spectra of 2–9 (in
CDCl3, see Figures S10–S35) contain
single sets of resonances, and the signals related to tpm are not
significantly affected by the nature of the varying ligand (L). More
precisely, the ring carbons resonate in the ranges 149.2–144.0
ppm (Cα), 109.0–107.3 ppm (Cβ), and 136.0–133.4 ppm (Cγ), whereas the
methylidyne group gives rise to resonances at about 12 ppm (1H) and 74 ppm (13C). Inequivalence of the NMR resonances
of the three pyrazolyl rings is in accordance with the chirality of
the metal atom. The 31P NMR spectra display the resonance
related to PPh3 falling at 44.8 (7)–52.5
(3) ppm; the resonance associated with the additional
phosphorus ligand in 9 and 7 occurs at 15.7
and 138.0 ppm, respectively.The structures of 3, 4, 5, 6, 7,
and 8 were confirmed
by single-crystal X-ray diffraction analyses; views of these structures
are shown in Figure with relevant bond lengths and angles reported in Table . Complexes 3–8 display a distorted octahedral geometry, as found in precursor 1 and related [RuCl(κ3–tpm)(PPh3)(L)]+ complexes.[37,38,52] The Ru(1)–N(3) and Ru(1)–N(5) distances
(Table ) are comparable
in 3–8 since they are trans to Cl(1) and P(1),
respectively, in all complexes. In contrast, the Ru(1)–N(1)
bond displays similar values in 3–6 [2.056(4),
2.045(5), 2.070(4), and 2.077(3) Å], being in the trans position to an aromatic N(7) ligand, whereas it is slightly elongated
in 7 [2.180(2) Å] and 8 [2.1428(19)
Å], where better π-acceptors P(OMe)3 and CNCy
are present in the trans position. The nature of
the varying ligand, L, slightly affects the Ru(1)–Cl(1) distance,
reaching the lowest values in complexes 7 and 8 [2.3950(10) and 2.3942(6) Å, respectively]; the strength of
the ruthenium–chloride bond is likely to be correlated with
the cytotoxic activity of the complexes (vide infra). The Ru(1)–C(21)
bond length in 8 [1.918(2) Å] is the shortest reported
for an octahedral RuII center bonded to CNXyl, for which
typical Ru–C values fall in the range 1.95–2.04 Å.[53−56] This is likely to be since CNXyl is trans to N(1)
in 8.
Figure 2
View of the molecular structures (ORTEP drawing) of the
cations
of 3, 4, 5, 6, 7, and 8. Displacement ellipsoids are at the
30% probability level. CCDC reference numbers are 2167597 (3), 2167598 (4), 2167599 (5), 2167600 (6), 2167601 (7), and 2167602 (8).
Table 1
Selected Bond Lengths (Å) and
Angles (°) for 3, 4, 5, 6, 7, and 8
3
4
5
6
7
8
Ru(1)–N(1)
2.056(4)
2.045(5)
2.070(4)
2.077(3)
2.180(2)
2.1428(19)
Ru(1)–N(3)
2.060(4)
2.055(5)
2.069(4)
2.061(3)
2.085(2)
2.0798(17)
Ru(1)–N(5)
2.103(4)
2.106(5)
2.104(4)
2.100(3)
2.131(2)
2.106(2)
Ru(1)–P(1)
2.3063(13)
2.3021(16)
2.3252(13)
2.3184(8)
2.3209(9)
2.3125(6)
Ru(1)–Cl(1)
2.3980(13)
2.4045(15)
2.4252(12)
2.4021(8)
2.3950(10)
2.3942(6)
Ru(1)–Xa
2.076(4)
2.103(5)
2.082(4)
2.094(3)
2.2181(10)
1.918(2)
N(1)–Ru(1)–N(3)
87.66(17)
87.71(18)
87.68(16)
87.29(11)
87.81(8)
87.79(7)
N(1)–Ru(1)–N(5)
84.36(17)
84.39(19)
85.02(17)
84.69(10)
81.02(8)
82.67(7)
N(3)–Ru(1)–N(5)
84.23(17)
84.1(2)
83.50(17)
84.15(11)
82.88(9)
82.06(7)
P(1)–Ru(1)–Cl(1)
95.74(5)
98.16(6)
96.68(4)
97.75(3)
91.04(3)
92.98(2)
P(1)–Ru(1)–Xa
94.14(12)
95.35(14)
94.84(12)
95.51(8)
93.35(3)
92.44(7)
Cl(1)–Ru(1)–Xa
88.62(12)
87.92(14)
87.37(12)
88.03(8)
96.29(3)
91.20(6)
X = N(7), 3–6; P(2), 7; C(21), 8.
View of the molecular structures (ORTEP drawing) of the
cations
of 3, 4, 5, 6, 7, and 8. Displacement ellipsoids are at the
30% probability level. CCDC reference numbers are 2167597 (3), 2167598 (4), 2167599 (5), 2167600 (6), 2167601 (7), and 2167602 (8).X = N(7), 3–6; P(2), 7; C(21), 8.The ligands of complexes 4, 5, and 6 contain NH (4 and 5) or OH (6) groups involved in hydrogen bonds. In particular,
the N(8)H(8)
benzimidazole group of 4 forms a hydrogen bond with the
chloride counterion Cl(3) [N(8)–H(8) 0.88 Å, H(8)···Cl(3)
2.15 Å, N(8)···Cl(3) 3.027(6) Å, N(8)–H(8)–Cl(3)
173.7°]. Similarly, the O(1)H(1a) group of 6 is
involved in a similar H-bond with the counterion [O(1)–H(1a)
0.848 Å, H(1a)···Cl(2) 2.19 Å, O(1)···Cl(2)
3.027(3) Å, O(1)–H(1a)–Cl(2) 175.7°]. In contrast,
the N(8)H(8) group of the pyrazole ligand of 5 forms
an intramolecular H-bond with the Cl(1) ligand [N(8)–H(8) 0.88
Å, H(8)···Cl(1) 2.525 Å, N(8)···Cl(1)
3.039(5) Å, N(8)–H(8)–Cl(1) 118.2°].
Solubility, Partition Coefficient, and Stability in Aqueous
Media
A detailed study on the behavior of complexes 1–9 in aqueous media was performed: experimental procedures
are provided in Experimental Section, and
the results are presented in Table .
Table 2
Solubility in Water (D2O), Octanol/Water Partition Coefficient (log Pow), and Residual Ruthenium Complex in D2O
(after 48 h) and DMSO-d6/DMEM-d (1:4 v/v, Except 1:3 v/v in the Case of 4; after 24 h) Solutions Maintained at 37 °C
compound
solubility/10–3 mol·L–1 (D2O, 21°C)a
log Pow
residual
complex % in D2Oa,b
residual complex
% in DMEM-d/ DMSO-d6a,b
1
1.18 ± 0.05
69c
88
2
4.5
–0.33 ± 0.07
89
51
3
3.4
–0.15 ± 0.03
100
100
4
2.4
0.58 ± 0.06
100
93
5
3.4
–0.05 ± 0.05
100
98
6
1.9
1.11 ± 0.07
100
100
7
3.1
–0.02 ± 0.05
100
95
8
1.1
0.34 ± 0.01
92
98
9
4.6
–0.20 ± 0.05
93
97
Calculated by 1H NMR
(Me2SO2 as the internal standard).
Sum of chloro (2–9) and aquo (2–9) complexes.
DMSO-d6/D2O (4:1 v/v) mixture.
Calculated by 1H NMR
(Me2SO2 as the internal standard).Sum of chloro (2–9) and aquo (2–9) complexes.DMSO-d6/D2O (4:1 v/v) mixture.NMR experiments (1H and 31P)
were used to
study the speciation of 2–9 in D2O; preparation of the samples required 2 h (t0) stirring of a suspension of each complex in D2O; then, NMR analyses of the solutions pointed out the occurrence
of a chloride–water exchange process (Scheme ).
