Structural determinants of p53-independence in anticancer ruthenium-arene Schiff-base complexes

Article abstract of DOI:10.1021/acs.molpharmaceut.6b00348

p53 is a key tumor suppressor gene involved in key cellular processes and implicated in cancer therapy. However, it is inactivated in more than 50% of all cancers due to mutation or overexpression of its negative regulators. This leads to drug resistance

Full text of DOI:10.1021/acs.molpharmaceut.6b00348

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Article
Structural Determinants of p53-Independence in
Anticancer Ruthenium-Arene Schiff-base Complexes
Mun Juinn Chow, Maria V. Babak, Daniel Yuan Qiang Wong,
Giorgia Pastorin, Christian Gaiddon, and Wee Han Ang
Mol. Pharmaceutics, Just Accepted Manuscript • DOI: 10.1021/acs.molpharmaceut.6b00348 • Publication Date (Web): 13 May 2016
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Structural Determinants of p53-Independence in Anticancer Ruthenium-
Arene Schiff-base Complexes
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Mun Juinn Chow1, 2, Maria V. Babak1, Daniel Yuan Qiang Wong1, Giorgia Pastorin2, 3, Christian
Gaiddon*4, 5 and Wee Han Ang*,1, 2
1Department of Chemistry, National University of Singapore, 3 Science Drive 3, 117543 Singapore
2NUS Graduate School for Integrative Sciences and Engineering
3Department of Pharmacy, National University of Singapore, 18 Science Drive 4, 117543 Singapore
4U1113 INSERM, 3 Avenue Molière, Strasbourg 67200, France
5Oncology section, FMTS, Université de Strasbourg, Strasbourg, France
KEYWORDS
Ruthenium Arene Schiff-Base Complexes, Anticancer, Structure-Activity Relationship Studies,
p53-independent activity.
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ABSTRACT
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p53 is a key tumor supressor gene involved in key cellular processes and implicated in cancer
therapy. However, it is inactivated in more than 50% of all cancers due to mutation or
overexpression of its negative regulators. This leads to drug resistance and poor
chemotherapeutic outcome as most clinical drugs act via a p53-dependent mechanism of action.
An attractive strategy to circumvent this resistance would be to identify new anticancer drugs
that act via p53-independent mode-of-action. In the present study, we identified 9 Ru (II)-Arene
Schiff-base (RAS) complexes able to induce p53-independent cytotoxicity and discuss structural
features that are required for their p53-independent activity. Increasing hydrophobicity led to an
increase in cellular accumulation in cells with a corresponding increase in efficacy. We further
showed that all 9 complexes demonstrated p53-independent activity. This was despite significant
differences in their physicochemical properties suggesting that the iminoquinoline ligand, a
common structural feature for all the complexes, is required for the p53-independent activity.
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INTRODUCTION
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p53 is a key tumor supressor gene involved in the mediation of cellular DNA repair, cell cycle
arrest, apoptosis-induction and neurotoxicity of anticancer drugs.1-2 It is inactivated in more than
50% of all cancers due to mutation or overexpression of p53 negative regulators.3 As a result,
these cancers are more resistant to drugs that act via p53-dependent pathways. For instance, p53-
dependent drugs such as oxaliplatin (OXP) or doxorubicin (DOX) are less effective in inducing
apoptosis in these cancer types even after causing significant DNA damage.4-5 Various strategies
have been explored for more effective treatment of cancers with defective p53 function. One
such strategy involves the resensitization of these cancers to chemotherapy by restoring p53
activity. This could be done through (i) the inhibition of the interaction between p53 and its
negative regulators or (ii) restoring misfolded mutant p53, using targeted small molecules or
selective peptide sequences.6 However, such treatment is often limited by the off-target effects,
low efficacy or development of resistance towards these ‘p53-activators’.6-7 A more attractive
strategy would be to identify new anticancer candidates that are able to bypass this resistance
mechanism entirely by inducing p53-independent antiproliferation.
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The success of Ru (III)-containing compounds such as KP1019, which received favorable
evaluations in preclinical studies,8-9 has lead to recent interest in the development of more active
Ru (II) complexes as anticancer agents.10-12 Some prominent examples of cytotoxic Ru (II)
complexes that have undergone preclinical studies are RAPTA-C, RM175 and RDC11.13-16
However, many of these Ru (II) complexes have p53-dependent activity and are subjected to the
same resistance mechanism of p53-mutated cancers. Previous studies indicated that RAPTA-C,
RM175 and RDC11 induced p53 and its target genes to various degrees.17-19 Many other reported
Ru (II) complexes also exert their antiproliferation effect via p53-dependent pathways.20-26 In
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contrast, only a limited number of Ru (II) complexes have been reported to demonstrate p53-
independent activity (Fig 1). Several Ru (II)-arene complexes bearing azopyridine,
iminopyridine or chloroquine ligand, and a imidazole-bearing phenanthroline Ru (II) complex
demonstrated similar toxicity in both wild type colorectal carcinoma HCT116 and p53-null
lineage.27-29 Another Ru (II)-arene complex bearing iminophosphorane ligand induced apoptotic
cell death without p53-induction.30 Other metal complexes of Pt and Os have also been shown to
act via p53-independent mechanism.27, 31-33 In all cases, the metal centre has little or no influence
on the p53-dependence of these complexes; the p53-independence of these complexes could
often be ‘switched on’ or ‘switched off’ by structural modification to the ligands.27, 33 In light of
this, a greater understanding of the structural determinents of p53-independence could lead to the
identification of new organoruthenium compounds with p53-independent activity, adding to the
limited pool of drugs for the effective treatment of resistant cancers with mutated p53 status.
