Synthesis, properties, and antitumor effects of a new mixed phosphine gold(I) compound in human colon cancer cells

Journal of Inorganic Biochemistry 124 (2013) 78–87

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Synthesis, properties, and antitumor effects of a new mixed phosphine gold(I) compound in human colon cancer cells Giulio Lupidi a, Luca Avenali a, Massimo Bramucci a, Luana Quassinti a, Riccardo Pettinari b, Hala K. Khalife c, Hala Gali-Muhtasib d, Fabio Marchetti e, Claudio Pettinari b,⁎ a

School of Pharmacy, Molecular Biological Section, Via Madonna delle Carceri, 62032 Camerino, Italy School of Pharmacy, Chemistry Section, Via S. Agostino 1, 62032 Camerino, Italy Department of Biology, American University of Beirut, Beirut, Lebanon d Department of Biology, Lebanese University, Beirut, Lebanon e School of Science and Technology, Chemistry Section, Via S. Agostino 1, 62032 Camerino, Italy b c

a r t i c l e

i n f o

Article history: Received 31 December 2012 Received in revised form 24 March 2013 Accepted 25 March 2013 Available online 29 March 2013 Keywords: Gold complexes Antineoplastic activity DNA fragmentation Spectroscopy Phoshine

a b s t r a c t The antineoplastic potential of a new stable mixed phosphine gold(I) complex containing tris(tert-butyl) phosphine (tBu3P) and bis(diphenylphosphino)ethene (dppet), namely [Au(tBu3P)(dppet)Cl], has been investigated in the human colon cancer HCT-116 cell line. The 31P NMR solution study, confirms the structural features observed in the solid state and, in addition, indicates partial formation of dinuclear cationic [Au(tBu3P)2]+ and [Au(dppet)2]+ species. The ionic character and strong Au–P bonds of this gold(I) species are similar to those of the most active antitumor gold compounds so far studied. The title compound was found to exhibit strong cytotoxicity, showing 85 fold greater toxicity than cisplatin (IC50 = 0.45 μM vs IC50 = 39.16 for cisplatin at 24 h) on the HCT-116 line. The cytotoxic effects were, at least partly, mediated by the induction of apoptotic cell death as evidenced by the sub-G1 cell accumulation, oligonucleosomal DNA fragmentation, caspase-3 activation and the release of cytochrome c from the mitochondria. The gold(I) compound showed little interaction with DNA measured through fluorescence quenching studies with calf thymus DNA. The inhibitory effect of the gold(I) compound on intracellular redox proteins has been also observed in pretreated HCT-116 cells. The compound was particularly effective in inhibiting thioredoxin reductase, that is likely responsible for the increased ROS production, and subsequent apoptosis induction via the mitochondrial pathway. © 2013 Elsevier Inc. All rights reserved.

1. Introduction The importance of metal complexes in cancer chemotherapy is best documented by the results obtained with platinum(II) compounds. Today, cisplatin is one of the most used anticancer drugs against a wide range of malignancies, including testicular carcinomas, ovarian tumors, head and neck cancers, bladder tumor and osteosarcoma [1]. However, significant side effects and drug resistance have limited its clinical applications. Recently, biological carriers conjugated to cisplatin analogs have improved specificity for tumor tissue, thereby reducing side effects and drug resistance [2]. The development of resistance to cisplatin represents a serious clinical problem that has prompted a great deal of investigation concerning the mechanisms by which tumor cells become resistant to this chemotherapeutic agent [3]. Platinum is not the only metal used in the treatment of cancer, various other transition metals have been used as anticancer drugs [4,5]. Among the non-platinum antitumor drugs, gold complexes, which are well known for their clinical anti-arthritic properties [6], have also attracted interest ⁎ Corresponding author. Tel.: +39 0737402234; fax: +39 0737 402007. E-mail address: [email protected] (C. Pettinari). 0162-0134/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.jinorgbio.2013.03.014

as potential antitumor agents [7–10]. Mechanistic studies suggest that, in contrast to cisplatin, DNA is not the primary target of these gold complexes. Their cytotoxicity is mediated by their ability to alter mitochondrial function and strongly inhibits both Se containing enzymes thioredoxin reductase (TrxR) and glutathione peroxidase [11–13] mechanisms that are different from those reported for platinum-derived drugs [7,8,14,15]. During the last two decades, a large variety of gold(I)] [16] and gold(III) [17] compounds have been reported to possess antiproliferative properties in vitro against several human tumor cell lines, qualifying them as excellent candidates for further pharmacological evaluation. Auranofin, the first gold(I) phosphine lipophilic complex introduced into clinical practice for chrysotherapy and for the treatment of rheumatoid arthritis [12,13,18], has been shown to induce cell death, with varying characteristics depending on the cell type [14,19]. It was shown that the same Auranofin possesses in vivo antitumor activity against P388 murine leukemia and in vitro cytotoxic potency against both B16 melanoma and P388 leukemia cells [20]. Moreover, in vitro studies indicated that Auranofin is able to overcome cisplatin resistance in human ovarian cancer cells [11,19], confirming the earlier assumption that a mechanism of action, different from the recognized DNA

