Free Radical Research, January 2015; 49(1): 67–77 © 2014 Informa UK, Ltd. ISSN 1071-5762 print/ISSN 1029-2470 online DOI: 10.3109/10715762.2014.979168
ORIGINAL ARTICLE
A new low molecular weight, MnII-containing scavenger of superoxide anion protects cardiac muscle cells from hypoxia/reoxygenation injury S. Nistri1, G. Boccalini1, A. Bencini2, M. Becatti3, B. Valtancoli2, L. Conti2, L. Lucarini4 & D. Bani1 1Department
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of Experimental & Clinical Medicine, Section of Anatomy & Histology, Research Unit of Histology & Embryology, University of Florence, Florence, Italy, 2Department of Chemistry, University of Florence, Florence, Italy, 3Department of Experimental & Clinical Biomedical Sciences “Mario Serio”, University of Florence, Florence, Italy, and 4Department of Neurosciences and Psychology, Drug Area and Child’s Health, Section of Pharmacology, University of Florence, Florence, Italy Abstract Reperfusion injury after oxygen starvation is a key pathogenic step in ischemic diseases. It mainly consists in oxidative stress, related to mitochondrial derangement and enhanced generation of reactive oxygen species (ROS), mainly superoxide anion (O2•2), and peroxynitrite by cells exposed to hypoxia. This in vitro study evaluates whether MnII(4,10-dimethyl-1,4,7,10-tetraazacyclododecane-1,7-diacetate).2H2O, or MnII(Me2DO2A), a new low molecular weight, MnII-containing O2• scavenger, has a direct protective action on H9c2 rat cardiac muscle cells subjected to hypoxia and reoxygenation. MnII(Me2DO2A) (1 and 10 μmol/l) was added to the culture medium at reoxygenation and maintained for 2 h. In parallel experiments, the inactive congener ZnII(Me2DO2A), in which ZnII replaced the functional MnII center in the same organic scaffold, was used as negative control. MnII(Me2DO2A) (10 μmol/l) significantly increased cardiac muscle cell viability (trypan blue assay), improved mitochondrial activity (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide test, membrane potential Δψ), reduced apoptosis (mitochondrial permeability transition pore opening, caspase-3, terminal deoxynucleotidyl transferase deoxyuridine triphosphate nick end labeling assay), decreased intracellular ROS levels (2¢,7¢-dichlorodihydrofluorescein diacetate and MitoSOX assays), and decreased protein nitroxidation (nitrotyrosine [NT] expression) and DNA oxidation (8-hydroxy-deoxyguanosine levels). Of note, ZnII(Me2DO2A) had no protective effect. The mechanism of MnII(Me2DO2A) relies on concentration-dependent removal of harmful O2• generated at reoxygenation from dysfunctional mitochondria in hypoxia-induced cells, as indicated by the MitoSOX assay. This study suggests that MnII(Me2DO2A) is a promising antioxidant drug capable of reducing O2•-mediated cell oxidative stress which occurs at reoxygenation after hypoxia. In perspective, MnII(Me2DO2A) might be used to reduce ischemia–reperfusion organ damage in acute vascular diseases, as well as to extend the viability of explanted organs before transplantation. Keywords: superoxide anion, ROS scavenger, H9c2 cardiac muscle cells, hypoxia, reoxygenation, oxidative stress
Introduction During oxidative metabolism, at different points in electron transport chain, O2 is reduced through one-electron paths to superoxide anion (O2•), a highly reactive radical which can oxidize and functionally compromise cell’s membrane lipids, proteins, and nucleic acids[1,2]. Normally, O2• levels are kept under the damage threshold by endogenous antioxidants, among which a key role is played by superoxide dismutases (SODs), which catalyze the dismutation of O2• to O2 and H2O2 by a reaction requiring a transition metal at the catalytic site [3–5]. However, oxidative stress causes a rapid inactivation of SODs, sparkling a vicious cycle that causes ever-increasing, harmful O2• tissue levels [6,7]. A paradigm of oxidative stress-related diseases includes ischemia–reperfusion, as it occurs during acute myocardial infarction and stroke [8]. In this process, the initial ischemic damage is then compounded by additional and more severe injury caused by reoxygenation upon blood flow restoration. Multiple interplaying mechanisms play a role in reperfusion-induced damage
and mainly involve oxidative stress [9]. In fact, mitochondrial dysfunction of hypoxia-exposed cells impairs the electron flow and enhances O2• formation and release [10,11]. Moreover, mitochondria can use the respiratory chain to reduce nitrite (NO2⫺) to nitric oxide (NO•) [12]. Because cytochrome oxidase produces NO• from nitrite at low O2 concentrations, the mitochondrially generated oxidants whose concentration increase under hypoxic conditions also include peroxynitrite (ONOO⫺), which is formed by the reaction between O2• and NO• [13]. In vivo, these mitochondrial mechanisms intrinsic to hypoxic cells are paralleled by inflammatory cell recruitment secondary to endothelial injury [14], which results in further enhancement of the generation of ONOO⫺. Over the past decade, ONOO⫺ has been identified as the effector of most, if not all, detrimental effects of excess NO• [15,16] and is regarded as the major harmful oxidant in myocardial hypoxia–reoxygenation injury [17]. The above notions underscore that O2• and NO• play a pivotal role in oxidative stress upon hypoxia–reoxygenation and hence can be suitable target for new pharmacological
Correspondence: Silvia Nistri, PhD, DSc, Department of Experimental and Clinical Medicine, Section of Anatomy and Histology, Research Unit of Histology and Embryology, University of Florence, viale G. Pieraccini, 6. I-50139 Florence, Italy.Tel: ⫹ 39 055 4271 387. Fax: ⫹ 39 055 4271 385. E-mail: silvia.nistri@unifi.it (Received date: 16 July 2014; Accepted date: 17 October 2014; Published online: 13 November 2014)
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68 S. Nistri et al. interventions. Theoretically, removal of excess O2• is the best approach to reduce oxidative stress, while leaving enough NO• to sustain residual vascular function [18]. In this context, extractive or recombinant SOD appears to be the logical choice. Indeed, transfection of H9c2 rat cardiomyocytes with Cu/Zn SOD was found to attenuate hypoxia–reoxygenation injury [19]. However, the pharmaceutical use of SOD is limited by its poor stability in water and intracellular penetration, immunogenicity, short half-life, and unfavorable yield/cost ratio [20,21]. Because of these limitations, pharmacological research has diverged toward non-peptidic, low molecular weight, Mn-containing compounds capable of efficiently catalyzing O2• dismutation as do authentic SODs [22]. Their efficacy relies on the wellknown chemical property of Mn ion to bind to and react with free radicals, including O2•. Several MnII and MnIII compounds with organic scaffolds, in particular, MnIII complexes with porphyrins, salen and its derivatives, and MnII chelates with cyclic polyamines and polyamine–polycarboxylates have been synthesized as reactive oxygen species (ROS) scavengers and tested for therapeutic properties in cellular and animal models of oxidative stress [23,24]. In recent years, we investigated the chemical and biological properties of a new polyamine–polycarboxylate MnII complex, MnII(4,10-dimethyl-1,4,7,10-tetraazacyclododecane-1,7-diacetate), also termed “MnII(Me2DO2A),” effective as O2• scavenger [25]. In principle, polyamine– polycarboxylate scaffolds represent an optimal tool for the synthesis of highly stable MnII complexes. In fact, these molecules are known for their ability to strongly bind a number of metals, from alkali and alkali earths to lanthanide and transition metal cations, including MnII. Moreover, these compounds are resistant to oxidizing and reducing agents, are highly soluble in aqueous media, and exhibit low toxicity and all properties that render them suitable for several biological and medicinal applications [26,27]. In particular, MnII(Me2DO2A) effectively reduced oxidative stress in cell culture models and oxidative tissue injury in animal models of inflammation [25,28,29]. The current study was designed to test the antioxidant effects of MnII(Me2DO2A) using H9c2 cardiac muscle cells subjected to hypoxia–reoxygenation in vitro. We used the inactive congener ZnII(Me2DO2A), in which the functional MnII was replaced with a ZnII ion lacking the capability to catalyze O2•decomposition, as control. Materials and methods Reagents The amount of MnII(Me2DO2A) required to perform the present experiments was kindly donated by the patent owner General Project Ltd., Montespertoli (Florence), Italy. The inactive compound ZnII(Me2DO2A) was synthesized in our laboratory from Me2DO2A following the same procedure as MnII(Me2DO2A). The formation constants of the ZnII complexes have been determined using the same procedure reported for the MnII complexes [25].