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Previously, a Ru (II)-Arene Schiff-base (RAS) complex bearing 1,3,5-triisopropylbenzene
(TPB) and iminoquinoline ligands, RAS-1T, was identified as a promising lead cytotoxic
compound with an unique mode-of-action.34 Preliminary investigations suggested that RAS-1T
acted via a mechanism distinct from classical alkylating agents including cisplatin (CDDP) and
other reported anticancer Ru (II) complexes. RAS-1T did not induce p53 overexpression and
was equipotent in epithelial cells BJwt (p53+/+) and BJshp53 (p53-/-). Further investigation of
RAS-1T and its hexamethylbenzene (HMB) analogue, RAS-1H, revealed that both compounds
induced non-apoptotic cell death via ER-stress pathways but the pathways were different for
each compound.35
In this work we report a structure-activity relationship study on an expanded set of RAS
complexes that are structurally related to RAS-1H (4) and RAS-1T (5), aimed to (i) investigate
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if p53-independent activity is a general feature of this class of iminoquinoline-containing Ru (II)
complexes and (ii) identify the key structural features that lead to their p53-independence. We
further discuss their activity against a panel of colorectal (HCT116 and SW480) and gastric
(AGS and KATOIII) cancer cells, the influence of the arene and chelating ligands on RAS
complexes’ stability to aquation and biological nucleophiles, as well as intracellular
accumulation.
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RESULTS & DISCUSSION
Synthesis and Characterization of RAS Complexes
RAS complexes 1 – 9 (Fig 2) were synthesized using the general reaction route as shown in
Scheme 1. Recently, we reported the synthesis and characterization of 4, 5 and 9.34-35 2-
Quinolinecarboxaldehyde and different aniline derivatives were reacted in dry EtOH or MeOH to
give the bidentate imine ligand with varying degree of purity. Most of the imine ligands could
not be isolated as pure compounds due to the facile hydrolysis of the imine bond and the crude
product was used directly for synthesis of the RAS complexes by addition of stoichiometric
amount of the corresponding [Ru(η6-arene)Cl2]2 in MeOH. Complexes 1 – 9 were further
purified and isolated via flash column chromatography. In addition, we report an improved
synthetic protocol of 8 and 9 that did not require purification by column chromatography.
Briefly, the imine ligand was prepared from 2-quinolinecarboxaldehyde and excess 3-
chloroaniline in toluene using a Dean-Stark apparatus and the crude ligand treated directly with
[Ru(η6-arene)Cl2]2 in MeOH. The product was filtered through a short plug of neutral alumina
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and recrystallized using vapour diffusion of diethyl ether into a saturated CH2Cl2 solution at 4
ºC.
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The H NMR spectra of 1 – 9 showed resonances typical of RAS complexes.34 The
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disappearance of the singlet peak corresponding to quinolinecarboxaldehyde at ca. 10.2 ppm and
the appearance of the peak corresponding to the imine proton at ca. 8.5-9.5 ppm indicated the
formation of the chelate ligand. The desymmetrization of the Ru complex due to chelation
resulted in additional splitting pattern of the arene proton signals (arene = toluene, cymene,
TPB). ESI-MS spectra of these compounds also showed the characteristic [M]+ molecular ion
peaks with Ru and Cl isotopic pattern. The purity of the complexes was confirmed by either
elemental analysis (EA) or RP-HPLC to be >95%.
Solid-state structural information was obtained for 4 and 9 via single crystal X-ray diffraction
studies (Fig 3) and selected structural data are given in Table 1 and Table 2. Single crystals
were obtained via slow diffusion of diethyl ether into saturated CH2Cl2 solution at 4 ºC. RAS
complexes 4 and 9 adopted the classical ‘piano-stool’ structure similar to most reported Ru(II)-
arene complexes with some notable deviations.26, 30, 36 The Ru-Carene bond ranges for 4 and 9 of
2.196-2.283 Å and 2.175-2.280 Å, respectively, are longer than RM175 (biphenyl) and RAPTA-
C (cymene) of 2.161-2.231 Å and 2.180-2.262 Å, respectively, presumably due to the sterically
encumbered HMB ligand. All corresponding bond length and angles between 4 and 9 were
similar, suggesting that changing the 4-OMe group to a 3-Cl group did not alter the core RAS
structure. The conformation adopted by the iminoquinoline ligands could be described by the
torsion angle θC22-N2-C23-C24 of 37.6(9)˚ and 34.6(6)˚ for 4 and 9, respectively, disrupting the
extended conjugation within the chelate ligand. This twisted conformation was presumably due
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to the steric bulk of the arene ligand. The same structural feature was observed in the recently
reported RAS complexes.34
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Effect of structural variation on stability and Log POW
Aqueous stability and Log POW are important factors that affect a potential drug’s
bioavailability and cell permeability,37 which can potentially influence the molecular target and
mode-of-action. To determine the stability of the 9 RAS complexes under investigation towards
aquation and biological nucleophiles, we monitored their UV-Vis profile in double distilled H2O
(ddH2O) and Hanks Balanced Salt Solution (HBSS) containing 10% fetal bovine serum (FBS)
over 24 h. Any reaction would manifest as a shifts in the UV-Vis profile, producing isosbestic
point(s) in the overlapping spectrums at different time points. The investigated RAS complexes
were generally stable in water (Fig S1). Slight shifts in the spectrums and the formation of
several isosbestic points were observed for 6, 7 and 8 after 24 h in water, which suggests that the
aquation of the Ru-Cl bond in 6, 7 and 8 occurred very slowly in the absence of other competing
nucleophiles at r. t..
Although all RAS complexes could be considered to be resistant to aquation in water, only
some could be considered stable towards interfering nucleophiles as seen in their stability in
serum-containing HBSS (Fig S2). This is particularly well illustrated by focusing on the slight
variations implemented when comparing 4, 5, 8 and 9 (Fig 4). In general, compounds with more
π-donating arene (HMB, TPB) and less π-acidic iminoquinoline (-OMe) ligands stabilized the
Ru-Cl bonds making it less susceptible towards substitution. For instance, 5 was stable in HBSS
with 10% FBS. However, changing the 4-OMe to the more π-acidic 3-CF3 or 3-Cl group (7 and
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8) labilised the Ru-Cl bond. Replacing the TPB arene with the more π-donating HMB in 9
resulted in a more stable complex resistant to Ru-Cl aquation. Furthermore, it cannot be ruled out
that compounds bearing labile benzene or toluene arene ligands (1 and 2) decomposed in serum-
containing buffer due to the displacement of arene ligand by biological nucleophiles (Fig S2).
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Log POW of 1 – 5 and 9 were determined using the shake-flask method.38 Log POW of 6, 7 and 8
could not be determined as their spectra were significantly shifted after the partitioning and
phase separation (data not shown), presumably due to non-specific interactions with the n-
octanol phase. The trend in the Log POW studies were as expected, where RAS complexes with
more hydrophobic ligands displayed higher Log POW values (Table 3).