G. Lupidi et al. / Journal of Inorganic Biochemistry 124 (2013) 78–87

damage induced by cisplatin, could underlie the cytotoxic activity of phosphine Au(I) drugs [6]. Another class of gold(I) phosphine derivatives containing diphosphine complexes as [Au(dppe)2] + (dppe = 1,2-bis(diphenylphosphino)ethane) was shown to exhibit a spectrum of antitumor activity in mouse tumor models [21,22]. [Au(dppe)2] +, and related tetrahedral Au(I) phosphine complexes, do not undergo ligand exchange reactions as readily as two-coordinate linear Au(I) complexes [21]. Their antitumor activity may stem from the lipophilic, cationic properties, as for other delocalized lipophilic cations that accumulate in mitochondria [6,23]. With the goal to overcome the toxicity problems, our approach has been to modify the gold(I) coordination environment in order to modulate their lipophilic/cationic properties and achieve more selective cytotoxicity. There is indeed much evidence suggesting that mitochondria play a critical role in the regulation of apoptosis (programmed cell death) by releasing several factors that lead to cell death. This function is added to those already known for mitochondria such as energy production, ion homeostasis control and hydrogen peroxide production. The level of the latter is modulated by two systems, present both in the cytosol and in mitochondria, which depend on glutathione, glutathione reductase and peroxidase [16,19] and thioredoxin, thioredoxin reductase and peroxidase [24,25]. In particular, it was found that auranofin, acting as a potent inhibitor of thioredoxin reductase, causes an alteration of the redox state of the cell, therefore creating the conditions for enhanced apoptosis [16,26]. Thioredoxin reductase is especially reactive with metal complexes, particularly platinum [26–28] and gold [29,30], and it is considered a highly specific target for antitumor agents [29]. Strong inhibition of mitochondrial thioredoxin reductase, attributable to binding of Au(I) to the redox-active selenocysteine residue [21,22,29,30], would eventually lead to altered mitochondrial functions and to the initiation of the apoptotic process. Following our previous studies on mixed phosphine gold(I) compounds with antitumor activity [31,32], we have decided to use a different gold(I) acceptor and a different diphosphine ligand. In detail, the introduction of the more basic and hindered tris(tert-butyl) phosphine (tBu3P) at the place of the triphenyl phosphine and the more rigid cis-1,2-bis(diphenylphosphino)ethene at the place of bis(diphenylphosphine)propane [31] or 1,2-bis(diphenylphosphine) ethane [32], should change the steric and electronic features of the Au(I) center and, consequently, also its physico-chemical properties in media. In the current study, we assessed the antitumor properties of the gold(I) mixed phosphine compound (Fig. 1A) in human colon cancer cell lines (HCT 116), including cisplatin as control. The gold(I) compound was also evaluated for its ability to inhibit TrxR, both (in cell lysate) and in intact HCT 116 cancer cells. Additionally, the reactive oxygen species (ROS) production and the ability to induce apoptosis, mediated by the direct inhibition of the thioredoxin reductase enzyme, were investigated. 2. Results and discussion 2.1. Synthesis and characterization [Au(tBu3P)(dppet)Cl] (complex 1) was obtained by adding the chelating cis-1,2-bis(diphenylphosphino)ethene (dppet) to a dichloromethane solution of the linear chlorotris(tert-butyl)phosphinegold(I), Au(tBu3P) Cl, in stoichiometric amounts (1:1) (Scheme 1). By addition of n-hexane an air- and moisture-stable precipitate was obtained, soluble in DMSO, acetonitrile, alcohol and chlorinated solvents. It is interesting to note that complex 1 can be obtained only by using strictly stoichiometric conditions (1:1) and not-coordinating solvents. The IR spectrum shows the bands of the phosphine ligands to be similar to those of analogous phosphine gold(I) complexes [31,32]. Gold derivatives are likely to exist as a neutral distorted tetrahedral species in the solid state, as previously observed for analogous [Au(dppe)(PPh3)Cl], however, in the absence of detailed X-ray structural data, also an ionic

79

Fig. 1. Gold (I) complex is more cytotoxic than cisplatin to human colon cancer cells. A) HCT-116 cells were treated with different concentrations of complex 1 (○,□) and cisplatin (●,■) for 24, or 48 h, respectively. Cell viability was determined by MTT assay and is reported as the percentage of viable cells. The results represent the mean value (± SD) of three independent experiments; P b 0.0001, Dunnett's test; *p b 0.01 vs vehicle.

formulation such as [Au(dppet)(t-but3P)]Cl, cannot be excluded. Conductivity measurements in DMSO (44.3 S cm2 mol−1) carried out for 1, typical of 1:1 electrolytes [33], seem to be in accordance with the latter hypothesis, but a dissociative process in strongly polar solvents can be expected even for a neutral derivative. Conductivity data in CH2Cl2 seems to support a non-ionic structure as that depicted in Scheme 1. Together with broad resonances for the cis CH= CH and the Ph groups of bridging dppet, in the 1H NMR spectrum of complex 1 carried out in CDCl3, two different signals were observed for the tert-butyl groups of the monophosphine, thus indicating the presence of an equilibrium in solution between at least two different gold(I) species. 31P{1H} NMR spectrum of complex 1 in CDCl3 (Supplementary material) showed four different resonances: that at 23.5 ppm can be assigned to a cationic [Au(dppet)2]+ species, while that at 97.4 ppm is due to a cationic [Au(tBu3P)2] + species, previously reported [34]. Furthermore, a doublet and a triplet have been respectively observed at 43.7 and 104.9 ppm, which can be both assigned to the neutral derivative 1. Both signals display the same 2J(P–P) coupling constant of 128 Hz, due to coupling of the P of tBu3P with that of dppet, further confirming our assignment. Hence, an equilibrium such as that shown in Scheme 2 is likely in chloroform. In contrast, the room temperature 31P{ 1H} NMR spectrum of complex 1 in DMSO-d6 shows only two signals due to the cationic [Au(dppet)2] + and [Au(tBu3P)2] + complexes [34]. This is a proof that in strongly coordinating solvents such as DMSO the equilibrium is completely shifted toward the cationic species and that the starting complex 1 cannot exist in DMSO. In order to further confirm the dissociation in solution of complex 1 and formation of cationic [Au(tBu3P)2] + and [Au(dppet)2] + species,

Ph Ph P t

PBut3 Au

Au(PBu 3)CI + dppet CH2CI2

P Ph Ph

Scheme 1. Synthesis of compound 1.

CI

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Ph

Ph Ph

Ph

P

Au P

CI

P

P

P

PBut3 Au

2

Ph

Ph

But3P

Ph Ph

PBut3

2 CI-

P

Ph

Ph

Au

Ph Ph

Scheme 2. Possible dissociation of compound 1 in solution.

we have also performed an ESI-MS (electrospray ionization mass spectrometry) spectrum in acetonitrile which, in the positive-ion mode, shows two peaks at m/z = 601 {[Au(tBu3P)2] +} and 989 {[Au(dppet)2] +}.

2.2. Complex 1 induces apoptosis in human colon cancer HCT-116 cells We have investigated the effect of complex 1 against HCT-116 cells. The compound efficacy was compared with the most widely used anticancer metal–drug, cisplatin. Complex 1 was found to decrease cell viability in a dose- and time-dependent fashion (Fig. 1A). IC50 values, calculated from the dose–survival curves (Table 1), showed that complex 1 was 34–87 times more active than cisplatin. The free ligand dppet had no effect on the viability of HCT-116 cells (data not shown). To determine if the decrease in cell viability is caused by necrosis, we treated the cells with acridine orange/ethidium bromide staining, which fluoresces green and red, respectively when intercalated into DNA. Of the two compounds, only acridine orange can cross the plasma membrane, whereas ethidium bromide is excluded from viable cells. If necrosis is the main pathway of cell death, then short times of incubation with the drug should result in the appearance of red stain. Fig. 2A and B shows that the majority of cells treated with complex 1 are exclusively green (i.e. acridine stained), confirming that the reduction in cell viability is not due to necrosis. After 4 and 6 h of treatment, we observed characteristic chromatin condensation (bright red inside cells) that may be indicative of apoptosis. To confirm that apoptosis is the main mechanism of cell death, we evaluated the induction of apoptosis by caspase-3 activation and cell cycle perturbation by PI staining of DNA content and flow cytometric analysis in HCT-116 cells treated with complex 1. As expected, complex 1 increased the pre-G1 population from 10 fold (at IC50) to 21 fold (2 × IC50) with respect to the control (Fig. 3) and this was accompanied by a significant decrease in the number of cells in the G0/G1 phase of the cell cycle. It has been previously reported that auranofin and other gold(I) derivatives are able to induce apoptosis in cancer cells through the activation of caspases [16,35,36]. Among the caspases, caspase-3 plays a critical role in the apoptosis signal pathway and this enzyme is commonly activated by numerous death signals and cleaves a variety of important cellular proteins [16,37], We investigated the ability of complex 1 to activate caspase-3 in HCT-116 cells treated for 18 h with IC50 concentration of this compound. Fig. 4 shows that the treatment markedly stimulated caspase-3 activity at short times with respect to the control. The enzyme activity returned to basal levels after 12 h.