Scheme 2
Chloride Dissociation Reaction of
Ruthenium(II) Tris(pyrazolyl)methane
Complexes in Water
NMR data of the resulting dicationic aquo species 2–9 are reported in the Supporting Information (Figures S36–S43); the representative spectra
of 3 dissolved in D2O are shown in Figures S44–S45. The aquo complexes were
detected in
a variable relative amount according to the ligand L (0–90%)
after 2 h, as sharp NMR signals; 2–9 became the
largely prevalent species in solution after 48 h in all cases (see
below) when the system had probably reached equilibrium. A significant
increase of the molar conductivity of a 10–3 M solution
of 6 in H2O was recognized over 24 h, in alinement
with the increase by one unit of the net cationic charge of the ruthenium
species (see Experimental Section). The pH
of a 10–3 M H2O solution of 3, this compound undergoing almost complete aquation over 48 h (Table S1), was monitored over time, revealing
a substantially constant and close to neutrality value [pH(t0) = 6.78, pH(24h) = 6.66, pH(48h) = 6.36]. This evidence reveals that the chloride/water
exchange process is not followed by deprotonation of the H2O ligand, differently from what is documented for RAPTA compounds.[57] The chloride/water substitution is fully reversible,
in that a mixture of 3 and 3 (isolated from the D2O solution) fully reverted
to 3 in CD3OD during 48 h (see page S32 and Figures S48–S49 in the SI).Note that leading ruthenium(II)–arene complexes, such as
RAPTA compounds, are considered prodrugs, and it has been proposed
that the dissociation of chloride ligand(s) in the physiological environment
is key to activation, enabling the binding of the metal center with
biosubstrates.[18,58−60] However, the
kinetics of chloride/water exchange typically occurs in a much shorter
time scale (≤30 min) for the arene complexes.[58−60]Compound 1 is insoluble in water, and its behavior
was evaluated in a DMSO-d6/D2O mixture. Under these conditions, chloride displacement was not
recognized, whereas partial substitution of PPh3 with one
solvent molecule was suggested by the appearance of a new signal at
−7.2 ppm in the 31P NMR spectrum.The D2O solubility of 2–9 was determined
by 1H NMR using dimethyl sulfone (Me2SO2) as the internal standard, ranging between 1.1 and 4.6 mM,
with reference to the sum of the chloro and aquo complex observed
after 2 h (see above).The octanol–water partition coefficients
(log Pow) of 1–9 were measured
by a UV–vis method; log Pow values of 2–9 were obtained approx. 20 min after
the dissolution and thus are representative of the monocationic chloro
complexes 2–9, as suggested by 1H and 31P NMR spectra recorded on a D2O solution of 3 after the same time (Figures S46–S47). The log Pow values are compiled in Table and reflect an overall amphiphilic or moderate
lipophilic character. More in detail, 1, which contains
two triphenylphosphine ligands, and 6, featured by a
hydroxy-pyridine ligand, revealed to be the most lipophilic compounds
of the series. Conversely, the incorporation of acetonitrile, 1-methylimidazole,
pyrazole, trimethylphosphite, and diethyl isocyanomethyl phosphonate
as ligands leads to slightly negative log Pow values.The D2O solutions of RuII–tpm complexes
were then maintained at 37 °C and monitored for 48 h. Apart from
the chloride dissociation process, no other changes were observed,
except for a minor degradation in the cases of 2, 8, and 9, ascribable to the release of L (see Experimental Section for details). In fact, signals
of dissociated PPh3 and tpm were not found in the NMR spectra.The behavior of 1–9 was also analyzed in deuterated
cell culture medium. The solutions were diluted with a variable amount
of DMSO-d6 to obtain an appreciable solubility
and then kept at 37 °C for 24 h. The compounds were found to
be stable even in these conditions, and only minor degradation of
the starting materials was detected by 1H and 31P NMR (1–12%, 49% for 2). The 31P
NMR spectra of 1 and 3–9 resembled
those recorded in D2O–DMSO-d6 and D2O solutions, respectively, while four different
signals were detected in the final 31P NMR spectrum of 2 (Figure S50). In the case of 2, in addition to chloride/water exchange, it is possible
that replacement of the acetonitrile ligand by solvent molecules occurs.
On the other hand, the NMR spectra of 3–9 in DMSO–DMEM
after 24 h only contained the resonances related to the starting complexes
and the respective aquo species; therefore, DMSO coordination must
be ruled out. The presence of ca. 0.1 M chloride ion in the medium
slowed down the chloride/water substitution from 1–9, being almost negligible in the initial solution, and decreased
the relative amount of aquo species after 48 h with respect to the
analogous experiments in pure D2O (see Table S1 in the Supporting Information). The lower percentage
of the aquo complex is exhibited by 7, 8, and 9, in accordance with the electron-withdrawing
property of trimethylphosphite and isocyanide ligands, presumably
strengthening the ruthenium–chloride bond.[61,62]
Antiproliferative Activity
Primary screening of antiproliferative
activity of the family of ruthenium–tpm complexes 1–9 was performed by the commonly used MTT assay on five cancer cell
lines of various origins: MCF-7 (breast), HeLa (cervical), 518A2 (melanoma),
HCT116 (colon), and RD (rhabdomyosarcoma). In addition, normal human
fibroblasts MRC5pd30 were used to assess the toxicity of the complexes
on a noncancerous cell line. The results are summarized in Table . The IC50 values (concentration that causes 50% inhibition of cell proliferation)
obtained for 1–9 are compared to those obtained
for clinically used cisplatin under the same experimental conditions.
Table 3
IC50 Values (μM)
Determined by the MTT Test after 72 h of Treatmenta
MCF-7
HeLa
518A2
HCT116
RD
MRC5pd30
SIb
1
2.4 ± 0.6
4.0 ± 0.4
2.6 ± 0.4
1.5 ± 0.1
2.2 ± 0.2
5.8 ± 0.7
2.3
2
32 ± 4
53 ± 4
26 ± 4
25 ± 2
26 ± 5
76 ± 1
2.4
3
38 ± 8
91 ± 3
33 ± 5
25 ± 1
27 ± 4
83.6 ± 0.5
2.0
4
37 ± 7
36 ± 1
35 ± 4
25 ± 2
25 ± 3
56.6 ± 0.7
1.8
5
32 ± 5
54 ± 6
31 ± 3
30 ± 2
23 ± 2
63 ± 4
1.9
6
46 ± 7
54 ± 14
38 ± 8
31 ± 2
38 ± 6
82 ± 4
2.0
7
6 ± 1
10 ± 2
6.8 ± 0.8
6.7 ± 0.4
6 ± 1
24 ± 1
3.4
8
10 ± 2
15 ± 1
10 ± 2
8 ± 2
6.6 ± 0.7
19.7 ± 0.4
2.0
9
43 ± 7
36 ± 2
38 ± 7
35 ± 6
24 ± 3
50.8 ± 0.4
1.5
cisplatin
13 ± 3c
14 ± 3c
2.6 ± 0.7d
8 ± 1c
4.6 ± 0.3
11.7 ± 0.8c
1.1
The results are expressed as mean
values ± SD from at least three independent experiments.
Selectivity index (SI) was calculated
as IC50 for noncancerous MRC-5pd30 vs the average IC50 value of cancer cell lines.
Data taken from ref (63).
Data
taken from ref (64).
The results are expressed as mean
values ± SD from at least three independent experiments.Selectivity index (SI) was calculated
as IC50 for noncancerous MRC-5pd30 vs the average IC50 value of cancer cell lines.Data taken from ref (63).Data
taken from ref (64).Most of the investigated compounds possess a moderate
activity
with IC50 values in tens of micromolar. The antiproliferative
activity appears significantly influenced by the nature of the ligand
(L) and only partially correlated with the log Pow values. Thus, among the tested compounds, three complexes
stand out. Namely, compound 1 featured the highest degree
of lipophilicity within the series (log Pow = 1.18) and 7 showed the best antiproliferative
activity with IC50 values in a single-digit micromolar
range in almost all of the investigated cancer cell lines. Furthermore,
the potency of these two complexes is comparable to or even better
than that of clinically used cisplatin, depending on the specific
cell line. On the other hand, the substantially lower activity of 6 (log Pow = 1.11), compared
to that of 1, is associated with the presence of the
benzimidazole ligand in place of triphenylphosphine.In addition
to 1 and 7, complex 8 (log Pow = 0.34), bearing
a cyclohexyl isocyanide ligand, also showed very good activity, still
roughly comparable to cisplatin. The beneficial effect of incorporating
the cyclohexyl moiety (Cy) within anticancer metal compounds was previously
recognized and attributed to its compact and hydrophobic structure.[65−67]Notably, the cytotoxic effects of 1–9 on
noncancerous
human skin fibroblasts MRC5pd30 were significantly lower, demonstrating
selectivity toward cancer over noncancerous cells.It should
be mentioned that complexes of formula [RuCl(κN-Py)(PPh3)(η6-p-cymene)]+ (Py = substituted pyridine) were previously
assessed for their cytotoxicity toward the HL60 leukemia tumor cell
line, displaying IC50 values in the 5–15 μM
range.[68]The MTT assay, employed
in the previously described experiment,
is based on the mitochondrial reduction of MTT dye to formazan in
living cells. However, several ruthenium complexes have been shown
to affect mitochondrial metabolism.[69,70] Moreover,
the ruthenium–tris-pyrazolylmethane complexes studied in this
work contain triphenylphosphine, which is known to impart specific
features and activate additional modes of action to the related metal
complexes.[68,71,72] The data obtained by the MTT assay could then be overestimated due
to the possible impact of the investigated compounds on mitochondrial
metabolism. Therefore, the experiments focused on antiproliferative
activity were repeated using an assay based on a mechanism other than
mitochondrial metabolism, namely, Sulphorhodamine B (SRB) assay. This
method relies on the stoichiometric binding of SRB dye to proteins
in cells. The amount of dye is a proxy for cell mass and thus the
number of cells in a sample. For SRB testing (and all further experiments
as well), the HCT116 cell line was chosen in which the most active
compound 1 exhibited the most promising anticancer activity
(the lowest IC50 value).As shown in Table S2 in the Supporting
Information, the values obtained by the SRB assay were similar to
(in the range of experimental error) or slightly lower than those
from the MTT assay. This indicates that mitochondrial succinate dehydrogenase
is likely not inhibited by the ruthenium complexes. Notably, the SRB
assay confirmed the same trend in the biological activity of all tested
complexes as found by MTT, with 1 being the most effective
drug, followed by 7 and 8.