Effects of ligand structural variations on cytotoxicity profile
To determine the effects of structural variation to cytotoxicity, we obtained the IC50 values of
the nine RAS complexes in colorectal cancer cell lines HCT116 and SW480, and gastric cancer
cell lines AGS and KATOIII (Table 3). In general, RAS complexes bearing the more
hydrophobic HMB and TPB arene ligands demonstrated the highest cytotoxicity in the cell lines,
with IC50 values in the low µM range. Complex 5 displayed the greatest efficacies with IC50 ca.
1.0 µM in HCT116, AGS and KATOIII and 4.1 µM in SW480, which was 34- and 7-fold more
cytotoxic than CDDP in AGS and KATOIII, respectively. For RAS complexes with 4-OMe (1 –
5), IC50 values decreased with the increasing hydrophobicity of the arene ligand, from the least
hydrophobic benzene in 1 to the most hydrophobic TPB in 5. Interestingly, starting from
complex 3 in the reverse order, the IC50 became higher in HCT116 and AGS versus SW480 and
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KATOIII, suggesting that hydrophobicity also affected selectivity between different cancer
types.
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Both 4 and 9 demonstrated biphasic dose-response curve in AGS and KATOIII cells,
respectively. AGS cell viability decreased with increasing concentration of 4 to reach a plateau
of 40% between 5 µM and 20 µM before dropping drastically to 0% above 20 µM (Fig 5a).34 A
similar biphasic profile was seen in KATOIII cells treated with 9 (Fig 5b). In contrast, the TPB
analogues (5 and 8) did not display such biphasic dose-response profiles. The basis for this
unique concentration-dependent mechanism of antiproliferative activity induced by HMB-
bearing RAS complexes (4 and 9) remains to be elucidated. This biphasic dose-response was not
observed in benzene, toluene, cymene or TPB RAS analogues. Few reported anticancer
compounds exhibit such biphasic dose-response profiles. One such compound is anti-tubule
taxane Docetaxal, which induced mitotic catastrophe at lower doses and p53-independent
apoptosis at higher doses in prostate cancer (PC3, DU145) and breast cancer cell lines (MCF7,
MDA-MB-231).39-42 A similar concentration-dependent dual mode-of-action for 4 and 9 would
be highly conceivable.
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RAS complexes accumulate in cells mainly by passive diffusion
One possibility for the higher cytotoxicity of compounds bearing hydrophobic arene ligands
could be attributed to better cellular uptake via passive diffusion.43 More efficient cellular
accumulation of the RAS compounds may result in a higher efficacy. The RAS complexes
evaluated were considered hydrophilic compounds by virtue of their Log Pow values. We
investigated whether varying the hydrophobicity of the arene ligands (1 – 5) affected cellular
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uptake in HCT116 cells, with 6 and OXP for comparison. Cell samples were centrifuged,
washed, and digested separately in concentrated HNO3 before their Ru/Pt contents were analyzed
by ICP-MS (Table S1).
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There was a positive correlation between hydrophobicity, cellular accumulation and
cytotoxicity for 1 – 5 (Fig 6a-b). When treated at the same concentration of 1.5 µM, the most
hydrophobic complex 5 accumulated 7.5-times more than 1, the least hydrophobic complex.
Interestingly, complex 6 accumulated 3 times less than complex 4 but had a higher cytotoxicity.
Hence, with the exception of complex 6, the higher uptake also corresponded with lower IC50
suggesting that uptake was partially responsible for the higher cytotoxicity of the more
hydrophobic RAS complexes. In addition, there was an interesting abrupt improvement of drug
intake at a Log Pow just above -1.5, suggesting that this value might be considered as a goal for
designing cytotoxic Ru (II) arene compound.
Since the RAS complexes are cationic and bear hydrophobic ligands, we investigated if
organic cation transporters (OCT) could be responsible for their activity in colorectal cancer in a
similar manner as OXP. OXP was effective in the treatment of colorectal cancer compared to
CDDP and attributed to the fact that OXP was a substrate for OCT1/2 and would be more
efficiently accumulated in OCT-overexpressing colorectal cancer cells.44 As the RAS complexes
are positively charged and contained hydrophobic ligands, they could possibly be substrates of
OCTs in a similar fashion to OXP. Hence HCT116 cells were treated with 5 and 6 for 7 h in the
presence and absence of OCT2-inhibitor cimetidine and compared to OXP as a positive control.
Cimetidine did not affect the viability of cells treated with 5 and 6, suggesting that both are not
substrates of OCT2 (Fig 7a-b). In contrast, the viability of cells treated with OXP increased in
the presence of cimetidine (Fig 7c), indicating reduced uptake of the cytotoxic OXP. Taken
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together, the data suggested that RAS complexes accumulated in the cell mainly via passive
diffusion. A general trend indicated that increasing hydrophobicity resulted in increased cellular
uptake and higher toxicity, which was characteristic of entry by passive diffusion. However,
exception such as in complex 6 indicated that additional mechanisms and physicochemical
determinants might exist and could participate in the uptake process. Although, OCT transporters
were not involved in the uptake of RAS complexes, the possibility that other energy-dependent
uptake pathways are involved in the cellular accumulation of RAS complexes cannot be
excluded.
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Structural determinant of p53-Independent mode-of-action
We determined previously that 4 and 5 induced cell death in AGS and HCT116 cells without
concomitant induction of p53.34-35 To further verify the p53-independence of the RAS
complexes, we evaluated the expression of p53 as well as downstream, cell cycle-arresting and
apoptosis-associated biomarkers in treated HCT116 cells. Cells were exposed to 4, 5, 8 and 9 for
2 h, 6 h and 24 h at their IC50 and IC75 values. In addition, their activities were probed in the
absence and presence of p53-inhibitor pifithrin-α. OXP was used as a positive control.