Table 1 IC50 values of complex 1 for tested cell lines. IC50 mM ± SD

Complex 1 Cisplatin

IC50 ratio cisplatin/gold(1)

24 h

48 h

24 h

48 h

0.45 ± 0.042 39.16 ± 4.37

0.29 ± 0.02 9.97 ± 0.67

87 –

34 –

2.3. Complex 1 causes DNA fragmentation, increases p53 expression and cytochrome c release The inabilities of cancer cells to undergo apoptosis represent a major problem in tumor therapy and efforts in finding a way to overcome the resistance to programmed cell death constitute an active research field. The release of cytochrome c from the mitochondrial inter-membrane space to the cytosol, where it gives rise to the apoptosome complex, is a critical step in the apoptotic process [38]. To provide further evidence for the apoptotic effect of complex 1, we investigated its ability to evoke genomic DNA fragmentation which is a key hallmark of programmed cell death. We also studied its effects on p53 expression and on the activities of mitochondrial apoptosis mediators in HCT-116 cells. For this purpose, genomic DNA was isolated from HCT-116 cells treated with IC50 concentration of complex 1 for 24 h and electrophoresis was performed on 1% agarose-gel. Fig. 5A shows the presence of typical DNA laddering, an increase in mono- and oligo-nucleosome formation, which is indicative of the activation of apoptotic processes. We then evaluated by western blotting the expression levels of the p53 protein and possible cytochrome c translocation from the mitochondria to the cytosol, which forms the basis for the initiation of apoptotic events. Interestingly, we observed a dose-dependent increase in the expression of p53 protein and a marked increase in cytochrome c levels in the cytosol (Fig. 5B). This net dose-dependent increase of cytochrome c in the cytosolic fraction of cell lysates, occurred with the consequent decrease of cytochrome c in the mitochondrial fraction which was confirmed by densitometric analysis (Fig. 6C). These results indicate that complex 1 induces apoptosis by increasing cytochrome c translocation from the mitochondria to the cytoplasm. 2.4. Complex 1 inhibits thioredoxin reductase activity and increases ROS production in colon cancer cells It is well-known that possible damage to the mitochondria, can generate hydrogen peroxide and, thus, play a crucial role in the apoptotic process [26–28]. The inhibition of the enzyme TrxR increases hydrogen peroxide concentration by preventing its removal, thus causing an imbalance in cell redox conditions leading to mitochondrial membrane permeabilization and swelling [26–28]. Therefore, we measured the modification of TrxR basal activity in HCT-116 cancer cells treated with IC50 concentrations of complex 1 by using the peroxide-sensitive fluorescent probe DCFH-DA (2′,7′-dichlorodihydrofluorescein diacetate). The activity of thioredoxin reductase enzyme was first assayed in cells treated with complex 1 and results show a dose-dependent inhibition of both cytosolic and mitochondrial TrxR enzyme isoforms (Fig. 6A). The activity of the enzyme was then assayed on the total protein extract of untreated cell lysates by directly adding complex 1 to the protein extract. This was done in order to demonstrate that the main target of complex 1 the TrxR protein itself while we do not explore in this work the effect of complex 1 on possible regulation of TrxR enzyme expression. The direct addition of complex 1 to untreated cell lysates also caused a strong inhibition in the activity of the enzyme as shown in Fig. 6B. Recent studies on the mechanism of gold(I)–diphosphine compounds revealed that they specifically inhibit the enzymatic activity of thioredoxin

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A

0.5 mM

B

0 mM

10x

40x

Fig. 2. Gold (I) complex exhibited apoptotic morphological changes in colon cancer HCT-116 cells. A,B) Cells were exposed to IC50 concentration of complex 1 for 0 and 6 h, stained with acridine orange and ethidium bromide, and observed by fluorescent microscopy. Viable cells show an intact, bright green nucleus; late apoptotic cells show a red/orange nucleus with chromatin condensation. 10× (A) and 40× (B) magnifications.

reductase by binding to selenocysteine residue, without targeting other well-known selenol and thiol groups containing biomolecules [39]. When the authors examined the effect of the powerful derivative on the expression of TrxR no significant changes were observed in the content of TrxR in cells treated at different time points [39]. Progress in our work on the study of action mechanism of compound 1 will be oriented also to demonstrate the effect the compound's treatment in proliferating cells on regulation of the TrxR gene. In biological systems ROS are constantly generated and eliminated and play an important role in a variety of normal biochemical functions and abnormal pathological processes [40]. ROS generation is determined by two opposite effects: the production and the removal of free radicals, and, at least in part, the toxic effects of metal-containing drugs. This mechanism is highly probable also for Pt(II) and Pt(IV) complexes. On the contrary, Pd(II) and Cd(II) complexes have no effect on ROS production, although they show considerable toxicity [41–43]. Hence, we have investigated the ROS production in colon cancer cells treated with complex 1 at the IC50 concentration. Fig. 5C shows a substantial increase in the level of ROS at medium-long times (>30 min), indicating that this increase was due mainly to their accumulation, rather than by a pro-oxidant effect of complex 1. This conclusion is supported by the fact that thioredoxin reductase is one of the

enzymes mainly involved in the depletion of the ROS pool and its demonstrated inhibition by our compound may be the reason for the increased level of ROS. 2.5. Complex 1 does not react with calf thymus DNA Prompted by the favorable biological effects of complex 1, we investigated whether its cytotoxic effects are a consequence of a direct interaction with nuclear DNA by evaluating its potential binding to calf-thymus DNA (ct-DNA). Previous studies have demonstrated that DNA is the primary intracellular target of many anticancer drugs. This drug–DNA interaction is known to lead to DNA damage, cell cycle arrest and subsequent cell death [11,12]. Of those studies, interactions between metallic anticancer agents and DNA are important in understanding the mechanisms of their anticancer activities [13–16]. To prove or rule out the possibility of a DNA–drug interaction, a competitive ethidium bromide (EB) binding assay was undertaken to understand the mode of ct-DNA interaction with complex 1. In this assay, the molecular fluorophore EB emits intense fluorescence in the presence of ct-DNA due to its strong intercalation between the adjacent DNA base pairs. The addition of the second molecule, which binds to DNA more strongly than EB would quench the DNA–EB fluorescence

Fig. 3. Gold (I) complex caused cell cycle arrest in G0/G1 phase in colon cancer HCT-116 cells. Cell cycle analysis by PI staining with different concentrations of complex 1.