Unless otherwise specified, the other reagents used for the experiments were from Sigma-Aldrich (Milan, Italy). Chemical characterization of MnII(Me2DO2A) and ZnII(Me2DO2A) The compound MnII(Me2DO2A) was synthesized in our laboratories and patented [30]. The organic scaffold contains a tetraamine macrocyclic moiety, capable of forming complexes with transition metals characterized by high thermodynamic stability, kinetic inertness, and poor tendency to bind alkali or alkaline metal cations [31]. It also possesses two carboxylic moieties as pendant arms, acting as additional binding sites for metal cations. Like most polyamine–carboxylate ligands, Me2DO2A is able to form stable complexes with alkali and alkaline-earth metal cations. Table I reports the stability constants of its complexes with MnII in comparison with MgII, CaII, and NaI (the constant of the complex with KI is very low—log K ⬍ 2—and cannot be reliably determined). Table I shows that MnII forms the most stable complex with the ligand in its fully deprotonated form, where the deprotonated carboxylic groups are likely coordinated with the metal center (Scheme 2), yielding the MnII(Me2DO2A) neutral complex (Figure 1). To rule out the possibility of exchange reactions with MgII, CaII, and NaI, whose intracellular concentrations are three or more orders of magnitude higher than MnII, competition experiments were carried out with Me2DO2A and MnII at very low concentrations (1 μM) in the presence of excess NaI (165 mM), CaII (2.5 mM), and MgII (1.2 mM) [25]. The competition diagram reported in Figure 2A shows that MnII remains completely bound to Me2DO2A, while CaII, MgII, and NaI appeared unbound by the ligand. Therefore, the MnII(Me2DO2A) complex is substantially unaffected by other metal ion competitors in the medium, and no detectable MnII release occurs because of complexation with other metals. At variance with MgII, CaII, and NaI, the MnII(Me2DO2A) complex is also able to bind acidic protons to give protonated complexes of the type MnII(HMe2DO2A)⫹ and MnII(H2Me2DO2A)2⫹ in water. However, the proton affinity of these MnII(Me2DO2A) species is rather weak and only minor amounts of the protonated complexes are formed at acidic pH values. In fact, proton binding would Table I. Stability constants of the NaI, MgII, CaII, MnII, and ZnII complexes with H2Me2DO2A (H2L) and its deprotonated forms— (HMe2DO2A) ⫺ (HL⫺) and (Me2DO2A)2 (L2⫺) (I ⫽ 0.1 M, 25°C). Reaction Na⫹ ⫹ L2⫺ ⫽ [NaL] ⫺ Ca2⫹ ⫹ L2⫺ ⫽ [CaL] Mg2⫹ ⫹ L2⫺ ⫽ [MgL] Mn2⫹ ⫹ L2⫺ ⫽ [MnL] [MnL] ⫹ H⫹⫺ ⫽ [Mn(HL)]⫹ [Mn(HL)]⫹ ⫹ H⫹⫺ ⫽ [Mn(H2L)]2⫹ Zn2⫹⫺ ⫹ L2⫺ ⫽ [ZnL] [ZnL] ⫹ H⫹ ⫽ [Zn(HL)]⫹ [Zn(HL)]⫹ ⫹ H⫹ ⫽ [Zn(H2L)]2⫹
Log K 2.08 (7) 7.67 (8) 5.74 (7) 14.73 (2) 4.53 (7) 3.96 (2) 15.91 (1) 4.25 (1) 3.90 (1)
Superoxide scavenger and H/R cardiac cells
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Figure 1. Polyamine–polycarboxylate scaffold (A) and 3D structure (B) of MnII(Me2DO2A).
imply detachment of the coordinated carboxylate group from the metal center, which is an unlikely and unfavored event. As a matter of fact, as shown in Figure 2B, the neutral MnII(Me2DO2A) complex can be regarded as the only species present in solution at neutral pH value. The stability of MnII(Me2DO2A) is also higher than that of other MnII complexes with endogenous ligands present
Figure 2. (A) Overall percentages of the species complexed with Mn(II), Ca(II), Mg(II), and Na(I) as a function of pH in competitive systems containing Mn(II) (1 μM), Ca(II), (2.5 mM), Mg(II) (1.5 mM), and Na(I) (165 mM), and Me2DO2A (1 μM). ∑[Mn(HxL)] ⫻ ⫹ ⫽ [MnL] ⫹ [Mn(HL)] ⫹ ⫹ [Mn(H2L)]2 ⫹; ∑[HxL4](x ⫺ 2) ⫹ ⫽ [L4]2 ⫺ ⫹ [HL4] ⫺ ⫹ [H2L4] ⫹ [H3L4] ⫹ ⫹ [H4L4]2 ⫹; (B) distribution curves of the complexes with Me2DO2A (25°C, I ⫽ 0.1 M). At physiological pH, MnII(Me2DO2A) does not release MnII, even in the presence of other metal cations; at pH: 6, MnII release is less than 5%.
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in the cellular environment, such as ATP (log K ⫽ 4.66 for the equilibrium MnII ⫹ ATP ⫽ [MnII(ATP)]) [32], glutathione (logK ⫽ 2.7 for MnII ⫹ GSH ⫽ [MnII(GSH)]) [33], and carboxylate anions (logK ⫽ 3.79, 1.68, and 0.92 for MnII complexation by citrate [34], maleate [35], and Llactate anions [36]), thus excluding substantial de-metallation of MnII(Me2DO2A) by biological chelators. MnII (Me2DO2A) was rather resistant to electrochemical oxidation in water solution, displaying an onset oxidation potential of 0.86 V and a formal E1/2 potential of 0.85 V [25]. MnII oxidation is characterized by a fast single-electron transfer process ([MnIIL] → [MnIIIL]⫹) followed by slow rearrangement of the carboxylate groups around MnIII. In our previous study, we demonstrated that MnII(Me2DO2A) had strong O2• scavenging properties in vitro and in vivo, with a kcat value of 1.1 ⫻ 106 M⫺ 1 s⫺ 1 [25,28]. The structural characteristics of MnII(Me2DO2A) resemble, in part, those of the M40403 compound, where MnII is bound by a pentaamine macrocycle to afford a positively charged complex. However, the catalytic ability in O2• dismutation of MnII(Me2DO2A) is about one order of magnitude lower than M40403, the kcat of the latter in water being 2 ⫻ 107 M⫺ 1 s⫺ 1 [8]. At variance with M40403, MnII(Me2DO2A) is a neutral complex and the metal cation is more coordinatively saturated and shielded by the simultaneous coordination of the tetraamine macrocycle and the two carboxylate anionic moieties, making it less accessible to the O2• anion in the first step of the catalytic process. On the other hand, binding of the carboxylate groups makes MnII(Me2DO2A) thermodynamically stable and, therefore, resistant to possible metal release. Moreover, the neutral charge of MnII(Me2DO2A) allows it to cross the cell membranes and attain significant intracellular concentrations [25]. Moreover, the presence of two methyl groups appended to amine groups prevents the nitrogen atoms to interact via hydrogen bonding with water molecules, and enhances the lipophilic features of the complex. Interestingly, the absence of negative charge and increased lipophilicity has been shown to facilitate accumulation of manganese complexes in mitochondria [24, 37]. This may suggest that MnII(Me2DO2A) can directly scavenge O2• in mitochondria. At the same time, the amine groups at positions 4 and 10 can be easily functionalized with other organic moieties: this property may be exploited to increase the lipophilic characteristics of the compounds by substituting the methyl groups with hydrophobic alkyl or aryl moieties. Me2DO2A displays a binding ability for ZnII similar to that observed for MnII. As shown in Table I, in aqueous solution this ligand forms a neutral ZnII(Me2DO2A) complex that is slightly more stable than the MnII complex. Similar to MnII(Me2DO2A), in ZnII(Me2DO2A) the metal is likely coordinated by both the tetraamine scaffold and the two carboxylate groups. Protonation of the carboxylate groups occurs at acidic pH values, generating ZnII(HMe2DO2A)⫹ and ZnII(H2Me2DO2A)2⫹. As for MnII(Me2DO2A), these features ensure that neither metal release at neutral or slightly acidic pH values nor
70 S. Nistri et al. transmetallation reaction can occur in solution. From an electrochemical point of view, the ZnII ion is extremely stable to reduction; the reduction potential of the ZnII/ Zn0 couple being ⫺ 0.76 V. At the same time, oxidation states higher than II are not accessible for zinc [38]. These characteristics make this complex inert to reducing and oxidizing agents present in cellular environment, including O2• or other ROS.