Western blot analysis was performed against p53 and its targets, cell cycle-regulating cyclin
D1 and p21 (Fig 8a).45-47 In general, there was a lack of induction of these biomarkers by 4, 5, 8
and 9 although p53-independent induction of p21 was observed with 4, 5 and 9 after 24 h
exposure.48 In contrast, OXP exhibited a concentration and time-dependent upregulation of p53,
cyclin D1 and p21. RT-qPCR analyses were performed on p53-regulated anti-apoptotic gene
BCL2 and pro-apoptotic genes NOXA and BAX (Fig 8b).49-51 Complexes 4 and 9 induced a slight
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increase in NOXA expression at the later time points (6 h and 24 h) while 5 and 8 did not cause
significant change in NOXA expression. Cells treated with 4, 5, 8 and 9 also induced negligible
change in expression of BAX and BCL2. In contrast, OXP treatment at IC75 led to 3-fold
induction of NOXA, 5.5-fold increase in BAX and a significant reduction in BCL2 after 24 h
exposure. To demonstrate functionally that the cytotoxic activities of 4, 5, 8 and 9 were
independent of p53, cells were co-incubated with pifithrin-α. Co-treatment of 4, 5, 8 and 9 with
pifithrin-α improved the efficacies of RAS complexes while in the case of OXP, co-treatment
incurred a 3-fold increase in IC50 suggesting that OXP exerted its cytotoxicity via a p53-
dependent pathway.
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The lack of significant induction of p53 and its downstream targets implied that these RAS
complexes exerted their antiproliferative effects via p53-independent pathways. On the other
hand, OXP significantly increased the expression of p53, cyclin D1, p21, NOXA and BAX and
decreased the expression of BCL2. This was in agreement with previous studies, which showed
that oxaliplatin induced p53-mediated cycle arrest and (to a smaller degree) p53-mediated
apoptosis in HCT116 cells.52-53 Furthermore, p53 inhibition with pifithrin-α did not adversely
affect RAS cytotoxicities compared to OXP. This further confirmed that the RAS complexes
acted via a p53-independent mode-of-action.
To ascertain that the p53-independent activity is a general characteristic of this class of
iminoquinoline-containing RAS complexes, we measured p53 expression following 24 h
treatment at IC50 concentrations for the remaining complexes 1, 2, 3, 6 and 7 as well as measured
viabilities of treated cells after 48 h in the absence and presence of pifithrin-α. Likewise, OXP
was used as a positive control. There was a lack of p53 induction by these RAS complexes and
their activities not negatively impacted by the presence of pifithrin-α, as seen by the similar or
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lower cell viabilities in cells co-treated with pifithrin-α (Fig S3), in keeping with the other RAS
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complexes.
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In general, all RAS complexes tested in this studies demonstrated p53-independent activity,
even though their physicochemical properties such as stability to nucleophiles and
hydrophobicity differed significantly. A similar class of Ru (II) arene complexes bearing imino-
and azo-pyridine ligands (Fig 1; top-right) has been shown to demonstrate p53-independent
activity only when the chloride ligand was substituted with iodide.27 It was reported that the
choice of halide ligand strongly influenced the p53-dependence on their class of compound. The
key difference with RAS complexes was that they demonstrated p53-independent activities
without the need for iodide substitution. The iminoquinoline and Ru (II)-arene functionalities
common to all 9 RAS complexes were crucial for the observed p53-independence cytotoxicity
and structural tuning at the non-halide sites could be used to fine-tune their antiproliferative
modes-of action.
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CONCLUSION
The current study gives insights to the structural determinants of cytotoxicity and p53-
independence in Ru (II)-arene complexes through a detailed chemical and in vitro SAR study of
9 RAS complexes. In general, the more hydrophobic complexes displayed higher cellular
accumulation and cytotoxicity across a panel of cancer cell lines. In addition, the lack of OCT
involvement in the uptake of RAS complexes suggested that entry occurred mainly via passive
diffusion, despite the fact that these complexes were hydrophilic in nature. RAS complexes also
displayed tolerance towards aquation, although analogues with benzene or toluene ligands were
significantly less stable towards biological nucleophiles as these arenes were much more prone
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to displacement. These complexes also exerted their anticancer activities via p53-independent
pathways as seen in the lack of activation of p53 and downstream targets, and their unchanged
activity in the presence of small molecule p53-inhibitors. Compounds such as this may have
important implication in the treatment of resistant cancers with mutated p53 or p53-null status.
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EXPERIMENTAL
Materials. All experimental procedures were carried out without additional precautions to
exclude air or moisture unless otherwise specified. All chemicals and solvents were used as
received unless otherwise specified. Dry ethanol and methanol were obtained by drying them
over molecular sieves 3-4 Å 24 h before use. RuCl3.xH2O was purchased from both Precious
Metals Online. [(η6-benzene)RuCl2]2, [(η6-toluene)RuCl2]2, [(η6-cymene)RuCl2]2, [(η6-
hexamethylbenzene)RuCl2]2 and [(η6-1,3,5-triisopropylbenzene)RuCl2]2 were synthesized
according to previously reported protocols.43, 54-56 Thiazolyl blue tetrazolium bromide (MTT),
Trizma® Base BioUltra, Nonidet P-40, DL-Dithiothreitol, Non-fat Dried Milk Bovine,
TWEEN® 20, Ponceau S and Cimetidine, were purchased from Sigma-Aldrich. All other
chemicals used were purchased from Sigma-Aldrich (Singapore). Ultrapure water used was
purified by a Milli-Q UV purification system (Sartorius Stedim Biotech SA). Gibco® Versene
solution, Gibco® Trypsin/EDTA solution, 10% SDS solution, Penicillin-Streptomycin (10 000
U/mL), Dulbecco's Phosphate-Buffered Saline (10x), TRIzol® Reagent and Applied
Biosystem® High Capacity cDNA Reverse Transcription Kit were purchased from Life
Technologies. HycloneTM RPMI 1640, DMEM medium and Fetal bovine serum (FBS) was
purchased from Thermo Fisher Scientific Inc. Bio-rad Protein Assay Dye Reagent Concentrate,
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40% Acrylamide/Bis solution, 10x Tris/glycine buffer, TEMED, 4x Laemmli Sample Buffer,
Nitrocellulose Membrane, 0.2 µm and 0.45 µm were purchased from Bio-rad Laboratories.
cOmplete, mini Protease Inhibitor Cocktail Tablets, RNase A, FastStart Universal Probe Master
(Rox) were purchased from Roche Diagnostics. LuminataTM Classico Crescendo Western HRP
Substrate were purchased from Merck Millipore Corporation.