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where F0 and F are the steady-state fluorescence intensities of DNA before and after the addition of complex 1, [1] is the concentration of complex 1 as quencher. Ksv is the Stern–Volmer dynamic quenching constant. Linear Stern–Volmer plots were obtained from the fluorescence titration and Ksv for the interaction of complex 1 and ct-DNA was found from the slope of the graph (Fig. 7B). The value of the Stern–Volmer constant was evaluated and is equal to Ksv = 1.55 (±0.05) 104 M−1, a value that is not so highly relevant, to indicate that the primary target of complex 1 is not DNA. The estimation of the binding parameters was made using Eq. (2) that considers the presence of a single class of binding sites. Eq. (2) starts from the classical Scatchard equation [47], and expresses the free ligand concentration at equilibrium as a function of the analytical, total concentration of the ligand [48]: Fig. 4. Caspase 3 activity in colon cancer HCT-116 cells after treatment with gold(I) complex as function of incubation time.

logðF 0 −F=F Þ ¼ n log KA þ n logð1=ð½1−f½ct  DNAðF 0 −F Þ=F 0 gÞ ð2Þ [43–45]. The extent of quenching of the fluorescence would reflect the extent of DNA binding of the second molecule. Fig. 7A shows the emission spectra of DNA–EB system upon increasing the amounts of complex 1. The emission intensity of the DNA–EB system decreased as the concentration of the gold–phosphine complex increased, which indicated that the complex could displace EB from the DNA–EB system. The result was due to the translocation of EB from a hydrophobic environment to an aqueous environment [38]. According to the Stern–Volmer equation, the quenching nature between DNA and complex 1 can be analyzed [44–46]: F 0 =F ¼ 1 þ Ksv ½1

ð1Þ

where KA and n are the binding constant and number of binding sites, and [1] and [ct-DNA] are the total complex 1 concentration and the total ct-DNA concentration, respectively. The linear fit according to Eq. (2), i.e. log(F0 − F) / F vs. log{1 / ([1] − [ct-DNA](F0 − F) / F0)}, is presented in Fig. 7C. The binding parameters, evaluated from the slope and intercept of these plots, are KA = 6.66 (±0.3)10 3 M−1 s−1 and n = 0.86. As it can be seen the value of intrinsic binding constant KA suggests low binding with DNA while the value of n about 1, suggesting that one molecule of complex 1 combines with one molecule of DNA in the drug to nucleic acid (d/DNA) molar ratio under study.

Fig. 5. Complex 1 causes DNA fragmentation, increases p53 expression and cytochrome c release. A) Gel electrophoresis of DNA fragments upon incubation of HCT-116 cells treated with IC50 concentration of complex 1 for 24 h at 37 °C. B) Western blotting for p53 expression vs β-actin used as control as well as cytosolic and mitochondrial expression of cytochrome c in HCT-116 cells lysate after treatment with IC50 and 2 × IC50 concentrations of complex 1. C) Densitometric analysis of band intensities. Results are reported as ratio between protein band and intensity of β-actin band.

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Fig. 7. Binding of Complex 1 to calf thymus DNA. A) Fluorescence emission spectra of EB–DNA in the absence (a) and in presence of increasing amounts of complex 1 (λex = 500 nm). B) Stern–Volmer plot of the fluorescence quenching of EB–DNA by complex 1. C) Plots for the determination of the binding parameters, according to Eq. (2). The experimental conditions are reported in Materials and methods.

Fig. 6. Complex 1 inhibits thioredoxin reductase activity and increases ROS production in colon cancer cells. A) TrxR activity in HCT-116 cells treated for 18 h with different concentrations of complex 1. B) hTrxR activity in untreated HCT-116 cell from which lysates were prepared and treated with different concentrations of complex 1. C) Evaluation of ROS levels in HCT-116 cells exposed to treatment with IC50 concentration of complex 1. (○) HCT-116 cells treated with IC50 concentration of Complex 1; (●) control.

3. Conclusion In this study, we synthesized the new mixed phosphine gold(I) complex [Au(tBu3P)(dppet)Cl] with high cytotoxic potential against HCT-116 human colon cancer cells. Even though the P NMR study shows that in solution the complex results in three species: [Au(tBu3P)(dppet)]+ itself,

[Au(tBu3P)2]+ and [Au(dppet)2]+ in this first approach we consider the total activity of the title compound. This compound was 85 fold more potent than cisplatin, suggesting its potential to be used as a therapeutic agent to overcome intrinsic cisplatin resistance. Treatment with complex 1 induced cell cycle arrest, which is consistent with previous data showing that this compound causes an accumulation of cells in the G0/G1 phase with a concomitant decrease in the G2/M population. Unlike cisplatin, the mechanism of cell cycle arrest did not involve interaction with DNA. There were only small effects on DNA conformation and stability, and the value of the binding constant obtained suggests a relatively weak binding to ct-DNA. The anticancer activity of complex 1 against HCT-116 cell line strongly supports a mechanism of action different from cisplatin. It has been previously shown that isolated mitochondria treated with auranofin and other gold compounds largely release cytochrome c [49], and our results agree with these and other findings [50]. Whereas cisplatin is effective in releasing cytochrome c only from sensitive cells [19,51,52]. This is because cisplatin acts directly on cellular DNA, while auranofin can specifically inhibit thioredoxin reductase [23,53]. Furthermore, it has been reported that the cytotoxicity of Au(I) phosphine compounds is mediated by their ability to inhibit mitochondrial human glutathione reductase