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Cell culture and treatments H9c2 embryonic rat cardiac muscle cells, obtained from European Collection of Cell Cultures (ECACC, Salisbury, UK), were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Invitrogen, Carlsbad, CA, USA), 2 mM of glutamine, 250 U/ml of penicillin G, and 250 μg/ml of streptomycin, in a humidified atmosphere with 5% CO2 at 37°C. These cells were subjected to hypoxia and reoxygenation (H ⫹ R), simulated in vitro by substrate starvation plus hypoxia followed by reoxygenation, as previously described [39] with minor modifications. The cells were incubated in DMEM with no serum or glucose and placed in a hypoxic chamber saturated with a 0.1% O2, 5% CO2, and ≈ 95% N2 gaseous mix, humidified, and warmed at 37°C, for 7 h. At the end of hypoxia, the cells were reoxygenated for 2 h by incubation in normoxic conditions in glucose-containing, serum-free DMEM. Normoxic control cultures were also prepared. Cells were treated with or without MnII(Me2DO2A) at 2 different doses (1 and 10 μmol/l) and added at reoxygenation, concurrently with the peak of ROS generation (H ⫹ MnII(Me2DO2A)⫹ R). The given MnII(Me2DO2A) doses were chosen on the basis of our previous in vitro studies [25]. As control for the specific capability of MnII(Me2DO2A) in suppressing oxidative stress by redox reaction, some cell viability experiments were performed using the inactive congener ZnII(Me2DO2A) at the same concentrations as MnII(Me2DO2A) added at reoxygenation (H ⫹ ZnII(Me2DO2A)⫹ R). Separate experiments were performed to assess the toxicity of MnII(Me2DO2A) and ZnII(Me2DO2A) in H9c2 cells. In these experiments, the two compounds were added to the cell cultures at increasing concentrations (0.1–100 μmol/l) for 2 and 24 h; cell viability was then assayed by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) test. Trypan blue viability assay The trypan blue exclusion method was used to assess cell viability. H9c2 cells (5 ⫻ 104/well) were seeded in 24-well plates. At the end of the treatments, the cells were gently detached by trypsin/ethylenediaminetetraacetic acid, resuspended in culture medium, and mixed 1:1 with 0.4% trypan blue solution. The final cell suspensions were counted under a phase contrast inverted microscope using a Burker chamber. Viable cells were expressed as percentage of the total counted cells.
MTT assay Cell mitochondrial function was measured using the MTT assay. H9c2 cells (5 ⫻ 104/well) were seeded in 24-well plates. At the end of the treatments, the culture medium was removed from each well and replaced with 300 μl of MTT stock solution, followed by 4-h incubation at 37°C. Then, 350 μl of dimethyl sulfoxide were added to each well to dissolve the formazan crystals. The plate was gently shaken for 10 min and read at 550 nm on a plate reader. Optical density was assumed as indicator of mitochondrial activity and, indirectly, cell viability. Mitochondrial membrane potential (Δψ) Mitochondrial membrane potential was assessed using tetramethylrhodamine methyl ester perchlorate (TMRM), a lipophilic potentiometric fluorescent dye that distributes between the mitochondria and cytosol in proportion to Δψ by virtue of its positive charge. At low concentrations, the fluorescence intensity depends on dye accumulation in mitochondria, which in turn is directly related to mitochondrial potential. For confocal microscope analysis, cells were cultured on glass coverslips and loaded for 20 min at 37°C with TMRM, dissolved in 0.1% dimethyl sulfoxide (DMSO) to a final concentration of 100 nM in the culture medium. The cells were fixed in 2% buffered paraformaldehyde for 10 min at room temperature and the TMRM fluorescence was analyzed under a confocal Leica TCS SP5 scanning microscope (Mannheim, Germany) equipped with a helium–neon laser source, using a 543-nm excitation wavelength, and with a Leica Plan Apo x63 oil immersion objective. Mitochondrial membrane potential was also quantified by flow cytometry. Single-cell suspensions were washed twice with PBS and incubated for 20 min at 37°C in the dark with TMRM dissolved in DMEM (100 nM). The cells were then washed, resuspended in PBS, and analyzed using a FACSCanto flow cytometer (Becton–Dickinson, San Jose, CA). Mitochondrial permeability transition pore opening Mitochondrial permeability, an index of mitochondrial dysfunction and early apoptosis, was measured by calcein fluorescence, as described [40]. The fluorescent probe calcein-(acetoxymethyl) AM freely enters the cells and emits fluorescence upon de-esterification. Coloading of cells with cobalt chloride, which cannot cross the mitochondrial membranes in living cells, quenches the fluorescence in the whole cell except mitochondria. During induction of mPTP, cobalt can enter mitochondria and quench calcein fluorescence, whose decrease can be taken as a measure of the extent of mPTP induction. H9c2 cells grown on glass coverslips were loaded with calcein-AM (3 μM) and cobalt chloride (1 mM) added to the culture medium for 20 min at 37°C. The cells were then washed in PBS, fixed in 2% buffered paraformaldehyde for 10 min at room temperature, and analyzed by a Leica TCS SP5 confocal laser scanning microscope equipped with an
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argon laser source, using 488-nm excitation wavelength, and with a Leica Plan Apo x63 oil immersion objective. Mitochondrial permeability was also monitored by flow cytometry: single-cell suspensions were incubated with calcein-AM (3 μM) and cobalt chloride (1 mM) for 20 min at 37°C, washed twice with PBS, and analyzed using a FACSCanto flow cytometer (Becton–Dickinson).
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Assessment of caspase-3 activity H9c2 cells seeded on glass coverslips were incubated with FAM FLICA™ Caspase 3&7 assay kit (Immunochemistry Technologies, Bloomington, MN, USA) for 30 min, following the manufacturer’s instructions. After incubation, the cells were thoroughly washed and fixed in 2% buffered paraformaldehyde for 10 min at room temperature. Fluorescence was detected by a confocal Leica TCS SP5 scanning microscope equipped with an argon laser source, using 488-nm excitation wavelength, and a Leica Plan Apo x63 oil immersion objective. Caspase-3 activity was also quantified by flow cytometry: single-cell suspensions were incubated with FAMFLICA™ for 30 min at 37°C, washed twice with PBS, and analyzed using a FACSCanto flow cytometer (Becton– Dickinson). TUNEL assay H9c2 cells were grown on glass coverslip and subjected to the different treatments. Cell death was studied with TUNEL assay for apoptosis, performed using a KlenowFragELTM DNA fragmentation detection kit (Calbiochem,
Figure 3. Evaluation of H9c2 cell viability by trypan blue assay. Reoxygenation (H ⫹ R) causes a marked reduction of the amounts of viable cells. This effect was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R), while ZnII(Me2DO2A) had no effects (H ⫹ Zn ⫹ R). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
Figure 4. Evaluation of H9c2 cell mitochondrial integrity and function by MTT assay, which reveals the efficiency of the respiratory chain (A), and TMRM assay, which evaluates the mitochondrial membrane potential, by confocal microscopy (B) and FACS analysis (C). Reoxygenation (H ⫹ R) causes a marked mitochondrial impairment, which was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). ZnII(Me2DO2A) (H ⫹ Zn ⫹ R) had no effects (A). FACS analysis confirms the microscopic findings of TMRM fluorescence as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose. Scale bars: 20 μm. Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
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Figure 5. Evaluation of apoptosis in H9c2 cells by mPTP and caspase-3 activity assayed by confocal microscopy (A) and FACS analysis (B). Reoxygenation (H ⫹ R) causes the extinction of calcein-AM fluorescence in mitochondria by increased cobalt ion permeation, while this effect was dose-dependently inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Similarly, cytoplasmic fluorescence related to activated caspase-3 was increased by H ⫹ R and markedly decreased by MnII(Me2DO2A). Scale bars: 20 μm. FACS analysis confirms the microscopic findings as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose.