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Instrumentation. H NMR spectrums were obtained using either a Bruker AMX 300, Avance
400 or AMX 500 spectrometer and the chemical shifts (δ) were reported in parts per million with
reference to residual solvent peaks. Electrospray-ionization Mass Spectrometry (ESI-MS) spectra
were obtained using Thermo Finnigan MAT ESI-MS System. UV-vis spectra were obtained
using the Shimadzu UV-1800 UV Spectrophotometer with a TCC-240A Temperate Controlled
Cell Holder. Ru concentrations were determined using the Optima ICP-OES (Perkin-Elmer)
operated by CMMAC, NUS. Elemental analyses of selected Ru complexes were carried out
using a Perkin-Elmer PE 2400 elemental analyzer by CMMAS, NUS.
HPLC analysis of compound purity. Determination of the purity of the RAS Complexes were
done using analytical HPLC on a Shimadzu Prominence System equipped with a DGU-20A3
Degasser, two LC-20AD Liquid Chromatography Pump, a SPD-20A UV/Vis Detector and a
Shim Pack GVP-ODS 2.0 mm 18 column (5 µM, 120Å, 250 mm x 4.60 mm i.d.) at r.t. at a flow
rate of 1.0 mL/min with detection at both 214 nm and 254 nm. The gradient elution conditions
were as follows: 20-80% solvent B over 30 min, where solvent A is 10 mM NH4OAc pH 7.0 and
solvent B is CH3CN.
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Synthesis of 1, [(η -benzene)RuCl(4-methoxy-N-(2-quinolinylmethylene)-aniline)]Cl. 2-
Quinolinecarboxaldehyde (78.6 mg, 0.5 mmol) and p-anisidine (61.5 mg, 0.5 mmol) was added
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to dry EtOH (10 mL) and and stirred at r.t. over 24 h. The solvent was removed in vacuo and
dried. The resultant crude 4-methoxy-N-(2-quinolinylmethylene)-aniline (94 mg, approx. 0.36
mmol) was treated with [(η6-benzene)RuCl2]2 (89.6 mg, 0.18 mmol) in MeOH (20 mL) and
stirred at r.t. over 12 h. The solvent was removed in vacuo and the resulting solid purified by
gradient elution column chromatography (1:1 v/v EtOH/CHCl3, Rf = 0.2; EtOH, Rf = 0.5). The
purified product was redissolved in CHCl3 and filtered through a syringe column to remove trace
amount of dissolved silica, before the solvent was removed in vacuo. The final product was then
dried in vacuo for 1 h to give a dark red solid. Yield: 40 mg (22%). 1H NMR (400 MHz, D2O): δ
8.68 (d, 3J= 9 Hz, 1H, quinolinyl), 8.57 (s, 1H, PhN=CH), 8.35 (d, 3JHH = 8 Hz, 1H, quinolinyl),
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8.07 (t, JHH = 8 Hz, 1H, quinolinyl), 8.01 (d, JHH = 8 Hz, 1H, quinolinyl), 7.87 (t, JHH = 8 Hz,
1H, quinolinyl), 7.81 (d, 3JHH = 8 Hz, 1H, quinolinyl), 7.74 (d, 3JHH = 9 Hz, 2H, C6H4OMe), 7.05
3
(d, JHH = 9 Hz, 2H, C6H4OMe), 5.83 (s, 6H, C6H6), 3.88 (s, 3H, OCH3) ppm. ESI-MS (+ve
mode): m/z = 477 [M]+. ). Purity of the complex was determined to be >95% pure by RP-HPLC
(% Purity): 95.8% at 214 nm and 96.4% at 254 nm; tr = 12.5 min.
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Synthesis of 2, [(η -toluene)RuCl(4-methoxy-N-(2-quinolinylmethylene)-aniline)]Cl. 2 was
obtained with [(η6-toluene)RuCl2]2 (94.6 mg, 0.18 mmol) and crude 4-methoxy-N-(2-
quinolinylmethylene)-aniline (94 mg, approx. 0.36 mmol) using the same protocol as used for 1
(Gradient elution: 1:4 v/v EtOH/CHCl3, Rf = 0.2; EtOH, Rf = 0.7). The final product was then
dried in vacuo for 1 h to give a dark red solid. Yield: 65 mg (35%). 1H NMR (400 MHz, DMSO-
3
3
d6): δ 9.16 (s, 1H, PhN=CH), 8.89 (d, JHH = 8 Hz, 1H, quinolinyl), 8.76 (d, JHH = 9 Hz, 1H,
3
3
quinolinyl), 8.31 (t, JHH = 8 Hz, 2H, quinolinyl), 8.15 (t, JHH = 7 Hz, 1H, quinolinyl), 7.99 (m,
3
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3H, quinolinyl /C6H4OMe), 7.22 (d, JHH = 9 Hz, 2H, C6H4OMe), 6.07 (t, JHH = 6 Hz, 1H,
C6H5), 6.01 (d, 3JHH = 6 Hz, 1H, C6H5), 5.81 (t, 3JHH = 6 Hz, 1H, C6H5), 5.71 (t, 3JHH = 6 Hz, 1H,
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C6H5), 5.37 (d, JHH = 6 Hz, 1H, C6H5), 3.90 (s, 3H, ArOCH3), 2.17 (s, 3H, CH3) ppm. ESI-MS
(+ve mode): m/z = 491 [M]+. Purity of the complex was determined to be >95% pure by RP-
HPLC (% Purity): 98.1% at 214 nm and 95.8% at 254 nm; tr = 14.4 min.