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(HGR) and TrxR irreversibly [16]. In particular, phosphol-containing gold(I) compounds are highly potent nanomolar inhibitors of both HGR and TrxR [7,16,54]. The enzyme TrxR is markedly expressed in tumor cell lines [55], and constitutes a target for cancer therapy [9,55–58]. Complex 1 potently inhibited TrxR in HCT-116 cells and a major consequence associated with this inhibition is the observed increase in ROS levels. Our data suggest that the redox metabolism and mitochondria are the likely cellular targets of complex 1. Considering that the decomposition of complex 1 in solution includes the formation of cations we can assert in our preliminary study that it could provide a combination of active species having effective antitumor activity as shown by the interesting results obtained for colon cancer cells but that we want to prove on a more large panel of tumors. Therefore, we are also working to perform a comparative study in solution for both species to determine their contribution on the cytotoxic activity observed. Similar behavior in solution was observed also for other complexes as Au(I) complex, chlorotriphenylphosphine-1,3-bis(diphenylphosphino)propanegold(I), [Au(DPPP)(PPh3)Cl] [32] that the 31P{ 1H} NMR spectrum in DMSO showed the existence of the following products: [Au(DPPP)(PPh3)Cl], [Au(DPPP)2]Cl, PPh3, and [Au(PPh3)2Cl]. In summary, as a consequence of their mechanism of action, targeting macromolecular synthesis, mitochondrial energy metabolism, and, to a minor extent, DNA, gold compounds may prove to be effective for the treatment of cisplatin-resistant cancers. Gold(I) complex, here reported and tested, effectively inhibits cell proliferation, induces apoptosis, and inhibits the growth of colon cancer cells and further confirms chelating diphosphines as a necessary feature for effective antitumor activity in their Au(I) derivatives. Given the poor long-term survival and prospects for patients who have late-stage colon cancer and the lack of successful treatments at this stage, this highly effective gold(I) complex could be an interesting therapeutic target against colon cancer. 4. Materials and methods Au(tBu3P)Cl, dppet, and all reagents were purchased from Aldrich (St. Louis, MO, USA) and used without further purification. The sample for microanalyses was dried in vacuo to constant weight (20 °C, ca. 0.1 Torr). Elemental analyses (C, H) were performed in-house with a Fisons Instruments 1108 CHNS-O elemental analyzer. IR spectra were recorded from 4000 to 600 cm−1 with a Perkin-Elmer Spectrum 100 FT-IR. 1H and 31P{1H} NMR spectra, referenced to Si(CH3)4 and external 85% H3PO4, respectively, were recorded on a 400 Mercury Plus Varian instrument operating at room temperature (400 MHz for 1H and 162.1 MHz for 31P). Peak multiplicities are abbreviated: singlet, s; doublet, d; triplet, t; multiplet, m. Melting points are uncorrected and were taken on an STMP3 Stuart scientific instrument and on a capillary apparatus. 4.1. Synthesis of complex [Au(tBu3P)(dppet)Cl] (1) Chloro[tris(tert-butyl)phosphine][cis-1,2-bis(diphenylphosphino) ethene]gold(I) have been synthesized upon addition of cis-1,2-bis (diphenylphosphino)ethene (0.079 g, 0.2 mmol) to a dichloromethane solution (10 ml) of chlorotris(tert-butyl)phosphinegold(I) (0.087 g, 0.2 mmol). The clear solution was stirred 4 h at room temperature, then the volume was reduced to one half and 5 ml of n-hexane were added. A colorless precipitate slowly formed, which was filtered off and the precipitate washed with diethyl ether (20 ml) to give 0.155 g (0.18 mmol, 90%) of the title compound. It is soluble in DMSO, chloroform, methanol, acetonitrile. M.p. 199 °C dec. Analysis calculated for C38H49AuClP3 (M.W. 831.15): C, 54.91; H, 5.94%. Found: C, 55.14, H, 5.76%. IR (cm−1): 3152w, 3053w ν(C–Harom), 1584 m, 1570w ν(C=C) 1481 m, 1433s 1395w, 1368w, 1306w, 1175m, 1094m, 1027w, 999w, 930w, 810w, 739s, 690vs. 1H NMR (CDCl3): δ, 1.53s, 1.56s, (27H, tBu3P), 7.12br (2H, CH=CHdppet), 7.35–7.70 (20H, m, C–Hdppet). 31P NMR

(CDCl3): δ, 23.6s (dppet of [Au(dppet)2] +), 43.7d (dppet of Au(tBu3P)(dppet)Cl, 2J(P–P) = 128 Hz), 97.4s (tBu3P of [Au(tBu3P)2]+), 104.9t (tBu3P of Au(tBu3P)(dppet)Cl, 2J(P–P) = 128 Hz). 1H NMR (DMSO-d6): δ, 1.49sbr, (27H, tBu3P), 7.93m (2H, CH = CHdppet), 7.15–7.38 (20H, m, C–Hdppet). 31P NMR (DMSO-d6): δ, 23.7s (dppet of [Au(dppet)2]+), 97.5s (tBu3P of [Au(tBu3P)2]+). ESI-MS (+, CH3CN) m/z (%) = 601 (100) [Au(tBu3P)2]+, 989 (20) [Au(dppet)2]+. Λm (DMSO, conc. 10−3 M): 44.3 S cm2 mol−1. Λm (CH2Cl2, conc. 10−3 M): 8.5 S cm2 mol−1. 4.2. Fluorescence quenching studies UV–Visible (UV–Vis) absorbance values were measured on a Perkin-Elmer Lambda-25 spectrophotometer in 10 mM phosphate buffer (pH 7.0) containing 50 mM NaCl at room temperature (25 °C). A stock solution of ct-DNA was prepared by dissolving the solid material in the same phosphate buffer. Solutions of DNA in the above buffer gave a ratio of UV absorbance at 260 and 280 nm, A260/A280 of 1.87, indicating that the DNA was sufficiently free of protein [35]. The concentration of DNA was determined by UV absorbance at 260 nm using the molar absorption coefficient ε 260 (6600 M−1 cm−1) [44,45]. The competitive binding experiment was carried out by maintaining the EB and ct-DNA concentration at 5 μM and 55.7 μM, respectively, while increasing the concentration of [Au(tBu3P)(dppet)Cl]. Fluorescence quenching spectra were recorded using a ISS-Greg 200 spectrofluorophotometer with an excitation wavelength of 500 nm and emission spectrum of 520–700 nm. Fitting was completed using GraphPad 4 software, where the value of the Stern–Volmer constant KSV was calculated. 4.3. Experiments with human colon cancer cell line Cisplatin and complex 1, along with the corresponding uncoordinated ligands, were dissolved in DMSO, just before the experiment, calculated amounts of drug solution were added to the growth medium to a final solvent concentration of 0.5%, which had no discernible effect on cell killing. MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), cis-platin, and all others materials were purchased from Sigma-Aldrich (St. Louis, MO, USA) and used without further purification. 4.4. Cell culture Human colon carcinoma cell line HCT-116 p53 wt were kindly provided by Dr. Carlos Galmarini (Institut National de la Sante et de la Recherche Medicale, Lyon, France). The HCT-116 cells were cultured in RPMI 1640 supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mmol/l glutamine and 10% heat-inactivated fetal bovine serum (HI-FBS). The cells were grown in a humidified atmosphere of 5% CO2 in air at 37 °C. Constituents and supplements of growth media were purchased from Sigma (St. Louis, MO, USA). 4.5. Viability assay Cytotoxicity assays were performed to establish the sensitivity of cancer cell lines and normal cell cultures to the experimental compounds. The growth inhibitory effect towards tumor cell lines was evaluated by means of MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay [59]. Briefly, cells were seeded in 96-well plates in 100 μl of growth medium (1 × 104 cells/ml). Cells were exposed to different concentrations of complex 1 (0.039–10 μM) at different time points. DMSO was utilized as vehicle control. Cell viability was then quantified by the ability of living cells to reduce the yellow dye MTT to an insoluble purple formazan dye crystals as product. At the end of the incubation, 10 μl MTT (5 mg/ml in phosphate buffer saline, PBS) per well was added and incubated for 4 h at 37 °C. The formazan product was then dissolved in 100 μl DMSO, after aspirating the medium. Absorbance was