Intracellular Accumulation
To reveal a possible relationship
between the cellular uptake and the in vitro activity
of the investigated complexes, the ruthenium content of HCT116 cells
after 24 h of incubation with complexes 1–9 was
determined by inductively coupled plasma mass spectrometry (ICP-MS).
The viabilities of the cells after the treatment ranged from 93 to
97%, as verified by the trypan blue exclusion assay so that the results
were not affected by elevated permeability of compromised cell membranes
of dying/dead cells. The results are summarized in Table .
Table 4
Accumulation of Ruthenium in HCT116
Cells after Treatment with Ruthenium Complexes (15 μM, 24 h)a,b
ng Ru/106 cells
log Pow
1
145.1 ± 5.9
1.18 ± 0.05
2
29.6 ± 2.9
–0.33 ± 0.07
3
66.1 ± 8.0
–0.15 ± 0.03
4
54.9 ± 1.3
0.58 ± 0.06
5
58.4 ± 3.0
–0.05 ± 0.05
6
27.2 ± 1.1
1.11 ± 0.07
7
63.3 ± 6.0
–0.02 ± 0.05
8
59.4 ± 0.6
0.34 ± 0.01
9
16.2 ± 1.4
–0.20 ± 0.05
log Pow values are also reported for comparison.
Data for intracellular Ru concentration
represent the mean ± standard error of the mean (SEM) from two
independent experiments.
log Pow values are also reported for comparison.Data for intracellular Ru concentration
represent the mean ± standard error of the mean (SEM) from two
independent experiments.The inspection of data in Tables and 4 reveals a correlation
between antiproliferative activity and intracellular accumulation
of the tested ruthenium complexes (Pearson’s correlation coefficient
calculated for IC50 and accumulated Ru r = −0.76, see also Figure S51).
This indicates that the ability of individual complexes to cross the
cell membrane and accumulate in cells significantly contributes to
their respective biological activity. In agreement with this view,
the most active complex 1 accumulates in cells much more
effectively than the other compounds.The overall collection
of data (speciation in aqueous solutions,
log Pow, average IC50, cellular uptake) suggests that the antiproliferative activity of
the complexes might be related to adequate lipophilicity of the active
species. This condition can be achieved with coligand L (see Scheme ) providing hydrophobic
character and/or disfavoring the conversion into more hydrophilic
biscationic complexes via chloride/water exchange (Scheme ). More precisely, 7 and 8 display log Pow values of −0.02 and 0.34, respectively; however the electron-withdrawing
character of L limits the elimination of the chloride and the consequent
formation of 7–8 (16% in DMEM-d/DMSO-d6 after 24 h, see Table S1). On the other hand, for instance, 4 and 6 are considerably more lipophilic than 7 and 8, but the former complexes generate a
higher fraction of biscations (4 and 6, respectively, 59 and
70% in DMEM-d/DMSO-d6), which are expected to be less prone than the parent monocationic
complexes to pass through the cell membrane.[73] Structural factors might also disfavor the penetration of the membrane
for [{Ru}–OH2]2+ species compared to
the corresponding [{Ru}–Cl]+.The three most
potent compounds, i.e., 1, 7, and 8, were selected for further studies to elucidate
their mechanism of action.
Mechanism of Cell Death Induced by Ru Complexes
The
previous test revealed that complexes 1, 7, and 8 possess an interesting antiproliferative activity.
However, these tests cannot distinguish between the cytostatic (growth
arrest and inhibition of division) and cytotoxic (loss of viability)
effects.[74,75] Therefore, we were interested in whether
such complexes could induce cell death and, if so, by what mechanism.
For this purpose, a commonly used Annexin-V/Propidium iodide (PI)
assay was used, and the results were evaluated by flow cytometry (FACS).
The typical densitograms obtained from FACS are shown in Figure S52, and Figure shows a quantitative evaluation of the results.
All tested complexes effectively induced apoptosis in HCT116 cells
(Annexin-V-positive/PI-negative cells), whereas the percentage of
cells undergoing necrosis was negligible (0.2–0.3%). This result
was confirmed by measuring apoptosis/necrosis in real time immediately
after the treatment (Figure S53).
Figure 3
Bar graph of
early apoptotic cells (% of the total population)
in HCT116 cells treated with Ru complexes 1, 7, and 8 for 24 h at their equitoxic concentration (4xIC50,72h) quantified by FACS. Positive controls staurosporine
(10 μM) and EtOH (5% v/v) were included as well-known apoptosis
and necrosis inducers, respectively. Data represent mean ± SEM
from three independent measurements; * indicates a significant (p < 0.05) difference from control, untreated cells.
Bar graph of
early apoptotic cells (% of the total population)
in HCT116 cells treated with Ru complexes 1, 7, and 8 for 24 h at their equitoxic concentration (4xIC50,72h) quantified by FACS. Positive controls staurosporine
(10 μM) and EtOH (5% v/v) were included as well-known apoptosis
and necrosis inducers, respectively. Data represent mean ± SEM
from three independent measurements; * indicates a significant (p < 0.05) difference from control, untreated cells.
Intracellular Distribution of Ru
To assess the subcellular
distribution of selected complexes (1, 7, and 8) in HCT116 cells, cell fractionation was carried
out following the treatment for 5 and 24 h. A FractionPREP Cell Fractionation
kit (BioVision) was employed for the assay. According to the manufacturer’s
information, the isolated fractions contain the nucleus (total nucleus
soluble proteins, including the nuclear membrane proteins), cytosol
(total cellular soluble proteins from cytoplasm), membrane/particulate
(total cellular membrane proteins including cellular organelles and
organelles membrane proteins), and cytoskeleton + DNA (total cellular
insoluble proteins, genomic DNA). In addition, the ruthenium content
in each fraction was determined by ICP-MS.As shown in Figure , ruthenium from
Ru–tpm complexes was detected primarily on the membrane/particulate
fraction, so approximately 92–98% of total intracellular Ru
was associated with this fraction. This means that the tested complexes
preferentially localize in the membrane of organelles like mitochondria
or endoplasmic reticulum. However, a small but not insignificant portion
of Ru was also associated with nuclear and genomic DNA-containing
fractions (2–8%), and the amount of Ru in these fractions increased
with incubation time. It has been shown that in the case of cisplatin,
whose anticancer mode of action is accepted as mediated by its interaction
with DNA,[76] only ca. 2–3% of intracellular
platinum reaches the nucleus and binds to DNA.[77,78] Thus, although the amount of ruthenium in fractions containing nuclear
components (DNA and proteins) is minor, the mechanism of antiproliferative
activity via interaction and chromatin damage cannot be excluded.
Figure 4
Relative
distribution of selected Ru complexes in subcellular fractions
after 5 and 24 h treatment of HCT116 cells with equimolar concentrations
(10 μM) of 1, 7, and 8. The sum of Ru concentration in all sample compartments was taken
as 100%. The experiment was performed in duplicate. Data represents
mean ± SEM from two measurements.
Relative
distribution of selected Ru complexes in subcellular fractions
after 5 and 24 h treatment of HCT116 cells with equimolar concentrations
(10 μM) of 1, 7, and 8. The sum of Ru concentration in all sample compartments was taken
as 100%. The experiment was performed in duplicate. Data represents
mean ± SEM from two measurements.Additionally, many Ru complexes from the literature
have been shown
to act via DNA-damaging mechanisms.[79] The
complexes studied in this work contain one chlorido ligand, which
is prone to substitution by water in a low-chloride environment such
as the intracellular one, thus possibly favoring the DNA binding.
Despite not collecting relevant evidence from the stability studies
(see Section 2), the additional potential role of κ3 to κ2 switching of tpm coordination should not
be ruled out in principle (see the Introduction). Next, experiments were performed to investigate the possibility
of DNA-damaging potency in the overall biological activity of the
three leading complexes.One of the experimental criteria applied
to prove DNA binding to
be responsible for the cytotoxicity of metal-based drugs is based
on the observation that the drug exhibits higher toxicity in the cells
deficient in DNA repair.[80,81] The reason lies in
the fact that the ability of DNA lesions to induce cell death is inversely
dependent on the capacity of the cells to repair the damage. Therefore,
an experiment focused on the effect of the studied complexes in wild-type
Chinese hamster ovary cell line CHO-K1 and its mutant NER-deficient
counterpart MMC-2 was performed to clarify the involvement of nuclear
DNA damage in mediating cellular sensitivity to 1, 7, and 8.As shown in Table S3, the MMC-2 cells
were significantly more sensitive to the treatment with cisplatin
(IC50 values being almost ten times lower than the IC50 values found for parental cells CHO-K1). This result confirms
that unrepaired DNA damage caused by cisplatin contributes markedly
to its antiproliferative activity, in agreement with the DNA-damage-mediated
mechanism of action of cisplatin. However, the IC50 values
found for selected ruthenium complexes were nearly the same for both
MMC-2 and parental CHO-K1 cells, indicating that NER-reparable DNA
lesions do not play a significant role in the mechanism of activity
of Ru–tpm complexes. Notwithstanding the above results, it
should also be noted that cytoskeletal proteins are also present in
the DNA-containing fraction. Therefore, to evaluate the possible effect
of Ru–tpm complexes on the cytoskeleton, the morphology of
cytoskeletal polymers, such as actin and tubulin filaments that participate
in many vital cell functions, including division, morphogenesis, phagocytosis,
and motility, was monitored by confocal microscopy. As shown in Figures S54 and S55, incubation with complexes 1, 7, or 8 did not significantly
affect the structure, shape, and layout of either tubulin (Figure S54) or actin (Figure S55) networks, even at concentrations causing a significant
antiproliferative effect (IC50). Thus, the antiproliferative
activity of the investigated complexes seems to be unlikely related
to the damage of the two major components of the cellular cytoskeleton,
i.e., actin or tubulin filaments.