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Synthesis of 3, [(η -cymene)RuCl(4-methoxy-N-(2-quinolinylmethylene)-aniline)]Cl. 3 was
obtained with [(η6-cymene)RuCl2]2 (109.72 mg, 0.179 mmol) and crude 4-methoxy-N-(2-
quinolinylmethylene)-aniline (94 mg, approx. 0.358 mmol) using the same protocol as used for 1
(Gradient elution: 1:4 v/v EtOH/CHCl3, Rf = 0.3; 1:1 v/v EtOH/CHCl3, Rf = 0.5). The final
1
product was then dried in vacuo for 1 h to give a dark orange solid. Yield: 125 mg (61%). H
3
NMR (400 MHz, DMSO-d6): δ 9.15 (s, 1H, PhN=CH), 8.91 (d, JHH = 8 Hz, 1H, quinolinyl),
3
3
3
8.73 (d, JHH = 9 Hz, 1H, quinolinyl), 8.32 (t, JHH = 7 Hz, 2H, quinolinyl), 8.17 (t, JHH = 7 Hz,
1H, quinolinyl), 8.01 (t, 3JHH = 7 Hz, 1H, quinolinyl), 7.96 (d, 3JHH = 9 Hz, 2H, C6H4OMe), 7.23
(d, 3JHH = 9 Hz, 2H, C6H4OMe), 6.11 (d, 3JHH = 6 Hz, 1H, i-PrC6H4Me), 5.89 (d, 3JHH = 6 Hz, 1H,
3
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i-PrC6H4Me), 5.75 (d, JHH = 6 Hz, 1H, C6H4), 5.39 (d, JHH = 6 Hz, 1H, C6H4), 3.91 (s, 3H,
ArOCH3), 2.32 (sept, 1H, 3JHH = 7 Hz, CH(Me)2), 2.24 (s, 3H, i-PrArCH3), 0.90 (d, 3JHH = 7 Hz,
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3H, CH(CH3)2), 0.74 (d, JHH = 7 Hz, 3H, CH(CH3)2) ppm. ESI-MS (+ve mode): m/z = 533
[M]+. Purity of the complex was determined to be >95% pure by RP-HPLC (% Purity): 99.4% at
214 nm and 96.0% at 254 nm; tr = 19.0 min.
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Synthesis of 4, [(η -hexamethylbenzene)RuCl(4-methoxy-N-(2-quinolinylmethylene)-
aniline)]Cl. Complex 4 was synthesized according to published procedure.35
6
Synthesis of 5, [(η -1,3,5-triisopropybenzene)RuCl(4-methoxy-N-(2-quinolinylmethylene)-
aniline)]Cl. Complex 5 was synthesized according to published procedure.34
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Synthesis of 6, [(η -1,3,5-triisopropybenzene)RuCl(N-(2-quinolinylmethylene)-1-naphthyl-
amine)]Cl. 2-Quinolinecarboxaldehyde (78.6 mg, 0.5 mmol) and 1-napthylamine (53.2 mg, 0.5
mmol) was added to dry EtOH (10 mL) and and stirred at r.t. over 24 h. The solvent was
removed in vacuo and dried. The resultant crude N-(2-quinolinylmethylene)-1-naphthylamine
(141 mg, approx. 0.5 mmol) was treated with [(η6-1,3,5-triisopropylbenzene)RuCl2]2 (188 mg,
0.25 mmol) in MeOH (20 mL) and stirred at r.t. over 12 h. The solvent was removed in vacuo
and the resulting solid purified by column chromatography (1:4 v/v EtOH/CHCl3, Rf = 0.6). The
purified product was redissolved in CHCl3 and filtered through a syringe column to remove trace
amount of dissolved silica, before the solvent was removed in vacuo. The final product was then
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dried in vacuo for 1 h to give a dark red solid. Yield: 198 mg (62%). H NMR (400 MHz,
CD3OD): δ 9.12 (s, 1H, PhN=CH), 8.91 (d, 3JHH = 12 Hz, 1H, quinolinyl), 8.86 (d, 3JHH = 11 Hz,
1H, quinolinyl), 8.18 (m, 8H, quinolinyl/napthyl), 7.72 (m, 3H, quinolinyl/napthyl), 5.86 (s, 3H,
C6H3), 2.18 (sept, 3JHH = 9 Hz, 3H, CHMe2), 1.07 (d, 3JHH = 9 Hz, 9H, CH(CH3)2), 0.98 (d, 3JHH
= 9 Hz, 9H, CH(CH3)2) ppm. ESI-MS (+ve mode): m/z = 623 [M]+. Purity of the complex was
determined to be >95% pure by elemental analysis. Anal. Calcd for C35H38Cl2N2Ru·H2O (%): C
62.12, H 5.96, N 4.14; Found: C 62.29, H 5.59, N 4.18.
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Synthesis of 7, [(η -1,3,5-triisopropybenzene)RuCl(3-trifluoromethyl-N-(2-quinolinyl-
methylene)aniline)]Cl. 2-Quinolinecarboxaldehyde (78.6 mg, 0.5 mmol) and 3-
(trifluoromethyl)aniline (62.5 µl, 0.5 mmol) was added to dry EtOH (10 mL) and and stirred at
r.t. over 24 h. The solvent was removed in vacuo and dried. The resultant crude 3-
trifluoromethyl-N-(2-quinolinylmethylene)aniline (150 mg, approx. 0.5 mmol) was treated with
[(η6-1,3,5-triisopropylbenzene)RuCl2]2 (188 mg, 0.25 mmol) in MeOH (20 mL) and stirred at r.t.
over 12 h. The solvent was removed in vacuo and the resulting solid purified by gradient elution
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column chromatography (1:4 v/v EtOH/CHCl3, Rf = 0.4; 1:1 v/v EtOH/CHCl3, Rf = 0.7). The
purified product was redissolved in CHCl3 and filtered through a syringe column to remove trace
amount of dissolved silica, before the solvent was removed in vacuo. The final product was then
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dried in vacuo for 1 h to give a light brown solid. Yield: 156 mg (46%). H NMR (400 MHz,
DMSO-d6): δ 9.23 (s, 1H, PhN=CH), 8.96 (d, 3JHH = 9 Hz, 1H, quinolinyl), 8.91 (d, 3JHH = 8 Hz,
1H, quinolinyl), 8.53 (s(broad), 1H, C6H4), 8.43 (d, 3JHH = 8 Hz, 1H, C6H4), 8.31 (t, 3JHH = 8 Hz,
3
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2H, quinolinyl), 8.16 (t, JHH = 9 Hz, 1H, quinolinyl/C6H4), 8.01 (t, JHH = 8 Hz, 2H,
quinolinyl/C6H4), 7.91 (t, 3JHH = 8 Hz, 1H, quinolinyl/C6H4), 5.60 (s, 3H, C6H3), 2.37 (sept, 3JHH
= 7 Hz, 3H, CHMe2), 1.16 (d, 3JHH = 7 Hz, 9H, CH(CH3)2), 0.84 (d, 3JHH = 7 Hz, 9H, CH(CH3)2)
ppm. ESI-MS (+ve mode): m/z = 641 [M]+. Purity of the complex was determined to be >95%
pure by elemental analysis. Anal. Calcd for C32H35Cl2F3N2Ru·3H2O (%): C 52.60, H 5.66, N
3.83; Found: C 52.43, H 5.46, N 3.84.