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measured at 540 nm using a Titertek Multiscan microElisa (Labsystems, Helsinki, Finland). Production of the formazan is therefore proportional to living cells in a cell population. Cell viability was calculated as the percentage ratio of the sample absorbance with respect to the reference control. IC50 represents the concentration of the compound showing a lethal effect on 50% of the cells. Values were calculated using Graph Pad Prism 4 computer program (GraphPad Software, S. Diego, CA, USA). 4.6. Acridine orange and ethidium bromide staining Characteristic apoptotic or necrotic morphological changes were investigated by fluorescent microscopy using the acridine orange and ethidium bromide staining method. Cells were treated with IC50 concentrations of complex 1 for 2, 4, and 6 h. After incubation, HCT-116 cells were trypsinized and washed with phosphate buffered saline (PBS). The cells were resuspended at 1 × 106 cells/ml in PBS. The cell suspension (25 μl) was mixed with 5 μl of the fluorochrome mixture (100 μg/ml acridine orange and 100 μg/ml ethidium bromide in PBS) for 3 min, and 10 μl of the samples was examined under an Olympus IX71 fluorescent inverted microscope with appropriate combination of filters and recorded with an Olympus DP70 digital camera. Viable cells presented an intact, bright green nucleus. Early apoptotic cells contained a bright green nucleus, with condensed chromatin, whereas late apoptotic cells contained a red/orange nucleus showing chromatin condensation [60]. 4.7. Apoptosis studies 4.7.1. Flow cytometry analysis of DNA content Drug-induced cell cycle effects and DNA fragmentation were analyzed by flow cytometry after DNA staining with propidium iodide (PI) according to Nicoletti et al. [60]. Briefly, cells were seeded in 100 mm dishes at a density of 1.2 × 105 cells for each well. They were then incubated and allowed to grow to 50% confluence after which they were treated with IC50 and 2 × IC50 concentrations of complex 1 for 24 h. They were then harvested by trypsin release and washed twice with PBS. The pellet was suspended in an ice-cold solution of 70% ethanol in PBS (v/v). After 2 h of fixation, cells were washed twice with PBS. Fixed cells were stained by suspension in a solution of propidium iodide dye (100 μg/ml final concentration; Molecular probes, Eugene, OR) and 1% RNase A (Sigma Chemical Co, St. Louis, MO). Distribution of cell cycle phases with different DNA contents was determined using a FACScan flow cytometer (Becton-Dickinson, San Jose, CA). Cells that were less intensely stained than G1 cells (sub G1 cells) in flow cytometric histograms were considered apoptotic cells. Analysis of cell cycle distribution and the percentage of cells in the G1, S and G2/M phases of the cell cycle were determined using Cell Quest. 4.7.2. DNA fragmentation Conventional agarose gel electrophoresis was done as described previously with some modifications [55]. For DNA fragmentation assay, HCT-116 cells (1 × 10 6 cells) were grown in microtiter plates, then exposed to complex 1 at a final concentration of 0.5 μM. After 24 h of incubation, cells were washed with PBS, trypsinized, and centrifuged at 500 ×g for 5 min. The pellet was suspended in 20 μl of lysis buffer (50 mM Tris–HCl, pH 8.0, 10 mM EDTA, 0.5% SDS, and 0.5 mg/ml proteinase K) and incubated for 2 h at 50 °C. RNAse was added to the lysate to a final concentration of 10 μg/ml, and the resulting suspension was incubated for 2 h at 50 °C. DNA was then precipitated by adding 5 μl of 3 M sodium acetate (pH 5.2) and 100 μl of ice-cold 100% ethanol to the solution, incubated on ice for 10 min and collected by centrifugation at 10,000 ×g for 10 min. The pellet was dissolved in 15 μl of sterile distilled water and 2.5 μl of sample buffer (30% glycerol, 0.25% bromophenol blue). DNA samples were resolved on a 1.8% agarose gel using 0.5 × TBE (0.089 mM Tris, 0.089 mM borate, and 2.5 mM EDTA, pH 8.3) as the running buffer, and visualized under a UV transilluminator

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after staining with ethidium bromide (0.5 μg/ml). The gel was finally digitally photographed with a Gel-Pro Imager (Kodak MI). 4.7.3. Caspase-3 assay Activation of caspases-3 was measured by using colorimetric Caspase-3 Assay Kit (Sigma, St. Louis, MO, USA). Cells were incubated with IC50 concentration of complex 1 for different times (1, 2, 4, 6, 8 and 12 h), were harvested, washed with PBS and the pellets were resuspended in lysis buffer (50 μl) for 20 min. Lysates were centrifuged at 20,000 ×g for 10 min at 4 °C, and supernatants were collected and added into 96-well plates. Final reaction buffer (50 μl) and caspase-3 colorimetric substrate (Ac-DEVDpNA 5 μl) were then added to each well. Plates were incubated at 37 °C for 2 h and optical density was measured at 405 nm with a Titertek Multiscan microElisa (Labsystems, Helsinki). 4.7.4. Western blotting HCT-116 cells (1 × 10 6 cells/ml) were plated in 35-mm culture dishes and maintained at 37 °C in an incubator for 48 h. Next, the medium was replaced with serum-free DMEM medium for a further 4 h. After this step, the cells were treated either with or without complex 1 at concentrations corresponding to IC50 and 2 × IC50 for 24 h. The cells were rinsed twice with PBS after removing the medium and scraped in 300 μl of buffer [20 mM [4-(2-hydroxyethyl)piperazin1-yl]ethanesulfonic acid (HEPES) (pH 7.9), 10 mM KCl, 1 mM EDTA, 1 mM EGTA, 1.5 mM MgCl2, 1.0 mM dithiothreitol, 250 mM sucrose] containing a protease inhibitor cocktail. The cells were homogenized by ten strokes in a Dounce homogenizer (B. Braun, Melsungen, Germany). To collect nuclei and debris, the homogenates were centrifuged twice at 750 ×g for 5 min at 4 °C. The supernatants were centrifuged at 10,000 ×g for 15 min at 4 °C, to collect mitochondriaenriched heavy membrane pellets. The resulting supernatants were centrifuged at 100,000 ×g for 1 h at 4 °C and the final supernatants were referred to as cytosolic fractions. The amount of protein was measured using the Lowry method and the proteins were stored at −70 °C for further experiments. These sample proteins were separated in a 12% polyacrylamide mini-gel at 100 V for 2 h at room temperature using a Mini-PROTEIN II electrophoresis cell (Bio-Rad). After electrophoresis, the proteins were transferred overnight onto nitrocellulose membranes (Millipore, Bedford, MA, USA). The membranes were incubated for 1 h at room temperature in TBS-T solution [10 mM Tris–HCl (pH 7.6), 150 mM NaCl and 0.1% (v/v) Tween-20] containing 5% (w/v) nonfat dry milk (NFDM). The membranes were subsequently incubated for 2 h at room temperature with rabbit polyclonal antibodies [1:3000; p53 and cytochrome c, Santa Cruz Biotechnology, CA, USA] in TBS-T solution containing 5% NFDM. After three washes with TBS-T, the membranes were incubated for 1 h at room temperature with horseradish peroxidase-conjugated goat antirabbit IgG (1:10,000; Santa Cruz Biotechnology, CA, USA) in TBS-T containing 5% NFDM. Antibodies were then detected by chemiluminescence and auto radiography using X-ray film. β-Actin was used as a loading control. 4.7.5. ROS production The intracellular ROS production was measured by using nonfluorescent 2′-7′-dichlorodihydrofluorescein diacetate (DCFH-DA). This compound is deacetylated by intracellular esterases to the nonfluorescent DCFH, which is oxidized to the fluorescent compound DCF by ROS [61]. For the measurement of ROS production, HCT-116 cells were pre-incubated with 10 μM DCFH-DA for 30 min at 37 °C, and then the cells were washed twice with PBS to remove the excess DCFH-DA. After that, the cells were treated with or without complex 1 at concentration corresponding to IC50. Finally, the fluorescence of DCF (2′-7′-dichlorofluorescein) was measured at an excitation wavelength of 485 nm and an emission wavelength of 530 nm at different times (0, 30, 60, 120, and 240 min) with a thermostated and stirred-equipped Hitachi F-4500 spectrofluorophotometer. The