San Diego, CA, USA), as reported in the manufacturer’s instructions. Briefly, H9c2 cells were incubated with 15 mg/ml of proteinase K for 5 min at room temperature. After rinsing in PBS, the cells were immersed in the Klenow Labeling Reaction Mixture containing deoxynucleotidyl transferase and unlabelled and biotin-labeled deoxynucleotides, and incubated at 37° C for 90 min in a humid atmosphere. Then, the cells were incubated with peroxidase-conjugated streptavidin for 30 min at room temperature and the signal was revealed with 3,3′diaminobenzidine. Finally, nuclear counterstaining was achieved using methyl green. Apoptotic nuclei were recognized by the presence of dark brown staining, at variance with those of viable cells, which instead appeared pale brown or green. TUNEL-positive nuclei were counted in 10 microscopic fields for each cell preparation. TUNEL apoptotic index was then expressed as relative percentage of TUNEL-positive nuclei on the total number of methylgreen-stained nuclei. Determination of intracellular ROS and mitochondrial superoxide H9c2 cells seeded on glass coverslips were loaded with the ROS-sensitive fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA; Invitrogen, CA, USA; 2.5 μmol/l) or the mitochondrial O2•-specific fluorescent probe MitoSOX (Invitrogen; 3 μmol/l)—dissolved in 0.1% DMSO and Pluronic acid F-127 (0.01% w/v)—which were
added to cell culture media for 15 min at 37°C. The cells were fixed in 2% buffered paraformaldehyde for 10 min at room temperature, and the H2DCFDA and MitoSOX fluorescence were analyzed using a Leica TCS SP5 confocal scanning microscope equipped with an argon laser source, using 488-nm and 543-nm excitation wavelength, respectively, and a Leica Plan Apo x63 oil immersion objective. ROS and mitochondrial O2• generation were also monitored by flow cytometry: briefly, single-cell suspensions were incubated with H2DCFDA (1 μmol/l) or MitoSOX (0.5 μmol/l) for 15 min at 37°C and immediately analyzed using a FACSCanto flow cytometer (Becton–Dickinson). Data were analyzed using FACSDiva software (Becton– Dickinson). Immunohistochemical localization and quantitation of nitrotyrosine NT, an index of protein nitrosylation by harmful ONOO⫺ generated during inflammation, was determined by immunocytochemistry as described previously [41]. Briefly, H9c2 cells were grown on glass coverslip and subjected to the different treatments. The cells were fixed with formaldehyde for 10 min and then incubated with rabbit polyclonal anti-NT antibody (Upstate Biotechnology, Buckingham, UK; 1:118) at 4°C overnight. Immune reaction was revealed by goat anti-rabbit IgG conjugated with biotin (1:200; Vector Lab, Burlingame, CA, USA) followed by incubation with
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the treatments, the cells were collected with TRIzol Reagent (Invitrogen) and DNA was isolated according to the manufacturer’s instructions. Then DNA was subjected to enzymatic digestion with 10 IU of P1 nuclease (Sigma-Aldrich) in 10 μL and incubated for 1 h at 37°C with 5 IU of alkaline phosphatase (Sigma-Aldrich) in 0.4 M of phosphate buffer, pH: 8.8. All of the procedures were performed in the dark under argon. The mixture was filtered using an Amicon Micropure-EZ filter (Millipore), and 50 μl of each sample was used for 8-OHdG determination. The values are expressed as ng 8-OHdG/ng total DNA.
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Statistical analysis The reported data are expressed as the mean ⫾ SEM of at least 3 independent experiments. Statistical comparison of differences between groups was carried out using one-way analysis of variance followed by Student–Newman–Keuls multiple comparison test. A p value of ⱕ 0.05 was considered significant. Calculations were done using GraphPad Prism 2.0 statistical program (GraphPad Software, San Diego, CA, USA).
Results MnII(Me2DO2A) preserves cardiac muscle cell viability impaired by H ⫹ R.
Figure 6. Evaluation of apoptosis in H9c2 cells by TUNEL assay. (A) Reoxygenation (H ⫹ R) causes a marked increase in the number of TUNEL-positive cell nuclei, which was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Scale bar: 10 μm. (B) The percentage of TUNEL-positive apoptotic cells was increased by H ⫹ R and significantly decreased by MnII(Me2DO2A). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
avidin–biotin complex (Vector Lab; 1:200). Negative controls were carried out by omitting the primary antibodies. Densitometric analysis of the intensity of NT was performed in 20 regions of interest (ROI) of 100 μm2 taken from digitized images, 10 per experimental group, using Scion Image Beta 4.0.2. Determination of 8-hydroxy-2’-deoxyguanosine 8-Hydroxy-2’-deoxyguanosine (8-OHdG) levels, an indicator of oxidative DNA damage, were determined in H9c2 cells using the Highly Sensitive 8-OHdG Check (JaICA, Japan), according to the manufacturer’s instructions. After
Evaluation of H9c2 cell viability using trypan blue assay (Figure 3) showed that reoxygenation caused a marked reduction in the amounts of viable cells. This effect was inhibited by MnII(Me2DO2A) (1 and 10 μmol/l) added at reoxygenation, when high levels of O2• are produced. The MnII(Me2DO2A)-induced cytoprotection showed a dosedependent trend, as it was expected based on the mechanism of action of MnII(Me2DO2A) which involves its functional MnII center to scavenge O2•. Of note, ZnII(Me2DO2A), made with a similar organic scaffold but lacking MnII, substituted for MnII(Me2DO2A) showed no cytoprotective effect (Figure 3). MnII(Me2DO2A) preserves cardiac muscle cell mitochondrial function impaired by H ⫹ R. Parallel experiments to explore mitochondrial integrity and function were carried out with the MTT assay, which reveals the efficiency of the respiratory chain (Figure 4A), and the TMRM assay, which evaluates the mitochondrial membrane potential (Figure 4B). The results of these experiments have consistently shown that reoxygenation caused a marked impairment of mitochondrial function in H9c2 cells. This detrimental effect was significantly blunted by MnII(Me2DO2A) (10 μmol/l) added at reoxygenation. Replacement of MnII(Me2DO2A) with ZnII(Me2DO2A) had no protective effect against mitochondrial dysfunction (Figure 4A, B).
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74 S. Nistri et al.
Figure 7. Evaluation of intracellular ROS and mitochondrial O2• production by H2DCFDA and MitoSOX assays. (A) By confocal microscopy, both these parameters were enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R) in a dose-related fashion. Scale bar: 20 μm. (B) FACS analysis confirms the microscopic findings as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose.
MnII(Me2DO2A) protects cardiac muscle cells from H ⫹ R-induced apoptosis The reduction of H ⫹ R-induced oxidative stress by MnII(Me2DO2A) resulted in a significant decrease in apoptotic cell death. Indeed, the occurrence of mPTP typical of early apoptosis (Figure 5A), the activation of caspase-3 (Figure 5B), and the percentage of TUNELpositive nuclei (Figure 6) were markedly increased in H9c2 cells exposed to H ⫹ R, while the addition of 1 and 10 μmol/l MnII(Me2DO2A) at reoxygenation significantly attenuated these changes (Figures 5 and 6). MnII(Me2DO2A) protects cardiac muscle cells from oxidative damage induced by H ⫹ R MnII(Me2DO2A) decreased H9c2 cell death by reducing the oxidative stress occurring at reoxygenation. In fact, determination of intracellular ROS and mitochondrial O2• by loading the cells with the fluorescent probes H2DCFDA and MitoSOX, respectively, showed that these oxidant species were markedly increased by H ⫹ R, whereas they were significantly reduced by 10 μmol/l and, to a lesser extent, 1 μmol/l of MnII(Me2DO2A) (Figure 7A, B). In keeping with these findings, the levels of immunoreactive NT, a marker of protein nitration which was enhanced upon H ⫹ R, were significantly reduced after the addition of 10 μmol/l of MnII(Me2DO2A)
at reoxygenation (Figure 8A, B). Similar findings were observed in the experiments performed to evaluate the degree of DNA oxidation (Figure 9): in this instance, 10 μmol/l of MnII(Me2DO2A) significantly reduced the levels of 8-OHdG in H9c2 cell lysates. MnII(Me2DO2A) has no toxic effects on cardiac muscle cells As shown by the MTT assay (Figure 10), MnII(Me2DO2A) had no toxic effect on H9c2 cells, even at 10–100-fold higher concentrations (100 μmol/l) and for longer exposure times (24 h) than those displaying significant biological effects. Similarly, ZnII(Me2DO2A) (0.1–100 μmol/l) was also innocuous for the cells (Figure 10). Discussion Formation of reactive oxygen and nitrogen species and the resulting nitroxidative stress are the most invoked pathogenic mechanisms of ischemia–reperfusion-dependent diseases [9–13] and justify the search for new drugs capable of limiting the overproduction of O2•, NO•, and ONOO⫺. In this context, previous studies have demonstrated that low-molecular-weight NO• scavengers and SOD-mimetic Mn-containing porphyrins can preserve the function of isolated rat myocardial mitochondria subjected
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Superoxide scavenger and H/R cardiac cells
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Figure 9. Evaluation of oxidative stress by 8-OHdG assay. 8-OHdG, an index of DNA oxidation, was enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
we have shown that MnII(Me2DO2A) has lipophilic properties which allow it to easily cross the cell membranes [25]. This property suggests that this scavenger was able to attain O2•-neutralizing levels within the cardiac muscle cells and exert antioxidative effects close to the mitochondrial site of O2• generation, as suggested by the results of the MitoSOX and TMRM membrane potential assays. Moreover,
Figure 8. Evaluation of oxidative stress by NT detection. (A) Immunoreactive NT, an index of protein nitration, was enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Scale bar: 10 μm. (B) Densitometric analysis of NT immunostaining. Significance of differences: °°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
to hypoxia–reoxygenation and can protect the heart from ischemia–reperfusion injury in in vivo models [42,43]. On a concurrent line of evidence, potentiation of O2• decomposition capability of H9c2 rat cardiomyocytes by transfection with Cu/Zn SOD was found to increase their resistance to hypoxia–reoxygenation damage [19]. The present cell culture model is intended to study the possible protection afforded by O2• scavenging on cardiac muscle cells subjected to hypoxia–reoxygenation-induced nitroxidative stress. In the noted experimental conditions, the O2• scavenger MnII(Me2DO2A), added at reoxygenation at low, micromolar concentrations, effectively prevented mitochondrial O2•generation, intracellular ROS generation, protein nitroxidation, and oxidative DNA damage, thereby reducing apoptotic cell death, improving mitochondrial function, and increasing cell viability. In a previous study,
Figure 10. Evaluation of the toxicity of MnII(Me2DO2A) and ZnII(Me2DO2A) on H9c2 cells by the MTT assay. Neither compounds show cytotoxic effects at any dose and exposure time assayed.