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Synthesis of 8, [(η -1,3,5-triisopropybenzene)RuCl(3-chloro-N-(2-quinolinylmethylene)-
aniline)]Cl. 2-Quinolinecarboxaldehyde (78.6 mg, 0.5 mmol) and 3-chloroaniline (52.9 mg, 0.5
mmol) was added to dry EtOH (10 mL) and stirred at r.t. over 24 h. The solvent was removed in
vacuo and dried. The resultant crude 3-chloro-N-(2-quinolinylmethylene)aniline (133 mg,
approx. 0.5 mmol) was treated with [(η6-1,3,5-triisopropylbenzene)RuCl2]2 (188 mg, 0.25 mmol)
in MeOH (20 mL) and stirred at r.t. over 12 h. The solvent was removed in vacuo and the
resulting solid purified by column chromatography (1:4 v/v EtOH/CHCl3, Rf = 0.6). The purified
product was redissolved in CHCl3 and filtered through a syringe column to remove trace amount
of dissolved silica, before the solvent was removed in vacuo. The final product was then dried in
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vacuo for 1 h to give a dark red solid. Yield: 89 mg (28%). H NMR (400 MHz, MeOD-d4): δ
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quinolinyl), 8.25 (m, 3H, quinolinyl/C6H4), 8.15 (t, JHH = 8 Hz, 1H, quinolinyl/C6H4), 8.06 (dt,
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Hz, 2H, quinolinyl/C6H4), 5.60 (s, 3H, C6H3), 2.49 (sept, 3JHH = 7 Hz, 3H, CHMe2), 1.23 (d, 3JHH
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= 7 Hz, 9H, CH(CH3)2), 0.97 (d, JHH = 7 Hz, 9H, CH(CH3)2) ppm. ESI-MS (+ve mode): m/z =
607 [M]+. Purity of the complex was determined to be >95% pure by elemental analysis. Anal.
Calcd for C31H35Cl3N2Ru·3H2O (%): C 53.41, H 5.93, N 4.02; Found: C 53.02, H 5.66, N 3.87.
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Synthesis
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9,
[(η -hexamethylbenzene)RuCl(3-chloro-N-(2-quinolinylmethylene)-
aniline)]Cl. Complex 9 was synthesized according to published procedure.34
Alternative synthetic route for 8 and 9. 2-Quinolinecarboxaldehyde (314 mg, 2 mmol) and 3-
chloroaniline (268 mg, 4 mmol) were placed to a 50 mL round-bottom flask equipped with a
Dean-Stark apparatus. 25 mL of toluene was added to dissolve the reactants and the resulting
brown solution was refluxed overnight. The mixture was allowed to cool and concentrated under
reduced pressure to give the dark brown oil which was dissolved in a minimal amount of diethyl
ether. The black insoluble residue was filtered and the solution was again concentrated under
reduced pressure and dried in vacuo to give a brown semi-solid. The crude product containing
small amount of unreacted 3-chloroaniline was used without additional purification. 1H NMR
(300 MHz, d6-DMSO): δ 8.79 (s, 1H, Ph-N=CH), 8.53 (d, J = 8.6 Hz, 1H, quinolinyl), 8.29 (d, J
= 8.6 Hz, 1H, quinolinyl), 8.12 (dd, J = 15.9, 7.9 Hz, 2H, quinolinyl), 7.90 – 7.83 (m, 1H,
quinolinyl), 7.76 – 7.69 (m, 1H, quinolinyl), 7.54 – 7.47 (m, 2H, quinolinyl), 7.43 – 7.35 (m, 2H,
quinolinyl). The corresponding [(η6-arene)RuCl2]2 (0.15 mmol) and 3-chloro-N-(2-
quinolinylmethylene)-aniline (180 mg, 0.675 mmol) were dissolved in 10 mL of dry MeOH
under N2 atmosphere and stirred for 48 h. A resulting brown solution was evaporated under
reduced pressure and the brown residue was washed with copious amounts of diethyl ether. The
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solid was subsequently dissolved in minimal amount of CH2Cl2 and eluted through a short layer
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neutral
Al2O3 (5 cm)
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CH2Cl2
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unreacted
3-chloro-N-(2-
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9
quinolinylmethylene)aniline and 3-chloroaniline. The product was eluted with CHCl3/MeOH
mixture (9:1) and the red filtrate was evaporated to dryness under reduced pressure. The red solid
was redissolved in a minimal amount of dichloromethane and centrifuged (13,000 rpm for 7 min)
to remove the undissolved residues. The red microcrystals (8) or needle-shaped X-ray quality
single crystals (9) were produced by slow diffusion of diethyl ether into the CH2Cl2 solutions.
Crystals were washed with diethyl ether and dried in vacuo.
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Complex 8. Yield: 38 mg (20%). Anal. Calcd, C31H35Cl3N2Ru·0.25 CH2Cl2 (%): C, 56.50; H,
5.39; N, 3.22. Found: C, 56.09; H, 5.04; N, 4.46.
Complex 9. Yield: 40 mg (22%). Anal. Calcd, C28H29Cl3N2Ru·2CH2Cl2·0.25 H2O (%): C, 50.45,
H, 4.60, N, 4.06. Found: C, 50.36, H, 5.01, N, 4.21.
Tissue culture. The human colorectal carcinoma cells HCT116, human colorectal
adenocarcinoma cells SW480, human gastric adenocarcinoma AGS, human gastric carcinoma
KATOIII were acquired from ATCC® (Manassa,VA). HCT116 and SW480 were cultured in
DMEM medium containing 10% FBS and 1% Penicillin/Steptomycin (complete DMEM). AGS
was cultured in RPMI 1640 medium containing 10% FBS and 1% Penicillin/Steptomycin.