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values were calculated as a relative of the DCF fluorescence intensities, compared to the control. 4.8. Enzyme inhibition 4.8.1. Inhibition of thioredoxin reductase by gold (I) complex in cell lysate Thioredoxin reductase activity of complex 1 was evaluated in HCT116 cell lysate. The whole procedure was performed at 4 °C. HCT-116 cells were grown in 75 cm 2 flasks and detached with scraper. The collected cells were centrifuged, washed with PBS, and lysed with RIPA buffer modified as follows: 150 mM NaCl, 50 mM Tris–HCl, 1% Triton X-100, 1% SDS, 1% sodium deoxycholate, 1 mM NaF, 1 mM EDTA, and immediately before use, an anti-protease cocktail (Roche, Basel, Switzerland) containing PMSF was added. 0.080 mg of protein was incubated in the presence of different concentrations of gold(I) mixed with phosphine derivative. The assay was performed in 0.2 M Na, K-phosphate buffer (pH 7.4) containing 2 mM EDTA, 0.25 mM NADPH and about 0.05–0.1 mg of bovine serum albumin. The reaction was initiated by the addition of 3 mM DTNB (5,5′-dithiobis (2-nitrobenzoic acid)) to both sample and reference control and the increase of absorbance was monitored at 412 nm over 5 min at 25 °C. Final values were recorded and plotted as percent of enzyme activities relative to the control. 4.8.2. Thioredoxin reductase activity in treated cells HCT-116 cells were grown in 75 cm2 flasks to reach the 50% confluence and treated with complex 1 at different concentrations for 18 h. At the end of incubation time, cells were collected, washed with PBS and centrifuged. Each sample was then lysed with RIPA buffer modified as follows: 150 mM NaCl, 50 mM Tris–HCl, 1% Triton X-100, 1% sodium deoxycholate, 1 mM NaF, 1 mM EDTA, and immediately before use, an anti-protease cocktail (Roche, Basel, Switzerland) containing PMSF was added. Samples were tested for thioredoxin reductase (0.080 mg proteins) as described above. Abbreviations used CT-DNA calf-thymus DNA DCF 2′,7′-dichlorofluorescein DCFH-DA 2′-7′-Dichlorodihydrofluorescein diacetate DTNB 5,50-dithiobis (2-nitrobenzoic acid) dppe 1,2-bis(diphenylphosphino)ethane dppet bis(diphenylphosphino)ethane EB ethidium bromide HEPES 4-(2-hydroxyethyl)piperazinethanesulfonic acid HGR glutathione reductase MTT 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide PBS phosphate buffered saline PMSF phenylmethanesulfonylfluoride ROS reactive oxygen species tBu3P tris(tert-butyl)phosphine TrxR thioredoxin reductase Supplementary data to this article can be found online at http:// dx.doi.org/10.1016/j.jinorgbio.2013.03.014. Acknowledgment Funding from UNICAM is acknowledged. References [1] L. Ronconi, D. Fregona, Dalton Trans. (2009) 10670–10680. [2] B. Köberle, M.T. Tomicic, S. Usanova, B. Kaina, Biochim. Biophys. Acta 1806 (2010) 172–182.