Real-Time Cell Growth Monitoring
The results described
above reveal DNA damage as an unlikely cause of the biological action
of the studied Ru complexes. Therefore, real-time impedance monitoring
of cellular responses was used further to elucidate their mechanism
of action. It has been shown that bioactive compounds produce specific
time-dependent cell response profiles (TCRPs), predictive of the mechanism
of action of the investigated molecules.[82−84] Furthermore,
a comparison of the shape characteristics of the TCRPs obtained for 1, 7, and 8 with those published
for classes of compounds acting through various mechanisms[82] revealed that the TRCPs of these three complexes
(Figure ) significantly
differ from those characteristic for DNA-damaging agents. This result
further supports the view that DNA is not a major target of the Ru–tpm
complexes tested in this work.
Figure 5
Interaction of HCT116 cells with 1, 7, and 8 at their 40, 160, and
160 μM concentrations,
respectively, monitored by a real-time cell analyzer (RTCA). The vertical
dashed lines indicate the start of the treatment after allowing the
cells to adhere to microelectrodes and grow for 24 h. Cell indices
were normalized to account for differences in cell counts across the
wells prior to the treatment.
Interaction of HCT116 cells with 1, 7, and 8 at their 40, 160, and
160 μM concentrations,
respectively, monitored by a real-time cell analyzer (RTCA). The vertical
dashed lines indicate the start of the treatment after allowing the
cells to adhere to microelectrodes and grow for 24 h. Cell indices
were normalized to account for differences in cell counts across the
wells prior to the treatment.Interestingly, TRCPs obtained for 1, 7, and 8 resemble the profile of deoxycycline,[82] a known inhibitor of mitochondria in cancer
cells. This inhibitor has been shown to impair mitochondrial function
by reducing mitochondrial membrane potential and mitochondrial respiration.[85] Thus, the result of this experiment suggested
that the mechanism of action of Ru–tpm complexes could be related
to the impairment of mitochondria. Moreover, the majority of the intracellular
Ru from 1, 7, and 8 was localized
in the fraction comprising cellular organelles, including mitochondria
(Figure ). In isolated
mitochondria, 467 ± 66, 127 ± 24, and 70 ± 15 pg Ru/106 cells were found when the cells were treated with 1, 7, and 8 respectively, indicating that
Ru–tpm complexes accumulate in these organelles, although not
exclusively. These facts support the hypothesis that the mitochondria
may represent one of the significant targets of biological action
of the tested complexes. Therefore, several functional assays were
used to confirm whether and how the Ru–tpm complexes affect
mitochondria in HCT116 cells.
Effect on Mitochondrial Membrane Potential
To provide
evidence for the hypothesis that Ru–tris-pyrazolylmethane complexes
tested in this work promote tumor cell death by a mitochondria-dependent
mechanism, changes in the mitochondrial membrane potential of HCT116
cells after treatment with 1, 7, and 8 were determined by the TMRE assay. TMRE (ethyl ester of
tetramethylrhodamine) is a cell-permeant fluorescent dye that accumulates
in negatively charged mitochondria in a charge-dependent manner and
is therefore used as a marker of mitochondrial membrane potential.
If mitochondria depolarize or lose their integrity, the TMRE fluorescence
intensity decreases accordingly.After 5 h treatment, the HCT116
cells were stained with TMRE, and fluorescence changes reflecting
mitochondrial membrane depolarization in response to the ruthenium
complexes were observed by flow cytometry. Quantitative analysis (Figures and S56) revealed a significant decrease of TMRE
fluorescence in treated cells compared with control (untreated) cells,
which indicated a marked mitochondrial depolarization in HCT116 cells
due to the action of ruthenium complexes. The effect was concentration-dependent
and more pronounced for 1 and 8. Instead, 7 was less potent in reducing mitochondrial membrane potential,
but its effect was still significant.
Figure 6
Bar graph showing median of TMRE intensity
normalized to the intensity
of the control. Cells were treated with equimolar and equitoxic concentrations
of Ru complexes. Data represent mean ± SEM from three independent
experiments.
Bar graph showing median of TMRE intensity
normalized to the intensity
of the control. Cells were treated with equimolar and equitoxic concentrations
of Ru complexes. Data represent mean ± SEM from three independent
experiments.
Effect on Oxidative Phosphorylation
Mitochondria are
key players in cellular bioenergetics, producing the majority of ATP
by oxidative phosphorylation (OXPHOS). Therefore, the next experiment
was performed to reveal whether the activity of the Ru complexes toward
mitochondria also comprises respiratory chain uncoupling and inhibition
of ATP syntheses. Mitochondrial Tox-Glo assay quantifies fluorescence
signal originating from cell membrane permeabilization (nonspecific
cell death) and luminescence signal generated by luciferase in the
presence of ATP. Thus, it can distinguish primary mitochondrial disfunction
from secondary cytotoxic events. Cells are treated and grown in a
glucose-containing or glucose-free (galactose-supplemented) medium.
In the presence of glucose, the cells may preferentially rely on glycolysis
to meet bioenergetics needs and are relatively unresponsive to mitochondrial
toxins. However, under glucose-free conditions (e.g., in the presence
of galactose), the cells necessarily use OXPHOS to generate ATP and
are more responsive to mitochondrial perturbation. If the drug disrupts
oxidative phosphorylation, then a decrease in the ATP signal becomes
observable.As demonstrated in Figure , well-known OXPHOS inhibitor Antimycin A
(used as a positive control) significantly decreased ATP production
by cells growing in galactose-supplemented media. Simultaneously,
the viability of cells was unaffected, as demonstrated by no changes
in fluorescence signals (Figure ). This indicates that a decrease in ATP synthesis
results from OXPHOS inhibition rather than the overall cytotoxic effect.
In contrast, no effect on ATP synthesis was observed for the investigated
Ru complexes, neither in glucose-containing nor glucose-free medium.
A decrease in ATP production was noticed only at the highest concentrations
of complexes 1 and 8; however, it was accompanied
by a decrease in viability. Thus, reducing ATP levels under these
conditions likely resulted from the cytotoxic effect instead of OXPHOS
inhibition. In summary, the primary mechanism of action of Ru–tris-pyrazolylmethane
complexes does not consist in inhibition of OXPHOS; in other words,
these compounds do not behave as specific mitotoxicants from the point
of view of OXPHOS.
Figure 7
Graphs showing the fluorescence signal intensity (in black,
cell
viability) and luminescence (in red, ATP) normalized to the untreated
control. Cells grown in a glucose-containing medium are marked with
a rectangle, while cells grown in a glucose-free medium are marked
with a circle. Antimycin A was used as a positive control of OXPHOS
inhibition.
Graphs showing the fluorescence signal intensity (in black,
cell
viability) and luminescence (in red, ATP) normalized to the untreated
control. Cells grown in a glucose-containing medium are marked with
a rectangle, while cells grown in a glucose-free medium are marked
with a circle. Antimycin A was used as a positive control of OXPHOS
inhibition.
Effect on Calcium Homeostasis
The previous results
revealed that the investigated ruthenium complexes, although affecting
mitochondria, do not uncouple mitochondrial energy metabolism. Thus,
another mechanism underlying the effect of the investigated Ru complexes
on mitochondria must be in play. Until now, multiple mechanisms of
mitochondrial toxicity have been reported besides OXPHOS inhibition.[86,87] Mitochondria are important in intracellular signal transduction
and tuning of calcium (Ca2+) homeostasis. In stress conditions,
intracellular Ca2+ is often elevated, and functional mitochondria
serve as a potent Ca2+-buffer system.[88] Much of the function of the mitochondria relies on Ca2+ homeostasis and effective Ca2+ signaling. Calcium
regulates mitochondrial dynamics and functionality, thus affecting
various cellular processes, including the mitochondrial pathway, enzyme
activity, etc. Then, the fine modulation of mitochondrial calcium
homeostasis plays a fundamental role in many processes involving these
organelles.[89] Cancer cells have therefore
evolved mechanisms to modulate mitochondrial Ca2+ transport
in order to sustain their metabolic demand and ensure their survival.[90] The complex role of mitochondrial calcium in
cancer has been thoroughly summarized in several recent reviews.[90−92] Therefore, we decided to assess the effect of selected complexes
on Ca2+ homeostasis using cytoplasmic and mitochondrial
Ca2+ fluorescent sensors. As demonstrated in Figure A, complex 1 reduced
Ca2+ flux into the mitochondria induced by ionomycin, a
potent ionophore commonly used for this assay. Similarly, a decrease
in Ca2+ flux into the mitochondria was also observed for
complexes 7, 8, and the less active 9 (Figure C), although their effects were less pronounced (7 and 8) or insignificant (9), in agreement with their
noticeably lower activity (Table ). This effect was accompanied by an increase in the
concentration of Ca2+ in the cytoplasm (Figure B,D) and was most prominent
for complex 1, consistently with its greatest antiproliferative
activity. The elevation of cytoplasmic calcium can be related to the
inhibition of mitochondrial Ca2+ intake, as cytoplasmic
calcium cannot be transferred to the mitochondria. However, it cannot
be ruled out that this increase could also be related to other concurrent
factors, such as a release of Ca2+ from the endoplasmic
reticulum or an effect on Ca2+ channels in the cytoplasmic
membrane. Thus, the tested complexes behave similarly to other well-known
mitochondrial Ca2+ uptake inhibitors.[93−97] The enhanced cytoplasmic concentration of Ca2+ ions can then lead to apoptosis via calcineurin-mediated
proapoptotic protein activation,[98] calpain
proteases activation,[99] or autophagy by
mTOR inhibition.[100] These data imply that
the mechanism of biological action of the new complexes may also be
mediated by disruption of Ca2+ homeostasis. On the other
hand, in addition to disrupting Ca2+ homeostasis, other
mechanisms may be involved in the biological effects of the here investigated
compounds.