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76 S. Nistri et al. MnII forms a highly stable complex at physiological pH with the polyamine–polycarboxylate scaffold. Other metal cations present in the cellular environment, including CaII, MgII, and KI, give remarkably less stable complexes than the functional MnII ion. The high stability of the MnII(Me2DO2A) prevents both de-metallation of the complex upon MnII complexation by cellular chelating agents and transmetallation reactions, due to complexation of the ligand to other metals present in the cellular medium. As a matter of fact, MnII(Me2DO2A) does not release MnII even in the presence of excess amount of CaII and other metal ions, and of broad variations of pH (at pH: 6, MnII release is less than 5%) [25]. These properties are particularly important in view of a possible pharmacological extension of MnII(Me2DO2A) to the protection of ischemic–reperfused heart in vivo: in fact, the injured myocardium undergoes an overload of CaII [44], which may potentially compete with the active MnII center of the compound, and prominent acidosis due to metabolic impairment. Moreover, MnII(Me2DO2A) is less susceptible to inactivation by oxidative stress conditions than endogenously or exogenously administered SOD [6,19]. The possible molecular mechanism of O2• scavenging by MnII(Me2DO2A) may consist of a catalytic cycle involving oxidation of MnII to MnIII by O2• and then reduction of the resulting MnIII complex by another O2• to form the initial MnII compound [25,28]. Interestingly, MnII in aqueous solution is oxidized via a single-electron process with redox potential higher than that of natural Mn-SODs. This suggests that, in the cellular environment, MnIII(Me2DO2A) reduction may occur upon reaction with O2• as well as other cytoplasmic reductants. To confirm these assumptions, the antioxidative effects of MnII(Me2DO2A) were completely lost when inactive ZnII was substituted for MnII. Taken together, these features allow MnII(Me2DO2A) to behave as an efficient O2• scavenger. Of note, MnII pentaazamacrocyclic complexes have been shown to react with ONOO⫺ and dismute NO•, albeit at lower rates than with O2• [45]. Whether MnII(Me2DO2A) may also be able to remove harmful ONOO⫺ and excess NO•, thereby increasing its antioxidant properties, remains a tantalizing matter for further investigation.
Conclusions Our study suggests that MnII(Me2DO2A) is a promising member of a new class of SOD-mimetic antioxidant drugs, capable of reducing O2•-mediated cell injury which occurs at reoxygenation after prolonged hypoxia. Further studies on animal models of ischemia and reperfusion are required to corroborate this assumption. In a drug-developing perspective, MnII(Me2DO2A) deserves to be further investigated and exploited to reduce ischemia–reperfusion organ damage in acute vascular diseases, as well as to supplement the incubation fluid of explanted organs and extend their viability in view of transplantation.
Acknowledgments The authors gratefully acknowledge Dr. Eng. Moreno Naldoni, CEO of General Project Ltd., for kind gift of MnII(Me2DO2A). Declaration of interest The authors declare that they have no financial, consulting, and personal relationships with other people or organizations that could bias the present work. This work was supported by research funds from the University of Florence issued to Silvia Nistri. Dr. Barbara Valtancoli is recipient of a research grant from the Ente Cassa di Risparmio di Firenze. References [1] Gerschman R, Gilbert DL, Nye SW, Dwyer P, Fenn WO. Oxygen poisoning and x-irradiation: a mechanism in common. Science 1954;119:623–626. [2] Murphy E, Steenbergen C. Mechanisms underlying acute protection from cardiac ischemia-reperfusion injury. Physiol Rev 2008;88:581–609. [3] McCord JM, Fridovich I. Superoxide dismutase: an enzymatic function for erythrocuprein (hemocuprein). J Biol Chem 1969;244:6049–6055. [4] Fridovich I. Superoxide dismutases. An adaptation to a paramagnetic gas. J Biol Chem 1989;264:7761–7764. [5] Johnson F, Giulivi C. Superoxide dismutases and their impact upon human health. Mol Aspects Med 2005;26:340–352. [6] Finkel T. Radical medicine: treating ageing to cure disease. Nat Rev Mol Cell Biol 2005;6:971–976. [7] Wang ZQ, Porreca F, Cuzzocrea S, Galen K, Lightfoot R, Masini E, et al. A newly identified role for superoxide in inflammatory pain. J Pharmacol Exp Ther 2004;309: 869–878. [8] Salvemini D, Wang ZQ, Zweier JL, Samouilov A, Macarthur H, Misko TP, et al. A nonpeptidyl mimic of superoxide dismutase with therapeutic activity in rats. Science 1999; 286:304–306. [9] Jaeschke H, Woolbright BL. Current strategies to minimize hepatic ischemia-reperfusion injury by targeting reactive oxygen species. Transplant Rev (Orlando) 2012;26:103–114. [10] Murphy MP. How mitochondria produce reactive oxygen species. Biochem J 2009;417:1–13. [11] Poyton RO, Ball KA, Castello PR. Mitochondrial generation of free radicals and hypoxic signaling. Trends Endocrinol Metab 2009;20:332–340. [12] Castello PR, Woo DK, Ball K, Wojcik J, Liu L, Poyton RO. Oxygen-regulated isoforms of cytochrome c oxidase have differential effects on its nitric oxide production and on hypoxic signaling. Proc Natl Acad Sci USA 2008;105:8203–8208. [13] Galkin A, Higgs A, Moncada S. Nitric oxide and hypoxia. Essays Biochem 2007;3:29–42. [14] Kokura S, Yoshida N, Yoshikawa T. Anoxia/reoxygenationinduced leukocyte-endothelial cell interactions. Free Radic Biol Med 2002;33:427–432. [15] Bencini A, Failli P, Valtancoli B, Bani D. Low molecular weight compounds with transition metals as free radical scavengers and novel therapeutic agents. Cardiovasc Hematol Agents Med Chem 2010;8:128–146. [16] Ferrer-Sueta G, Radi R. Chemical biology of peroxynitrite: kinetics, diffusion, and radicals. ACS Chem Biol 2009;4: 161–177.