[3] J.M. Piulats, L. Jiménez, X. García del Muro, A. Villanueva, F. Viñals, J.R. Germà-Lluch, Clin. Transl. Oncol. 11 (2009) 780–786. [4] S. Rafique, M. Idrees, A. Nasim, H. Akbar, A. Athar, Biotechnol. Mol. Biol. Rev. 5 (2010) 38–45. [5] P.J. O'Dwyer, J.P. Stevenson, S.W. Johnson, Cisplatin, in: B. Lippert (Ed.), chemistry and biochemistry of leading anticancer drug: Clinical status of cisplatin, carboplatin and other platinum-based antitumor drugs, Wiley-VCH, Zürich, 1999, pp. 31–72. [6] S. Berners-Price, in: E. Alessio (Ed.), Bioinorganic Medicinal Chemistry: Gold-Based Therapeutic Agents: A New Perspective, Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim, Germany, 2011. [7] I. Ott, R. Gust, Arch. Pharm. 340 (2007) 117–126. [8] P. Köpf-Maier, Eur. J. Clin. Pharmacol. 47 (1994) 1–16. [9] X. Wang, Z. Guo, Dalton Trans. (2008) 1521–1532. [10] I. Kostova, Anticancer Agents Med. Chem. 6 (2006) 19–32. [11] C. Marzano, V. Gandin, A. Folda, G. Scutari, A. Bindoli, M.P. Rigobello, Free Radic. Biol. Med. 42 (2007) 872–881. [12] K.P. Bhabak, G. Mugesh, Inorg. Chem. 48 (2009) 2449–2455. [13] C. Chiellini, A. Casini, O. Cochet, C. Gabbiani, G. Ailhaud, C. Dani, L. Messori, E.Z. Amri, Chem. Biodivers. 5 (2008) 1513–1520. [14] L. Oehninger, R. Rubbiani, I. Ott, Dalton Trans. 42 (2013) 3269–3284. [15] A. Casini, L. Messori, Curr. Top. Med. Chem. 11 (2011) 2647–2660. [16] M.J. McKeage, L. Maharaj, S.J. Berners-Price, Coord. Chem. Rev. 232 (2002) 127–135. [17] A. Casini, C. Hartinger, C. Gabbiani, E. Mini, P.J. Dyson, B.K. Keppler, L. Messori, J. Inorg. Biochem. 102 (2008) 564–575. [18] C.F. Shaw III, Chem. Rev. 99 (1999) 2589–2600. [19] J.C. Lima, L. Rodriguez, Anticancer Agents Med. Chem. 11 (2011) 921–928. [20] C.K. Mirabelli, R.K. Johnson, D.T. Hill, L.F. Faucette, G.R. Girard, G.Y. Kuo, C.M. Sung, S.T. Crooke, J. Med. Chem. 29 (1986) 218–223. [21] I. Ott, Coord. Chem. Rev. 253 (2009) 1670–1681. [22] P.J. Barnard, S.J. Berners-Price, Coord. Chem. Rev. 251 (2007) 1889–1902. [23] M.P. Rigobello, L. Messori, G. Marcon, M.A. Cinellu, M. Bragadin, A. Folda, G. Scutari, A. Bindoli, J. Inorg. Biochem. 98 (2004) 1634–1641. [24] R. Rubbiani, I. Kitanovic, H. Alborzinia, S. Can, A. Kitanovic, L.A. Onambele, M. Stefanopoulou, Y. Geldmacher, W.S. Sheldrick, G. Wolber, A. Prokop, S. Wölfl, I. Ott, J. Med. Chem. 53 (2010) 8608–8618. [25] K.F. Tonissen, G. Di Trapani, Mol. Nutr. Food Res. 53 (2009) 87–103. [26] A.B. Witte, K. Anestål, E. Jerremalm, H. Ehrsson, E.S. Arnér, Free Radic. Biol. Med. 39 (2005) 696–703. [27] E.S. Arnér, H. Nakamura, T. Sasada, J. Yodoi, A. Holmgren, G. Spyrou, Free Radic. Biol. Med. 31 (2001) 1170–1178. [28] K. Becker, C. Herold-Mende, J.J. Park, G. Lowe, R.H. Schirmer, J. Med. Chem. 44 (2001) 2784–2792. [29] E. Vergara, A. Casini, F. Sorrentino, O. Zava, E. Cerrada, M.P. Rigobello, A. Bindoli, M. Laguna, P.J. Dyson, ChemMedChem 5 (2010) 96–102. [30] M. Colonnello, E. Mini, B. Caciagli, M.A. Cinellu, A. Bindoli, C. Gabbiani, L. Messori, J. Med. Chem. 48 (2005) 6761–6765. [31] F. Caruso, M. Rossi, J. Tanski, C. Pettinari, F. Marchetti, J. Med. Chem. 46 (2003) 1737–1742. [32] F. Caruso, C. Pettinari, F. Paduano, R. Villa, F. Marchetti, E. Monti, M. Rossi, J. Med. Chem. 51 (2008) 1584–1591. [33] W.J. Geary, Coord. Chem. Rev. 7 (1971) 81–122. [34] P. Diversi, A. Cuzzola, F. Ghiotto, Eur. J. Inorg. Chem. (2009) 545–553. [35] M.P. Rigobello, A. Folda, B. Dani, R. Menabò, G. Scutari, A. Bindoli, Eur. J. Pharmacol. 582 (2008) 26–34. [36] O. Rackham, S.J. Nichols, P.J. Leedman, S.J. Berners-Price, A.A. Filipovska, Biochem. Pharmacol. 74 (2007) 992–1002. [37] S. Kagawa, J. Gu, T. Honda, J.T. McDonnell, G.S. Swisher, A.J. Roth, B. Fang, Clin. Cancer Res. 7 (2001) 1474–1480. [38] A.J. Sánchez-Alcázar, A. Khodjakov, E. Schneider, Cancer Res. 61 (2001) 1038–1044. [39] Y. Wang, M. Liu, R. Cao, M. Yin, Q. Liu, N. Huang, J. Med. Chem. 56 (2013) 1455–1466. [40] M.J. Matês, M.F. Sânchez-Jiménez, Int. J. Biochem. Cell Biol. 32 (2000) 157–170. [41] S. Nafisi, Z. Norouzi, DNA Cell Biol. 28 (2009) 469–477. [42] N. Ercal, H.N. Gurer-Orhan, N. Aykin-Burns, Curr. Top. Med. Chem. 1 (2001) 529–539. [43] M. Schmid, S. Zimmermann, H.F. Krug, B. Sures, Environ. Int. 33 (2007) 385–390. [44] R. Bera, B.K. Sahoo, K.S. Ghosh, S. Dasgupta, Int. J. Biol. Macromol. 42 (2008) 14–21. [45] N. Akbay, Z. Seferoğlu, E. Gök, J. Fluoresc. 19 (2009) 1045–1051. [46] A. Alonso, M.J. Almendral, Y. Curto, J.J. Criado, E. Rodrıguez, J.L. Manzano, Anal. Biochem. 355 (2006) 157–164. [47] J.R. Lakowicz, Principles of Fluorescence Spectroscopy, 2nd ed. Kluwer Academic/ Plenum Publishers, New York, 1999. [48] G. Lupidi, E. Camaioni, H. Khalifé, L. Avenali, E. Damiani, F. Tanfani, A. Scirè, J. Pharm. Sci. 101 (2012) 2564–2573. [49] M.P. Rigobello, G. Scutari, R. Boscolo, A. Bindoli, Br. J. Pharmacol. 136 (2002) 1162–1168. [50] K.M. Debatin, D. Poncet, G. Kroemer, Oncogene 21 (2002) 8786–8803. [51] S. Tanida, T. Tsutomu Mizoshita, K. Ozeki, H. Tsukamoto, T. Kamiya, H. Kataoka, D. Sakamuro, T. Joh, Int. J. Surg. Oncol. (2012) 1–8. [52] A.M. Florea, D. Büsselberg, Cancers 3 (2011) 1351–1371. [53] K. Hill, G.W. McCollum, M.E. Boeglin, R.F. Burk, Biochem. Biophys. Res. Commun. 234 (1997) 293–295.

G. Lupidi et al. / Journal of Inorganic Biochemistry 124 (2013) 78–87 [54] S. Gromer, L.D. Arscott, C.H. Williams Jr., R.H. Schirmer, K. Becker, J. Biol. Chem. 273 (1998) 20096–20101. [55] M.P. Rigobello, G. Scutari, A. Folda, A. Bindoli, Biochem. Pharmacol. 67 (2004) 689–696. [56] D.T. Lincon, E.M. Ali Emadi, K.F. Tonissen, F.M. Clarke, Anticancer. Res. 23 (2003) 2425–2433. [57] M. Deponte, S. Urig, L.D. Arscott, K. Fritz-Wolf, R. Reau, H.-M.C. Christel, S. Koncarevic, M. Meyer, E. Davioud-Charvet, P.D. Ballou, H.C. Williams Jr., K. Becker, J. Biol. Chem. 280 (2005) 20628–20637.

87

[58] Y. Liu, Y. Li, G. Zhao, Curr. Drug Targets 13 (2012) 1432–1444. [59] M. Page, N. Bejaoui, B. Cinq-Mars, P. Lemieux, Int. J. Immunopharmacol. 10 (1988) 785–793. [60] I. Nicoletti, G. Migliorati, M.C. Pagliacci, J. Immunol. Methods 139 (1991) 271–279. [61] C.P. LeBel, H. Ischiropoulos, S.C. Bondy, Chem. Res. Toxicol. 5 (1992) 227–231.