Figure 8
(A, C) Effect on the mitochondrial influx of calcium. Cells pretreated
for 2.5 h with 2.5, 5, or 10 μM Ru complex 1 (panel
A) or with 10 μM 1, 7, 8, or 9 (panel C) were stained with 5 μM Rhod-2
(mitochondrial Ca2+ indicator) and treated with 5 μM
ionomycin. (B, D) Effect on the cytosolic concentration of Ca2+. Cells were treated for 2.5 h with 2.5, 5, or 10 μM
Ru complex 1 (panel B) or with 10 μM 1, 7, 8, or 9 (panel D) and
subsequently stained with 5 μM Fluo-4 (cytoplasmic Ca2+ indicator). In both experiments, the fluorescence intensity was
recorded in PBS with 2 mM CaCl2.
(A, C) Effect on the mitochondrial influx of calcium. Cells pretreated
for 2.5 h with 2.5, 5, or 10 μM Ru complex 1 (panel
A) or with 10 μM 1, 7, 8, or 9 (panel C) were stained with 5 μM Rhod-2
(mitochondrial Ca2+ indicator) and treated with 5 μM
ionomycin. (B, D) Effect on the cytosolic concentration of Ca2+. Cells were treated for 2.5 h with 2.5, 5, or 10 μM
Ru complex 1 (panel B) or with 10 μM 1, 7, 8, or 9 (panel D) and
subsequently stained with 5 μM Fluo-4 (cytoplasmic Ca2+ indicator). In both experiments, the fluorescence intensity was
recorded in PBS with 2 mM CaCl2.
Cytotoxicity in Three-Dimensional (3D) Spheroids
To
emphasize the possibility that the tested complexes might be promising
candidates for further preclinical testing and to improve the relevance
of our in vitro results, we used 3D cell cultures
(spheroids), which are much better at replicating in vivo environment
than traditional two-dimensional (2D) cultures. Cells grow in complex
3D cultures in an environment closely reflecting the tumor microenvironment,
such as nutrient and oxygen gradients, intercellular and cell-extracellular
matrix interactions, and heterogeneity.[101] Hence, 3D growth of immortalized established cell lines or primary
cell cultures is regarded as a more stringent and representative model
for performing in vitro drug screening.[101−103] Moreover, 3D cell culture models using human cells can circumvent
the drawbacks of animal models that, aside from the high cost and
ethical considerations, cannot always recapitulate human diseases
or capture the side effect of drugs accurately. Also, the tumor spheroids
have exhibited several features of the in vivo solid
tumors. The similarities in the drug responsiveness among the tumor
spheroids and the animal models might largely be due to their similarities
in enhanced cellular interactions via adhesion and secretion of soluble
factors of tumors that lead to low pH and hypoxia.[104]Therefore, we assessed the ability of 1, 7, and 8 to inhibit spheroid formation
and growth. HCT116 cells were cultured under 3D cell culture conditions
for 96 h to grow up to a tissue mass of around 100 μm in diameter,
as described in Experimental Section. The
spheroids were then treated with various concentrations of complexes
for an additional 72 h. The Cell Titer-Glo 3D cell viability assay
was used to determine IC50 values. The results (Table ) confirm that Ru–tpm
complexes tested in this experiment, which showed very good activity
in conventional 2D cell cultures, also exhibit substantial activity
in the 3D spheroids formed from HCT116 cells being significantly more
effective than clinically used cisplatin. According to what is observed
on 2D cultures, 1 excels in its activity, being approximately
18-fold more effective than cisplatin.
Table 5
IC50 Values (μM)
for the HCT116 Cell Line Determined by the Cell Titer-Glo Test after
72 h of Treatment
compound
HCT116 (3D)
1
2.5 ± 0.6
7
12.2 ± 2.5
8
12.1 ± 2.7
cisplatin
44.6 ± 1.8
The effect on the morphology of 3D spheroids of HCT116
cells is
shown in Figure S57. The spheroids formed
by the control, untreated cells displayed round-shape morphology with
a well-defined surrounding edge (Figure S57, panel A). However, after treatment with 7, 8, or cisplatin (Figure S57, panels C–E),
spheroids displayed heterogeneous morphology with several dissociated
cell clumps. Furthermore, these dissociated cell clumps were found
to a much greater extent in the samples treated with 1; under the condition of the experiment, the spheroids treated with
this complex were almost completely disintegrated (Figure S57, panel B). Thus, also this morphologic study showed
superior activity of Ru–tris-pyrazolylmethane complexes in
the 3D spheroids formed from colon cancer HCT116 cells.
Conclusions
The search for metal drugs alternative
to platinum compounds, which
are currently administered in clinical treatment against several types
of tumors, is an ultimate demand of research. Ruthenium complexes
have aroused a great interest in this regard due to their versatile
anticancer activity and the scarce toxicity of the metal element,
and some ruthenium(II)–arene compounds (RAPTA complexes) are
pointing to clinical trials with great promise. Tris-pyrazolylmethane
(tpm) is a versatile ligand behaving as a neutral six-electron donor
like the arene moiety, but parallel studies on the anticancer potential
of Ru–tpm species are lacking in the literature, apparently
due to some synthetic drawbacks. Here, we report a straightforward
route to access a family of novel, robust cationic ruthenium(II)–tpm
complexes differing from each other in one key ligand, modulating
both the amphiphilic character and the strength of the ruthenium–chloride
bond, which may be implicated in the activation mechanism.Three
complexes of the series, showing a favorable combination
of these two factors, displayed micromolar potency against a panel
of human cancer cells (comparable to that of conventional cisplatin).
Notably, the promising potency of these leading Ru–tpm complexes
was bolstered by the results obtained with the 3D spheroids formed
from cancer cells, which are much better at replicating in vivo environments
than traditional 2D cultures. Moreover, these complexes demonstrated
selectivity toward cancer over noncancerous cells. Our further data
prove that Ru–tpm complexes effectively induced apoptosis in
cancer cells, whereas the percentage of cells undergoing necrosis
was negligible. The complexes are taken up in large amounts by cancer
cells, and a correlation was observed between antiproliferative activity
and intracellular accumulation. In vitro growth inhibition
studies were completed by investigating the mechanism by which the
studied complexes inhibit the growth of cancer cells. The results
of these experiments showed that the mitochondria represent a significant
target. Although mitochondria are key players in cellular bioenergetics,
producing the majority of ATP by oxidative phosphorylation (OXPHOS),
the primary mechanism of action of Ru–tpm complexes does not
consist of inhibition of OXPHOS. In contrast, we conclude, based on
the present findings, that the biological action is mediated by disruption
of calcium homeostasis due to the inhibiting of mitochondrial calcium
intake. To the best of our knowledge, this is the first study demonstrating
a mechanism of antiproliferative activity of ruthenium complexes in
cancer cells that involves the regulation of mitochondrial calcium
homeostasis. However, the regulation of mitochondrial calcium homeostasis
may not be the only mechanism by which the present complexes act.
Ruthenium complexes are generally considered to be multifactorial
agents, so the biological activity of Ru–tpm complexes may
include other factors whose identification was beyond the scope of
this work. Nevertheless, disruption of calcium homeostasis undoubtedly
contributes to the overall activity of these complexes. Work is currently
underway to evaluate other possible mechanistic contributions to the
overall biological activity and the in vivo tumor efficacy of this
new class of antitumor metal complexes.
Experimental Section
General Remarks
Reactants and solvents were purchased
from Alfa Aesar, Merck, Strem, or TCI Chemicals and were of the highest
purity available. Tris(1-pyrazolyl)methane (tpm) was prepared according
to the published procedure.[105] Reactions
were conducted under a N2 atmosphere using standard Schlenk
techniques, and all products were stored in air once isolated. All
compounds are >95% pure by elemental analysis. Solvents were used
as received unless otherwise stated. Toluene and diethyl ether were
dried with the solvent purification system mBraun MB SPS5, while methanol
was distilled from calcium hydride and isopropanol from magnesium.
IR spectra of solid samples were recorded on an Agilent Cary630 FTIR
spectrometer. IR spectra were processed with Spectragryph software.[106] NMR spectra were recorded at 298 K on a JEOL
JNM-ECZ500R instrument equipped with a Royal HFX Broadband probe.