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Superoxide scavenger and H/R cardiac cells [17] Xie YW, Kaminski PM, Wolin MS. Inhibition of rat cardiac muscle contraction and mitochondrial respiration by endogenous peroxynitrite formation during posthypoxic reoxygenation. Circ Res 1998;82:891–897. [18] Seal JB, Gewertz BL. Vascular dysfunction in ischemiareperfusion injury. Ann Vasc Surg. 2005;19:572–584. [19] Liu J, Hou J, Xia ZY, Zeng W, Wang X, Li R, et al. Recombinant PTD-Cu/Zn SOD attenuates hypoxiareoxygenation injury in cardiomyocytes. Free Radic Res 2013; 47:386–393. [20] McCord JM, Edeas MA. SOD, oxidative stress and human pathologies: a brief history and a future vision. Biomed Pharmacother 2005;59:139–142. [21] Slemmer JE, Shacka JJ, Sweeney MI, Weber JT. Antioxidants and free radical scavengers for the treatment of stroke, traumatic brain injury and aging. Curr Med Chem 2008;15: 404–414. [22] Riley DP, Weiss RH. Manganese macrocyclic ligand complexes as mimics of superoxide dismutase. J Am Chem Soc 1994;116:387–388. [23] Bani D, Bencini A. Developing ROS scavenging agents for pharmacological purposes: recent advances in design of manganese-based complexes with anti-inflammatory and antinociceptive activity. Curr Med Chem 2012;19:4431–4444. [24] Batinic-Haberle I, Tovmasyan A, Roberts ERH, Vujaskovic Z, Leong KW, Spasojevic I. SOD Therapeutics: latest insights into their structure-activity relationships and impact on the cellular redox-based signaling pathways. Antioxid Redox Signal 2014;20:2372–2415. [25] Failli P, Bani D, Bencini A, Cantore M, Di Cesare Mannelli L, Ghelardini C, et al. A novel manganese complex effective as superoxide anion scavenger and therapeutic agent against cell and tissue oxidative injury. J Med Chem 2009;52:7273–7283. [26] Bianchi A, Calabi L, Corana F, Fontana S, Losi P, Maiocchi A, et al. Thermodynamics and structural properties of Gd(III) complexes with polyamino-polycarboxylic ligands: basic compound for the development of MRI contrast agents. Coord Chem Rev 2000;24:309–393. [27] Merbach AE, Tóth E. The Chemistry of Contrast Agents in Medical Magnetic Resonance Imaging. New York: John Wiley & Sons; 2001. [28] Cinci L, Masini E, Bencini A, Valtancoli B, Mastroianni R, Calosi L, Bani D. Suppression of allergen-induced respiratory dysfunction and airway inflammation in sensitized guinea pigs by Mn(II)(Me(2)DO2A), a novel superoxide scavenger compound. Free Radic Biol Med 2010;48:1525–1534. [29] Di Cesare Mannelli L, Bani D, Bencini A, Brandi ML, Calosi L, Cantore M, et al. Therapeutic effects of the superoxide dismutase mimetic compound MnIIMe2DO2A on experimental articular pain in rats. Mediat Inflamm 2013:2013:905360. [30] Bani D, Bencini A, Cantore M, Failli P, Giorgi C, Valtancoli B. Manganese and cobalt complexes, their preparation and use. International patent Appl. No. PCT/IB2009/053859, University of Florence, Italy, March 11th, 2010. Owned by General Project Ltd., Montespertoli (Florence), Italy.
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[31] Lindoy LF. The Chemistry of Macrocyclic Ligand Complexes Cambridge, UK: Cambridge University Press; 1989. [32] Taqui-Khan M, Martell AE. Thermodynamic quantities associated with the interaction of adenosine triphosphate with metal ions. J Am Chem Soc 1966;88:668–671. [33] Martin RB, Edsall JT. The association of divalent cations with glutathione. J Am Chem Soc 1959;81:4044–4047. [34] Amico P, Bonomo RP, Calì R, Cucinotta V, Daniele PG, Ostacoli G, Rizzarelli E. Ligand-ligand interactions in metal complexes. Thermodynamic and spectroscopic investigation of simple and mixed copper(II) and zinc(II) substituted malonate complexes with 2,2′-bipyridyl in aqueous solution Inorg Chem 1989;28:3555–3561. [35] Martell AE, Smith RM. NIST Standard Reference Database 46 Version 8.0 Gaithersburg. USA: NIST; 2004. [36] Thun H, Guns W, Verbeek F. The stability constants of some bivalent metal complexes of α-hydroxyisobutyrate and lactate. Anal Chim Acta 1967;37:332–336. [37] Spasojevic I, Li AM, Tovmasyan A, Rajic Z, Salvemini D, St. Clair D, et al. Accumulation of porphyrin-based SOD mimics in mitochondria is proportional to their lipophilicity. S. cerevisiae study of ortho Mn(III) N-alkylpyridylporphyrins. Free Radic Biol Med 2010;S199:49. [38] Cotton FA, Wilkinson G, Murillo CA, Bochmann M. Advanced Inorganic Chemistry, 6th ed New York: Wiley-Interscience; 1999. [39] Zhang ZL, Fan Y, Liu ML. Ginsenoside Rg1 inhibits autophagy in H9c2 cardiomyocytes exposed to hypoxia/reoxygenation. Mol Cell Biochem 2012;365:243–250. [40] Petronilli V, Miotto G, Canton M, Brini M, Colonna R, Bernardi P, Di Lisa F. Transient and longlasting openings of the mitochondrial permeability transition pore can be monitored directly in intact cells by changes in mitochondrial calcein fluorescence. Biophys J 1999;765:725–754. [41] Masini E, Cuzzocrea S, Mazzon E, Muià C, Vannacci A, Fabrizi F, Bani D. Protective effects of relaxin in ischemia/ reperfusion-induced intestinal injury due to splanchnic artery occlusion. Br J Pharmacol 2006;148:1124–1132. [42] Robin E, Derichard A, Vallet B, Hassoun SM, Neviere R. Nitric oxide scavenging modulates mitochondrial dysfunction induced by hypoxia/reoxygenation. Pharmacol Rep 2011; 63:1189–1194. [43] Masini E, Cuzzocrea S, Mazzon E, Marzocca C, Mannaioni PF, Salvemini D. Protective effects of M40403, a selective superoxide dismutase mimetic, in myocardial ischaemia and reperfusion injury in vivo. Br J Pharmacol 2002;136:905–917. [44] Bourdillon PD, Poole-Wilson PA. Effects of ischaemia and reperfusion on calcium exchange and mechanical function in isolated rabbit myocardium. Cardiovasc Res 1981;15: 121–130. [45] Filipović MR, Koh AC, Arbault S, Niketić V, Debus A, Schleicher U, et al. Striking inflammation from both sides: manganese(II) pentaazamacrocyclic SOD mimics act also as nitric oxide dismutases: a single-cell study. Angew Chem Int Ed Engl 2010;49:4228–4232.