KATOIII was cultured in RPMI 1640 medium containing 20% FBS and 1%
Penicillin/Steptomycin. All cell lines were grown at 37 °C in a humidified atmosphere of 95%
air and 5% CO2. Experiments were performed on cells within 20 passages.
Determination of Log Pow. Log Pow of the RAS complexes were determined using the shake
flask method.38 The RAS complex were predissolved in ddH2O that was presaturated with n-
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octanol (for 24 h and left to stand until phase separation occurs). The UV-vis spectrum for each
samples was obtained and the absorbances at the λmax of each compound were determined. Equal
volume of n-octanol was added to each sample solution and the heterogeneous mixtures shaked
for 2 h before centrifuging at 4000 rpm for 1 min to achieve phase separation. The final
absorbance of the aqueous phase at the λmax of each compound were determined and their water-
octanol partition coefficient were calculated. All experiments were done in triplicate.
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Inhibition of cell viability assay. The anti-proliferation activity the RAS Complexes on
exponentially growing cancer cells were determined using MTT assay as described previously.57
HCT116, SW480, AGS and KATOIII were seeded at 10 000 cells per well (100 µL) in Cellstar®
96-well plates (Greiner Bio-One) and incubated for 24 h. Thereafter, cancer cells were exposed
to drugs at different concentration in media for 48 h. The final concentration of DMSO in
medium was < 1% (v/v) at which cell viability was not significantly inhibited. The medium was
removed and replaced with MTT solution (100 ꢀL, 0.5 mg/mL) in media and incubated for an
additional 45 min. Subsequently, the medium was aspirated, and the purple formazan crystals
dissolved in DMSO (100 ꢀL). The absorbance due to the dissolved purple formazan was then
obtained at 565 nm. Inhibition to cell viability was evaluated with reference to the IC50 value,
which is defined as the concentration needed for a 50% reduction of survival based on the
survival curves. IC50 values were calculated from the dose - response curves (cell viability vs
drug concentration) obtained in repeated experiments and adjusted to actual [Ru] administered,
which was determined using ICP-OES. The experiments were performed in 4 replicates for each
drug concentration and were carried out at least three times independently.
For cell viability assays involving inhibitors, Cimetidine (1.5 mM) were added together with test
compounds and incubated for 7 h. Thereafter, the drugs were removed and replace with fresh
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media for the remainder of the 48 h. Pilfithrin-α (10 µM) was incubated with drugs for the entire
48 h duration. Cell viability in the absence and presence of inhibitor was normalized against
untreated control. Experiments were performed in 3 replicates and carried out at least three times
independently.
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Drug uptake studies. HCT116 cells were grown on Cellstar® 6-well plates (Greiner Bio-One)
at a density of 500 000 cells/well for 24 h before being treated with complexes 1 – 5, 6 and
oxaliplatin at 1.5 µM for 24 h. Thereafter, the drug-containing media was removed and washed
once with EDTA/PBS followed by trypsinization. The cells were collected, counted and digested
separately with 69% nitric acid. The samples were diluted to a final concentration of 2% nitric
acid and the Ru or Pt content quantified via ICP-MS. Experiments were performed three times
independently.
Antibodies and Western blot protocol. HCT116 cells were grown on Cellstar® 6-well plates
(Greiner Bio-One) at a density of 500 000 cells/well for 24 h before being treated with
complexes 4, 5, 8 and 9 at IC50 and IC75 for 2 h, 6 h and 24 h. Oxaliplatin treatment at IC50 and
IC75 was used as a positive control for several experiments. The cells were lysed with lysis buffer
[100 µL, 1% NP40, 150 mM NaCl, 50 mM Tris-HCl (pH 8.0), protease inhibitor]. The cell
lysate were transferred to separate 2 mL tubes and sonicated for 10 s. The samples were then
centrifuged at 13000 rpm, 4˚C for 15 min. The supernatant liquid containing the proteins were
collected and total protein content of each sample was quantified via Bradford’s assay. 50 µg of
proteins from each sample were reconstituted in loading buffer [5% DDT, 1x Protein Loading
Dye] and heated at 95˚C for 5 min. The protein mixtures were resolved on a 10% SDS-PAGE gel
by electrophoresis and transferred to a nitrocellulose membrane. The proteins bands were
visualized via enhanced chemiluminescence imaging (PXi, Syngene) after treatment with the
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primary antibodies and the appropriate secondary antibodies. Equal loading of protein was
confirmed by comparison with actin. The following antibodies were used: p53 (FL-393), p21 (F-
5) and Cyclin D1 (H-295) from Santa Cruz Biotechnologies. β-Actin (ab75186) from Abcam.
ECL Anti-rabbit IgG (NA934V) and ECL Anti-mouse IgG (NA931) from GE Healthcare Life
Sciences. All antibodies were used at 1:1000 dilutions except for actin (1:10000), anti-mouse and
anti-rabbit (1:5000).
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Primers and qPCR protocol. Treatment conditions for HCT116 cells were similar to the
protocol in western blot. RNA was extracted using TRIzol® Reagent and reverse transcription
was performed with 2 µg of the extracted RNA using Applied Biosystem® High Capacity cDNA
Reverse Transcription Kit with an Applied Biosystem® 2720 Thermal Cycler. Quantitative PCR
was done on the resulting cDNA using FastStart Universal Probe Master (Rox) with Applied
Biosystem® 7500 Real Time PCR System. The relative starting quantities of genes of interest
were normalized against the housekeeping genes TBP and samples were done in duplicates. The
specificity of the amplification was controlled by a melting curve. The gene and Assay ID of
TaqMan probes are as follows: NOXA (Hs00560402_m1), BAX (Hs00180269_m1), BCL2
(Hs00608023_m1) and TBP (Hs00427620_m1).
ACKNOWLEDGEMENTS
The authors thank staff of CMMAC, NUS for performing elemental analysis and ICP-OES
analysis. They also thank Ligue contre le cancer, CNRS, European COST action CM1105,
Ministry of Education and the National University of Singapore (R143-000-638-112) for funding
the research presented in this study.
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