Chemical shifts (expressed in parts per million) are referenced to
the residual solvent peaks (1H, 13C)[107] or external standard (31P to H3PO4). 1H and 13C{1H} NMR spectra were assigned with the assistance of 1H–13C (gs-HSQC and gs-HMBC) correlation experiments.[108] Elemental analyses were performed on a Vario
MICRO cube instrument (Elementar). Conductivity measurements were
performed at 25 °C using an XS COND 8 instrument (cell constant
= 1.0 cm–1).[109,110] pH measurements were
performed with an Orion pH meter equipped with a Hamilton glass pH
electrode. ESI-Q/ToF flow injection analysis was carried out using
a 1200 Infinity HPLC (Agilent Technologies), coupled to a Jet Stream
ESI interface (Agilent) with a Quadrupole-Time-of-Flight tandem mass
spectrometer 6530 Infinity Q-TOF (Agilent Technologies).
Synthesis and Characterization of Complexes
[RuCl(κ3-tpm)(PPh3)2]Cl, 1 (Chart )
The title compound was prepared using a slightly modified literature
procedure (Chart ).[38,39]
Chart 1
Synthesis and Structure
of 1 (Labeling Refers to Carbon
Atoms)
Step (a): Synthesis of [RuCl2(PPh3)3]
A solution of RuCl3·3H2O (350 mg, 1.34 mmol) and triphenylphosphine (1.75 g, 6.70 mmol)
in distilled methanol (20 mL) was heated at reflux for 4 h. Thus,
complex [RuCl2(PPh3)3] precipitated
as a brown-purple solid. This solid was separated by filtration under
a N2 atmosphere, washed with methanol and three times with
diethyl ether, and finally dried under vacuum for several hours. The
isolated material contained minor amounts of ineliminable PPh3 and O=PPh3, according to 31P
NMR (resonances at −5.5 and 29.2 ppm, respectively), which
did not affect step (b).
Step (b): Synthesis of [RuCl2(κ3-tpm)(PPh3)2]
Complex [RuCl2(PPh3)3], as obtained from step (a), and an
equimolar amount of tpm (286 mg, 1.34 mmol) were taken in 30 mL of
anhydrous toluene under vigorous agitation. A yellow solid rapidly
precipitated, and the mixture was left to stir overnight. The solid
was filtered, washed with toluene and three times with diethyl ether,
and finally dried under vacuum. Yellow solid, yield 1.13 g (95%).
Anal. calcd for C46H40Cl2N6P2Ru: C, 60.66; H, 4.43; N, 9.23; Cl, 7.79. Found: C,
60.35; H, 4.47; N, 9.13; Cl, 7.90. IR (solid state): ṽ/cm–1 = 3138w, 3121w, 3101w, 3061w, 3054w, 1515w, 1480m,
1467m, 1454m, 1440m, 1433m, 1411m, 1404m, 1383w, 1303m, 1291m, 1259m,
1254m, 1222m, 1187m, 1090s, 1075m, 1048m, 1024w, 998w, 988w, 852m,
799m, 776m, 768m, 754s, 742s, 696s, 691s, 681m. 1H NMR
(CDCl3): δ/ppm = 12.21 (s, 1H, CδH); 8.86 (d, 1H, 3JHH = 2.7
Hz, CγH trans to P); 8.64 (d, 2H, 3JHH = 2.8 Hz, CγH trans to Cl);
7.29–7.26 (m, 3H, C4H); 7.15–7.07 (m, 12H,
C3H + C2H); 6.83 (d, 2H, 3JHH = 2.2 Hz, CαH trans to P);
5.98 (t, 2H, 3JHH = 2.6 Hz,
CβH trans to P); 5.50 (t, 1H, 3JHH = 2.7 Hz, CβH trans to Cl);
5.12 (d, 1H, 3JHH = 2.4 Hz,
CαH trans to Cl). 31P{1H} NMR
(CDCl3): δ/ppm = 40.1[38]
Preparation of [RuCl(κ3-tpm)(PPh3)(NCMe)]Cl, 2 (Chart )
A solution of 1 (200 mg, 0.22
mmol) in 25 mL of acetonitrile (MeCN) was heated at reflux for 3 h.
The solvent was evaporated under reduced pressure, and the obtained
solid was washed with diethyl ether and dried under vacuum. Yellow
solid, yield 144 mg (95%). Anal. calcd for C30H28Cl2N7PRu: C, 52.26; H, 4.09; N, 14.22; Cl,
10.28. Found: C, 52.08; H, 3.98; N, 14.26; Cl, 10.40. IR (solid state):
ṽ/cm–1 = 3109w, 3055w, 2958w, 2919w, 2278w
(ṽN=C), 1620w-br, 1507w, 1483w, 1450w, 1433m, 1408m,
1375w, 1289m, 1277w, 1252w, 1223w, 1187w, 1090s, 1053w, 1048w, 997w,
987w, 857w, 971s, 779s, 767s, 750s, 756s, 695s. 1H NMR
(CDCl3): δ/ppm = 12.29 (s, 1H, CδH); 8.90, 8.75, 8.71 (d, 3H, 3JHH = 2.9 Hz, CγH); 8.14 (d-br, 1H, CαH); 6.91, 6.55 (d, 2H, 3JHH = 2.2 Hz, CαH); 7.41–7.27 (m, 15H, C3H + C4H + C6H); 6.43 (s-br, 1H CβH); 6.07, 5.96 (t, 2H, 3JHH = 2.6 Hz, CβH); 2.16 (s, 3H, C2H). 13C{1H} NMR (CDCl3):
δ/ppm = 148.0, 147.2, 144.1 (Cα); 135.6, 135.4,
133.6 (Cγ); 134.1 (C4, 3JPC = 9.4 Hz); 132.3 (C3, 1JPC = 42.3 Hz); 130.1 (C6);
128.3 (C5, 4JPC =
9.3 Hz); 127.9 (C1); 108.3, 108.2, 108.0 (Cβ); 74.4 (Cδ); 4.7 (C2). 31P{1H} NMR (CDCl3): δ/ppm = 48.1 (Chart ).
Chart 2
Structure of 2 (Labeling Refers to Carbon
Atoms)
General Procedure for the Synthesis of Complexes [RuCl(κ3-tpm)(PPh3)(L)]Cl
A solution of 1 and the proper ligand L in ethanol or anhydrous isopropanol
was heated at reflux for a variable time. After cooling to room temperature,
the volatiles were evaporated under reduced pressure. The crude product
was redissolved in the minimum volume of dichloromethane, precipitated
with diethyl ether, filtered, and dried under vacuum.
Crystal data and collection details
for 3·2CHOH, 4·0.5EtO·solv, 5·CHCl·2HO, 6·solv, 7·CHCOCH·HO, and 8·2HO are reported
in Table . Data were
recorded on a Bruker APEX II diffractometer equipped with a PHOTON2
detector using Mo Kα radiation. The structures were solved by
direct methods and refined by full-matrix least-squares based on all
data using F2.[111] Hydrogen atoms were fixed at calculated positions and refined using
a riding model.[112] The crystals of 4·0.5EtO·solv and 6·solv contain some highly disordered solvent
molecules that have been treated using the SQUEEZE routine of PLATON.[113,114] The refined crystal structures of 4·0.5EtO·solv and 5·CHCl·2HO contain
large difference peaks; these are located in the proximity of disordered
chloride ions or CH2Cl2 molecules.
Table 6
Crystal Data and Measurement Details
for 3·2CH, 4·0.5Et, 5·CH, 6·solv, 7·CH, and 8·2H
3·2CH3OH
4·0.5Et2O·solv
5·CH2Cl2·2H2O
6·solv
7·CH3COCH3·H2O
8·2H2O
formula
C34H39Cl2N8O2PRu
C37H36Cl2N8O0.5PRu
C32H35Cl4N8O2PRu
C33H30Cl2N7OPRu
C34H42Cl2N6O5P2Ru
C35H40Cl2N7O2PRu
FW
794.67
803.68
837.52
743.58
848.64
793.68
T, K
100(2)
100(2)
100(2)
100(2)
100(2)
100(2)
λ, Å
0.71073
0.71073
0.71073
0.71073
0.71073
0.71073
crystal system
triclinic
monoclinic
triclinic
triclinic
monoclinic
monoclinic
space group
P1̅
P21/c
P1̅
P1̅1̅
C2/c
P21/c
a, Å
10.1428(8)
20.5145(12)
9.7016(8)
11.1525(4)
41.370(8)
17.110(2)
b, Å
13.2743(11)
20.6593(12)
10.0588(8)
12.4862(5)
9.756(2)
10.1025(13)
c, Å
13.9235(11)
19.0486(11)
19.2103(16)
14.7685(6)
19.152(4)
21.524(3)
α, deg
72.776(2)
90
97.728(2)
87.2220(10)
90
90
β, deg
77.299(2)
100.126(2)
103.181(3)
73.5550(10)
91.34(3)
106.679(8)
γ, deg
85.255(3)
90
97.790(2)
89.3910(10)
90
90
cell volume, Å3
1746.4(2)
7947.3(8)
1781.7(3)
1970.09(13)
7728(3)
3564.1(8)
Z
2
8
2
2
8
4
Dc, g·cm–3
1.511
1.343
1.561
1.253
1.459
1.479
μ, mm–1
0.692
0.607
0.828
0.606
0.674
0.677
F(000)
816
3288
852
756
2488
1632
crystal size, mm
0.16 × 0.13 × 0.10
0.22 × 0.16 × 0.14
0.22 × 0.18 × 0.15
0.18 × 0.15 × 0.12
0.24 × 0.21 × 0.18
0.18 × 0.13 × 0.10
θ limits, deg
1.5649–26.998
1.467–25.998
2.074–25.997
1.633–27.227
1.970–26.000
1.9759–26.998
reflections collected
32 233
142 792
29 925
27 297
54 659
72 769
independent reflections
75 857 [Rint = 0.0850]
15 547 [Rint = 0.1422]
6987 [Rint = 0.0724]
8722 [Rint = 0.0465]
7594 [Rint = 0.0553]
7780 [Rint = 0.0673]
data/restraints/parameters
7585/17/428
15 547/8/871
6987/39/445
8722/0/407
7594/130/473
7780/6/445
goodness on fit on F2 a
1.193
1.075
1.036
1.097
1.038
1.064
R1 (I > 2σ(I))b
0.0628
0.0799
0.0610
0.0486
0.0336
0.0308
wR2 (all data)c
0.1366
0.1881
0.1604
0.1053
0.0772
0.0767
largest diff. peak and hole, e Å–3
1.972/–1.232
3.653/–1.407
3.443/–2.006
1.132/–0.668
1.028/–0.793
0.651/–0.568
Goodness on fit on F2 = [Σw(FO2 – FC2)2/(Nref – Nparam)]1/2, where w = 1/[σ2(FO2) + (aP)2 + bP], where P = (FO2 + 2FC2)/3; Nref = number of reflections used in the refinement; Nparam = number of refined parameters.