ORIGINAL ARTICLE
A new low molecular weight, MnII-containing scavenger of superoxide anion protects cardiac muscle cells from hypoxia/reoxygenation injury S. Nistri1, G. Boccalini1, A. Bencini2, M. Becatti3, B. Valtancoli2, L. Conti2, L. Lucarini4 & D. Bani1 1Department
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of Experimental & Clinical Medicine, Section of Anatomy & Histology, Research Unit of Histology & Embryology, University of Florence, Florence, Italy, 2Department of Chemistry, University of Florence, Florence, Italy, 3Department of Experimental & Clinical Biomedical Sciences “Mario Serio”, University of Florence, Florence, Italy, and 4Department of Neurosciences and Psychology, Drug Area and Child’s Health, Section of Pharmacology, University of Florence, Florence, Italy Abstract Reperfusion injury after oxygen starvation is a key pathogenic step in ischemic diseases. It mainly consists in oxidative stress, related to mitochondrial derangement and enhanced generation of reactive oxygen species (ROS), mainly superoxide anion (O2•2), and peroxynitrite by cells exposed to hypoxia. This in vitro study evaluates whether MnII(4,10-dimethyl-1,4,7,10-tetraazacyclododecane-1,7-diacetate).2H2O, or MnII(Me2DO2A), a new low molecular weight, MnII-containing O2• scavenger, has a direct protective action on H9c2 rat cardiac muscle cells subjected to hypoxia and reoxygenation. MnII(Me2DO2A) (1 and 10 μmol/l) was added to the culture medium at reoxygenation and maintained for 2 h. In parallel experiments, the inactive congener ZnII(Me2DO2A), in which ZnII replaced the functional MnII center in the same organic scaffold, was used as negative control. MnII(Me2DO2A) (10 μmol/l) significantly increased cardiac muscle cell viability (trypan blue assay), improved mitochondrial activity (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide test, membrane potential Δψ), reduced apoptosis (mitochondrial permeability transition pore opening, caspase-3, terminal deoxynucleotidyl transferase deoxyuridine triphosphate nick end labeling assay), decreased intracellular ROS levels (2¢,7¢-dichlorodihydrofluorescein diacetate and MitoSOX assays), and decreased protein nitroxidation (nitrotyrosine [NT] expression) and DNA oxidation (8-hydroxy-deoxyguanosine levels). Of note, ZnII(Me2DO2A) had no protective effect. The mechanism of MnII(Me2DO2A) relies on concentration-dependent removal of harmful O2• generated at reoxygenation from dysfunctional mitochondria in hypoxia-induced cells, as indicated by the MitoSOX assay. This study suggests that MnII(Me2DO2A) is a promising antioxidant drug capable of reducing O2•-mediated cell oxidative stress which occurs at reoxygenation after hypoxia. In perspective, MnII(Me2DO2A) might be used to reduce ischemia–reperfusion organ damage in acute vascular diseases, as well as to extend the viability of explanted organs before transplantation. Keywords: superoxide anion, ROS scavenger, H9c2 cardiac muscle cells, hypoxia, reoxygenation, oxidative stress
Introduction During oxidative metabolism, at different points in electron transport chain, O2 is reduced through one-electron paths to superoxide anion (O2•), a highly reactive radical which can oxidize and functionally compromise cell’s membrane lipids, proteins, and nucleic acids[1,2]. Normally, O2• levels are kept under the damage threshold by endogenous antioxidants, among which a key role is played by superoxide dismutases (SODs), which catalyze the dismutation of O2• to O2 and H2O2 by a reaction requiring a transition metal at the catalytic site [3–5]. However, oxidative stress causes a rapid inactivation of SODs, sparkling a vicious cycle that causes ever-increasing, harmful O2• tissue levels [6,7]. A paradigm of oxidative stress-related diseases includes ischemia–reperfusion, as it occurs during acute myocardial infarction and stroke [8]. In this process, the initial ischemic damage is then compounded by additional and more severe injury caused by reoxygenation upon blood flow restoration. Multiple interplaying mechanisms play a role in reperfusion-induced damage
and mainly involve oxidative stress [9]. In fact, mitochondrial dysfunction of hypoxia-exposed cells impairs the electron flow and enhances O2• formation and release [10,11]. Moreover, mitochondria can use the respiratory chain to reduce nitrite (NO2⫺) to nitric oxide (NO•) [12]. Because cytochrome oxidase produces NO• from nitrite at low O2 concentrations, the mitochondrially generated oxidants whose concentration increase under hypoxic conditions also include peroxynitrite (ONOO⫺), which is formed by the reaction between O2• and NO• [13]. In vivo, these mitochondrial mechanisms intrinsic to hypoxic cells are paralleled by inflammatory cell recruitment secondary to endothelial injury [14], which results in further enhancement of the generation of ONOO⫺. Over the past decade, ONOO⫺ has been identified as the effector of most, if not all, detrimental effects of excess NO• [15,16] and is regarded as the major harmful oxidant in myocardial hypoxia–reoxygenation injury [17]. The above notions underscore that O2• and NO• play a pivotal role in oxidative stress upon hypoxia–reoxygenation and hence can be suitable target for new pharmacological
Correspondence: Silvia Nistri, PhD, DSc, Department of Experimental and Clinical Medicine, Section of Anatomy and Histology, Research Unit of Histology and Embryology, University of Florence, viale G. Pieraccini, 6. I-50139 Florence, Italy.Tel: ⫹ 39 055 4271 387. Fax: ⫹ 39 055 4271 385. E-mail: silvia.nistri@unifi.it (Received date: 16 July 2014; Accepted date: 17 October 2014; Published online: 13 November 2014)
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68 S. Nistri et al. interventions. Theoretically, removal of excess O2• is the best approach to reduce oxidative stress, while leaving enough NO• to sustain residual vascular function [18]. In this context, extractive or recombinant SOD appears to be the logical choice. Indeed, transfection of H9c2 rat cardiomyocytes with Cu/Zn SOD was found to attenuate hypoxia–reoxygenation injury [19]. However, the pharmaceutical use of SOD is limited by its poor stability in water and intracellular penetration, immunogenicity, short half-life, and unfavorable yield/cost ratio [20,21]. Because of these limitations, pharmacological research has diverged toward non-peptidic, low molecular weight, Mn-containing compounds capable of efficiently catalyzing O2• dismutation as do authentic SODs [22]. Their efficacy relies on the wellknown chemical property of Mn ion to bind to and react with free radicals, including O2•. Several MnII and MnIII compounds with organic scaffolds, in particular, MnIII complexes with porphyrins, salen and its derivatives, and MnII chelates with cyclic polyamines and polyamine–polycarboxylates have been synthesized as reactive oxygen species (ROS) scavengers and tested for therapeutic properties in cellular and animal models of oxidative stress [23,24]. In recent years, we investigated the chemical and biological properties of a new polyamine–polycarboxylate MnII complex, MnII(4,10-dimethyl-1,4,7,10-tetraazacyclododecane-1,7-diacetate), also termed “MnII(Me2DO2A),” effective as O2• scavenger [25]. In principle, polyamine– polycarboxylate scaffolds represent an optimal tool for the synthesis of highly stable MnII complexes. In fact, these molecules are known for their ability to strongly bind a number of metals, from alkali and alkali earths to lanthanide and transition metal cations, including MnII. Moreover, these compounds are resistant to oxidizing and reducing agents, are highly soluble in aqueous media, and exhibit low toxicity and all properties that render them suitable for several biological and medicinal applications [26,27]. In particular, MnII(Me2DO2A) effectively reduced oxidative stress in cell culture models and oxidative tissue injury in animal models of inflammation [25,28,29]. The current study was designed to test the antioxidant effects of MnII(Me2DO2A) using H9c2 cardiac muscle cells subjected to hypoxia–reoxygenation in vitro. We used the inactive congener ZnII(Me2DO2A), in which the functional MnII was replaced with a ZnII ion lacking the capability to catalyze O2•decomposition, as control. Materials and methods Reagents The amount of MnII(Me2DO2A) required to perform the present experiments was kindly donated by the patent owner General Project Ltd., Montespertoli (Florence), Italy. The inactive compound ZnII(Me2DO2A) was synthesized in our laboratory from Me2DO2A following the same procedure as MnII(Me2DO2A). The formation constants of the ZnII complexes have been determined using the same procedure reported for the MnII complexes [25].
Unless otherwise specified, the other reagents used for the experiments were from Sigma-Aldrich (Milan, Italy). Chemical characterization of MnII(Me2DO2A) and ZnII(Me2DO2A) The compound MnII(Me2DO2A) was synthesized in our laboratories and patented [30]. The organic scaffold contains a tetraamine macrocyclic moiety, capable of forming complexes with transition metals characterized by high thermodynamic stability, kinetic inertness, and poor tendency to bind alkali or alkaline metal cations [31]. It also possesses two carboxylic moieties as pendant arms, acting as additional binding sites for metal cations. Like most polyamine–carboxylate ligands, Me2DO2A is able to form stable complexes with alkali and alkaline-earth metal cations. Table I reports the stability constants of its complexes with MnII in comparison with MgII, CaII, and NaI (the constant of the complex with KI is very low—log K ⬍ 2—and cannot be reliably determined). Table I shows that MnII forms the most stable complex with the ligand in its fully deprotonated form, where the deprotonated carboxylic groups are likely coordinated with the metal center (Scheme 2), yielding the MnII(Me2DO2A) neutral complex (Figure 1). To rule out the possibility of exchange reactions with MgII, CaII, and NaI, whose intracellular concentrations are three or more orders of magnitude higher than MnII, competition experiments were carried out with Me2DO2A and MnII at very low concentrations (1 μM) in the presence of excess NaI (165 mM), CaII (2.5 mM), and MgII (1.2 mM) [25]. The competition diagram reported in Figure 2A shows that MnII remains completely bound to Me2DO2A, while CaII, MgII, and NaI appeared unbound by the ligand. Therefore, the MnII(Me2DO2A) complex is substantially unaffected by other metal ion competitors in the medium, and no detectable MnII release occurs because of complexation with other metals. At variance with MgII, CaII, and NaI, the MnII(Me2DO2A) complex is also able to bind acidic protons to give protonated complexes of the type MnII(HMe2DO2A)⫹ and MnII(H2Me2DO2A)2⫹ in water. However, the proton affinity of these MnII(Me2DO2A) species is rather weak and only minor amounts of the protonated complexes are formed at acidic pH values. In fact, proton binding would Table I. Stability constants of the NaI, MgII, CaII, MnII, and ZnII complexes with H2Me2DO2A (H2L) and its deprotonated forms— (HMe2DO2A) ⫺ (HL⫺) and (Me2DO2A)2 (L2⫺) (I ⫽ 0.1 M, 25°C). Reaction Na⫹ ⫹ L2⫺ ⫽ [NaL] ⫺ Ca2⫹ ⫹ L2⫺ ⫽ [CaL] Mg2⫹ ⫹ L2⫺ ⫽ [MgL] Mn2⫹ ⫹ L2⫺ ⫽ [MnL] [MnL] ⫹ H⫹⫺ ⫽ [Mn(HL)]⫹ [Mn(HL)]⫹ ⫹ H⫹⫺ ⫽ [Mn(H2L)]2⫹ Zn2⫹⫺ ⫹ L2⫺ ⫽ [ZnL] [ZnL] ⫹ H⫹ ⫽ [Zn(HL)]⫹ [Zn(HL)]⫹ ⫹ H⫹ ⫽ [Zn(H2L)]2⫹
Log K 2.08 (7) 7.67 (8) 5.74 (7) 14.73 (2) 4.53 (7) 3.96 (2) 15.91 (1) 4.25 (1) 3.90 (1)
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Figure 1. Polyamine–polycarboxylate scaffold (A) and 3D structure (B) of MnII(Me2DO2A).