R1 =
Σ||FO| – |FC||/Σ|FO|.
wR2 =
[Σw(FO2 – FC2)2/Σw(FO2)2]1/2, where w = 1/[σ2(FO2) + (aP)2 + bP], where P =
(FO2 + 2FC2)/3.
Goodness on fit on F2 = [Σw(FO2 – FC2)2/(Nref – Nparam)]1/2, where w = 1/[σ2(FO2) + (aP)2 + bP], where P = (FO2 + 2FC2)/3; Nref = number of reflections used in the refinement; Nparam = number of refined parameters.R1 =
Σ||FO| – |FC||/Σ|FO|.wR2 =
[Σw(FO2 – FC2)2/Σw(FO2)2]1/2, where w = 1/[σ2(FO2) + (aP)2 + bP], where P =
(FO2 + 2FC2)/3.
Behavior in Aqueous Media
(a) Solubility in Water
A suspension of the selected
ruthenium complex (3–5 mg) in a D2O solution (0.7
mL) containing Me2SO2 as the internal standard[115] (3.36 × 10–3 M) was
vigorously stirred at 21 °C for 2 h. The resulting saturated
solution was filtered over celite, transferred into an NMR tube, and
analyzed by 1H NMR and 31P NMR spectroscopy
(delay time = 3 s; number of scans = 20). The concentration (solubility)
was calculated by the relative integral (starting complex + aquo complex,
CβH signal in the 1H spectrum) with respect
to Me2SO2 (δ/ppm = 3.14). The results
are compiled in Table . NMR data are given in the Supporting Information (Figures S36–S47).
(b) Stability in D2O and DMSO-d6/D2O
The same samples prepared at point
(a) were used in this experiment, apart from complex 1 that was dissolved in DMSO-d6/D2O 4:1 v/v solution (0.7 mL; [Ru] = approx. 3 mg) containing
Me2SO2 as the standard.[115] After 1H and 31P{1H} NMR
analyses described above at point (a) (time = t0), samples were heated at 37 °C for 48 h. After cooling
to room temperature, 1H and 31P NMR spectra
were recorded. The percentage of starting complex + related aquo complex
was calculated by the relative integral (CβH signal
in the 1H spectrum) with respect to Me2SO2 (c = 3.3 × 10–3 mol·L–1; δ/ppm = 3.14 in D2O; δ/ppm
= 2.95 in DMSO-d6/D2O 4:1 v/v),
see Table and S1.
Partition coefficients (Pow); IUPAC: KD partition
constant,[116] defined as Pow = corg/caq, where corg and caq are the molar concentrations of the selected
compound in the organic and aqueous phase, respectively, were determined
by the shake-flask method and UV–vis measurements.[62,117,118] Deionized water and 1-octanol
were vigorously stirred for 24 h to enable saturation of both phases
and then separated by centrifugation. A stock solution of the selected
ruthenium compound (ca. 2 mg) was prepared by first adding MeOH (50
μL, to help solubilization), followed by octanol-saturated water
(2.5 mL). The solution was diluted with octanol-saturated water (ca.
1:3 v/v ratio, cRu ≈ 10–4 M, so that 1.5 ≤ A ≤ 2.0 at λmax), and its UV–vis spectrum was recorded (Aaq0). An aliquot of the solution (Vaq =
1.2 mL) was transferred into a test tube, and water-saturated octanol
(Vorg = Vaq = 1.2 mL) was added. The mixture was vigorously stirred for 20 min
at 21 °C then centrifuged (5000 rpm, 5 min). The UV–vis
spectrum of the aqueous phase was recorded (Aaqf), and the partition
coefficient was calculated as Pow = (Aaq0 – Aaqf)/Aaqf, where Aaq0 and Aaqf are the
absorbances in the aqueous phase before and after partition with the
organic phase, respectively.[118] For 1, an inverse procedure was followed, starting from a solution
of the compound in water-saturated octanol. The partition coefficient
was calculated as Pow = Aorgf/(Aorg0 – Aorgf), where Aorg0 and Aorgf are the absorbances
in the organic phase before and after partition with the aqueous phase,
respectively. The wavelength of the maximum absorption of each compound
(280–380 nm range) was used for UV–vis quantitation.
The procedure was repeated three times for each sample (from the same
stock solution); the results are given as mean ± standard deviation
(Table ). Naphthoquinone
was used as a reference compound (log P =
1.8 ± 0.2; literature:[119] 1.71).
(d) Stability in Cell Culture Medium
Powdered DMEM
cell culture medium (1000 mg/L glucose and l-glutamine, without
sodium bicarbonate and phenol red; D2902; Sigma-Aldrich) was dissolved
in D2O (10 mg/mL), according to the manufacturer’s
instructions. The solution of deuterated cell culture medium (“DMEM-d”) was treated with Me2SO2 (6.6 × 10–3 M) and NaH2PO4/Na2HPO4 (0.10 M, pD = 7.5)[120−122] and then stored at 4 °C under N2. The selected ruthenium compound (2–3 mg) was dissolved
in DMSO-d6 (0.14 mL; 0.18 mL for 4) and then diluted with DMEM-d up to 0.75
mL total volume (cRu ca. 4 × 10–3 M). The mixture was stirred for 30 min, then filtered
over celite, and transferred into an NMR tube. The resulting yellow
solution was analyzed by 1H and 31P NMR (delay
time = 3 s; number of scans = 20) and then heated at 37 °C for
24 h. After cooling to room temperature, NMR analyses were repeated.
Compound 1 was instead dissolved in a DMSO-d6/DMEM-d 4:1 v/v solution (0.7 mL; [Ru]
= approx. 3 × 10–3 M) containing Me2SO2 as the internal standard.[115] The percentage of the starting complex + related aquo complex was
calculated by the relative integral with respect to Me2SO2 (δ/ppm = 3.16 in DMSO-d6/DMEM-d 1:4 and 1:3 v/v; δ/ppm = 2.95
in DMSO-d6/DMEM-d 4:1
v/v).
Biological Studies
Cell Lines, Culture Conditions, and Stock Solutions of Ru Complexes
The human cervical carcinoma HeLa cells and human colorectal carcinoma
cells HCT116 were kindly supplied by Professor B. Keppler, University
of Vienna (Austria). Human rhabdomyosarcoma RD cells were purchased
from the American Type Culture Collection (ATCC, Manassas, VA). Human
breast cancer MCF-7 cells, human skin melanoma 518A2 cells, and human
MRC5pd30 cells derived from normal lung tissue were purchased from
the European Collection of Authenticated Cell Cultures (ECACC) (Salisbury,
U.K.). Chinese hamster ovary CHO-K1 cell line (wild type) and its
derivative MMC-2 carrying the ERCC3/XPB mutation (NER-deficient) cell
line were kindly supplied by Dr. M. Pirsel, Cancer Research Institute,
Slovak Academy of Sciences, Bratislava (Slovakia).All of the
cell lines were cultivated in DMEM medium (high glucose 4.5 g L–1, PAA, Pasching, Austria) supplemented with gentamycin
(50 μg mL–1, Serva, Heidelberg, Germany) and
10% heat-inactivated fetal bovine serum (PAA). The medium for MRC5pd30
cells was further supplemented by 1% nonessential amino acids (Sigma-Aldrich,
Prague, Czech Republic). All cells were cultured as adherent monolayers
in a humidified incubator at 37 °C in a 5% CO2 atmosphere
and subcultured twice a week with an appropriate plating density.For biological studies, stock solutions of Ru complexes were prepared
by dissolving the compounds in DMSO to a final concentration of 10
mM and subsequently diluted to the media to the required concentration.
The concentration of Ru in media used in the experiment was verified
by flameless atomic absorption spectrometry (FAAS). The final DMSO
concentration in the cell culture medium did not exceed 1% (v/v) to
avoid DMSO toxicity.
Antiproliferative Activity
In vitro antiproliferative activity of Ru complexes was determined by the
MTT or, alternatively, SRB assay as already described,[84] after 72 h of incubation of cells with various
concentrations of the Ru complex. The reported IC50 values
are an average of three independent experiments, each consisting of
three replicates per concentration.
Cellular Uptake
In these experiments, 1 × 106 HCT116 cells were seeded on 100 mm Petri dishes. After overnight
preincubation in a drug-free medium, the cells were treated with the
Ru complexes (15 μM) for 24 h. Afterward, the cells were extensively
washed with PBS (37 °C), detached using 0.25% trypsin, washed
twice with ice-cold PBS, and counted by an automatic cell counter.
The cell pellets were digested using a microwave acid (HCl) digestion
system (CEM Mars). The quantity of Ru taken up by the cells was determined
by inductively coupled plasma mass spectrometry (ICP-MS). All experiments
were carried out in triplicate.
Annexin-V/PI Staining
The type of cell death (apoptosis/necrosis)
caused by studied Ru complexes was determined by flow cytometry using
Annexin-V and propidium iodide staining after 24 h of treatment. HCT116
cells were seeded in a six-well plate at a density of 150000 cells/well.
After overnight cultivation, the cells were treated with studied complexes
and incubated for 24 h. Afterward, the cells from individual wells
were collected. Pellets were resuspended in the Annexin-V/PI staining
solution (BD Pharmingen), and the samples were analyzed using a BD
FACSverse flow cytometer.
Real-Time Apoptosis/Necrosis
Type of cell death and
its kinetics were measured using the Real Time-Glo Annexin-V Apoptosis
and Necrosis Assay (Promega). HCT116 cells were seeded at a density
of 8 × 103 cells/50 μL in a 96-well black plate
and incubated overnight. The cells were then treated with the Ru complexes,
and immediately afterward, kinetic analysis began. Staurosporine (10
μM) and ethanol (5%) were used as positive controls of apoptosis
and necrosis, respectively. Luminescence (integration of 1000 ms)
and fluorescence (λex: 485 nm; λem: 535 nm) were detected by a SPARK reader (Tecan, Manedorf, Switzerland).
Real-Time Cell Growth Monitoring
We employed an xCELLigence
RTCA SP Instrument (ROCHE) for monitoring cell growth in real time.
First, the background of the 96-well E-Plate was read (100 μL
of cultivation media). Then, the cells were added to the E-Plate (2000
cells/well in 50 μL of media), and the measurement was started.
After 21 h, Ru complexes were added at various concentrations, and
impedance was monitored for 72 h. An arbitrary unit CI (cell index)
is a quantitative measure in which the status of the cells (number
and morphology of attached cells) is reflected.
Cell Fractionation
The HCT116 cells were seeded at
a density of 1.5 × 106 cells/10 mL Petri dish and
incubated overnight. Then, the cells were treated with 10 μM
Ru complexes, incubated for 5 or 24 h, harvested, and exhaustively
washed with PBS. The cell pellets were processed by the FractionPREP
Cell Fractionation Kit (BioVision) according to the manufacturer’s
instructions, yielding four subcellular fractions: cytosol, membrane/particulate,
nuclear, and cytoskeletal. The Ru content in each fraction was evaluated
by ICP-MS. The measurement was performed in triplicate.
Isolation of Mitochondria
The HCT116 cells were seeded
at a density of 3 × 106 cells/10 mL Petri dish and
incubated overnight. The cells were treated with Ru complexes (10
μM) for 24 h, harvested, and washed with ice-cold PBS, and mitochondrial
fractions were extracted by the Mitochondrial Isolation Kit (MITOISO2,
Sigma-Aldrich) according to the manufacturer’s instructions.
The Ru content in each sample was evaluated by ICP-MS. The measurement
was performed in triplicate.
Effect on Mitochondrial Membrane Potential
Effects
of Ru complexes on mitochondrial membrane potential were assessed
by TMRE staining of HCT116 cells after treatment. First, HCT116 cells
were seeded on a six-well plate at a 1.5 × 105 cells/well
density. The next day, the cells were treated with equimolar (10 μM)
and equitoxic (twofold or fourfold IC50,72h) concentrations
of Ru complexes for 5 h. Then, the cells were collected and stained
with 100 nM TMRE in a complete DMEM medium for 30 min at 37 °C
in the dark. The TMRE-containing medium was then replaced with PBS,
and cells were analyzed by a BD FACSverse flow cytometer (λex = 488 nm, λem = 586 nm).
Effect on Oxidative Phosphorylation
The Mitochondrial
Tox-Glo assay was used to determine whether studied Ru complexes influence
oxidative phosphorylation in HCT116 cells. Two culture media were
prepared: glucose-containing, serum-free RPMI medium (RPMI 1640, + l-glutamine, 10 mM D-glucose (Gibco, Thermo Fisher Scientific))
supplemented with sodium bicarbonate and glucose-free, serum-free
RPMI medium (RPMI 1640, + l-glutamine, no glucose (Gibco,
Thermo Fisher Scientific)) supplemented with 10 mM galactose (Sigma)
and sodium bicarbonate. Cells were seeded at a density of 1.5 ×
103 cells/well in 50 μL in a 96-well black plate
in media containing either glucose or galactose (vide supra) and incubated
overnight. Then, the cells were treated with 50 μL of Ru complexes
in various concentrations as indicated. Cells were cultivated for
90 min at 37 °C. Cell membrane permeability and ATP quantity
were determined by the Mitochondrial Tox-Glo assay according to the
manufacturer’s protocol. The first step was to add a fluorogenic
peptide substrate (bis-AAF-R110) to measure the dead cell protease
activity. Bis-AAF-R110 substrate cannot cross the intact membrane
of live cells and therefore gives an insignificant signal with viable
cells relative to nonviable cells. The second step of the procedure
is adding an ATP detection reagent, resulting in cell lysis and generating
a luminescent signal proportional to the amount of ATP present. The
final fluorescent and luminescent signals were detected on multimode
reader SPARK (Tecan, Manedorf, Switzerland).
Calcium Flux
The distribution of calcium ions in cytosol
and mitochondria was studied using calcium-sensitive fluorescent probes.
HCT116 cells were seeded at a density of 2 × 105 cells/well
on six-well plates. After overnight cultivation, the cells were treated
with Ru complexes at indicated concentrations for 2.5 h. The culture
medium was then replaced with either 5 μM Rhod-2 or Fluo-4 in
PBS supplemented with 2 mM CaCl2 and incubated for 30 min
at 37 °C. At last, cells were harvested and suspended in PBS
(with 2 mM CaCl2) or in PBS (with 2 mM CaCl2) supplemented with ionomycin (5 μM) for 30 min at 37 °C
and analyzed using a BD FACSverse flow cytometer.
Cytotoxicity in Colonospheres
A spontaneous spheroid
formation was used to generate colonospheres derived from the HCT116
cell line. The HCT116 cells were seeded on 96-well ultralow attachment
plates (1400 cells/well) and cultivated in a tumor sphere-forming
medium (DMEM/F12, supplemented with B27 (Invitrogen), BSA, bFGF (10
ng mL–1), and EGF (20 ng mL–1))
for 4 days. After the incubation period, colonospheres were treated
with Ru complexes for another 3 days. The viability of colonospheres
was determined by the Cell Titer-Glo 3D cell viability assay (Promega)
according to the manufacturer’s instructions. The reported
IC50 values are the average of three independent experiments,
each consisting of three replicates per concentration level. Bright-field
images of the spheroids were taken as well to determine the effect
on the morphology of the spheroids.
Confocal Microscopy Imaging of Actin and Tubulin
HCT116
cells were seeded on coverslips precoated with 0.1% gelatin in six-well
culture plates at a density of 1 × 105 cells/well.
The following day, the cells were treated with the tested compounds
at concentrations corresponding to IC50 for 24 h. Following
the treatment, the cells were washed with PBS, fixed with 4% p-formaldehyde,
washed, permeabilized with 0.1% Triton X-100, and blocked. Samples
for actin staining were blocked with 1.5% BSA for 1 h and then stained
with Alexa Fluor 488-conjugated Phalloidin (Thermo Fisher Scientific,
1:50 dilution, 20 min). Samples for tubulin staining were blocked
with 5% goat serum for 1 h and incubated with primary antibody (anti-α-tubulin,
Abcam, 1:200 dilution, 1 h) and Alexa Fluor-conjugated secondary antibody
(goat antirabbit, Abcam, 1:500 dilution, 1 h). Both groups of samples
were mounted with ProLong Diamond Antifade Mountant with DAPI (Invitrogen).
Cells were visualized on a confocal microscope Leica TCS SP8 SMD.
Figure 1
Scheme 1
Figure 2
Scheme 2
Figure 3
Figure 4
Figure 5
Figure 6
Figure 7
Figure 8
Chart 1
Chart 2
Chart 3
Chart 4
Chart 5
Chart 6
Chart 7
Chart 8
Chart 9