imply detachment of the coordinated carboxylate group from the metal center, which is an unlikely and unfavored event. As a matter of fact, as shown in Figure 2B, the neutral MnII(Me2DO2A) complex can be regarded as the only species present in solution at neutral pH value. The stability of MnII(Me2DO2A) is also higher than that of other MnII complexes with endogenous ligands present
Figure 2. (A) Overall percentages of the species complexed with Mn(II), Ca(II), Mg(II), and Na(I) as a function of pH in competitive systems containing Mn(II) (1 μM), Ca(II), (2.5 mM), Mg(II) (1.5 mM), and Na(I) (165 mM), and Me2DO2A (1 μM). ∑[Mn(HxL)] ⫻ ⫹ ⫽ [MnL] ⫹ [Mn(HL)] ⫹ ⫹ [Mn(H2L)]2 ⫹; ∑[HxL4](x ⫺ 2) ⫹ ⫽ [L4]2 ⫺ ⫹ [HL4] ⫺ ⫹ [H2L4] ⫹ [H3L4] ⫹ ⫹ [H4L4]2 ⫹; (B) distribution curves of the complexes with Me2DO2A (25°C, I ⫽ 0.1 M). At physiological pH, MnII(Me2DO2A) does not release MnII, even in the presence of other metal cations; at pH: 6, MnII release is less than 5%.
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in the cellular environment, such as ATP (log K ⫽ 4.66 for the equilibrium MnII ⫹ ATP ⫽ [MnII(ATP)]) [32], glutathione (logK ⫽ 2.7 for MnII ⫹ GSH ⫽ [MnII(GSH)]) [33], and carboxylate anions (logK ⫽ 3.79, 1.68, and 0.92 for MnII complexation by citrate [34], maleate [35], and Llactate anions [36]), thus excluding substantial de-metallation of MnII(Me2DO2A) by biological chelators. MnII (Me2DO2A) was rather resistant to electrochemical oxidation in water solution, displaying an onset oxidation potential of 0.86 V and a formal E1/2 potential of 0.85 V [25]. MnII oxidation is characterized by a fast single-electron transfer process ([MnIIL] → [MnIIIL]⫹) followed by slow rearrangement of the carboxylate groups around MnIII. In our previous study, we demonstrated that MnII(Me2DO2A) had strong O2• scavenging properties in vitro and in vivo, with a kcat value of 1.1 ⫻ 106 M⫺ 1 s⫺ 1 [25,28]. The structural characteristics of MnII(Me2DO2A) resemble, in part, those of the M40403 compound, where MnII is bound by a pentaamine macrocycle to afford a positively charged complex. However, the catalytic ability in O2• dismutation of MnII(Me2DO2A) is about one order of magnitude lower than M40403, the kcat of the latter in water being 2 ⫻ 107 M⫺ 1 s⫺ 1 [8]. At variance with M40403, MnII(Me2DO2A) is a neutral complex and the metal cation is more coordinatively saturated and shielded by the simultaneous coordination of the tetraamine macrocycle and the two carboxylate anionic moieties, making it less accessible to the O2• anion in the first step of the catalytic process. On the other hand, binding of the carboxylate groups makes MnII(Me2DO2A) thermodynamically stable and, therefore, resistant to possible metal release. Moreover, the neutral charge of MnII(Me2DO2A) allows it to cross the cell membranes and attain significant intracellular concentrations [25]. Moreover, the presence of two methyl groups appended to amine groups prevents the nitrogen atoms to interact via hydrogen bonding with water molecules, and enhances the lipophilic features of the complex. Interestingly, the absence of negative charge and increased lipophilicity has been shown to facilitate accumulation of manganese complexes in mitochondria [24, 37]. This may suggest that MnII(Me2DO2A) can directly scavenge O2• in mitochondria. At the same time, the amine groups at positions 4 and 10 can be easily functionalized with other organic moieties: this property may be exploited to increase the lipophilic characteristics of the compounds by substituting the methyl groups with hydrophobic alkyl or aryl moieties. Me2DO2A displays a binding ability for ZnII similar to that observed for MnII. As shown in Table I, in aqueous solution this ligand forms a neutral ZnII(Me2DO2A) complex that is slightly more stable than the MnII complex. Similar to MnII(Me2DO2A), in ZnII(Me2DO2A) the metal is likely coordinated by both the tetraamine scaffold and the two carboxylate groups. Protonation of the carboxylate groups occurs at acidic pH values, generating ZnII(HMe2DO2A)⫹ and ZnII(H2Me2DO2A)2⫹. As for MnII(Me2DO2A), these features ensure that neither metal release at neutral or slightly acidic pH values nor
70 S. Nistri et al. transmetallation reaction can occur in solution. From an electrochemical point of view, the ZnII ion is extremely stable to reduction; the reduction potential of the ZnII/ Zn0 couple being ⫺ 0.76 V. At the same time, oxidation states higher than II are not accessible for zinc [38]. These characteristics make this complex inert to reducing and oxidizing agents present in cellular environment, including O2• or other ROS.
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Cell culture and treatments H9c2 embryonic rat cardiac muscle cells, obtained from European Collection of Cell Cultures (ECACC, Salisbury, UK), were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Invitrogen, Carlsbad, CA, USA), 2 mM of glutamine, 250 U/ml of penicillin G, and 250 μg/ml of streptomycin, in a humidified atmosphere with 5% CO2 at 37°C. These cells were subjected to hypoxia and reoxygenation (H ⫹ R), simulated in vitro by substrate starvation plus hypoxia followed by reoxygenation, as previously described [39] with minor modifications. The cells were incubated in DMEM with no serum or glucose and placed in a hypoxic chamber saturated with a 0.1% O2, 5% CO2, and ≈ 95% N2 gaseous mix, humidified, and warmed at 37°C, for 7 h. At the end of hypoxia, the cells were reoxygenated for 2 h by incubation in normoxic conditions in glucose-containing, serum-free DMEM. Normoxic control cultures were also prepared. Cells were treated with or without MnII(Me2DO2A) at 2 different doses (1 and 10 μmol/l) and added at reoxygenation, concurrently with the peak of ROS generation (H ⫹ MnII(Me2DO2A)⫹ R). The given MnII(Me2DO2A) doses were chosen on the basis of our previous in vitro studies [25]. As control for the specific capability of MnII(Me2DO2A) in suppressing oxidative stress by redox reaction, some cell viability experiments were performed using the inactive congener ZnII(Me2DO2A) at the same concentrations as MnII(Me2DO2A) added at reoxygenation (H ⫹ ZnII(Me2DO2A)⫹ R). Separate experiments were performed to assess the toxicity of MnII(Me2DO2A) and ZnII(Me2DO2A) in H9c2 cells. In these experiments, the two compounds were added to the cell cultures at increasing concentrations (0.1–100 μmol/l) for 2 and 24 h; cell viability was then assayed by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) test. Trypan blue viability assay The trypan blue exclusion method was used to assess cell viability. H9c2 cells (5 ⫻ 104/well) were seeded in 24-well plates. At the end of the treatments, the cells were gently detached by trypsin/ethylenediaminetetraacetic acid, resuspended in culture medium, and mixed 1:1 with 0.4% trypan blue solution. The final cell suspensions were counted under a phase contrast inverted microscope using a Burker chamber. Viable cells were expressed as percentage of the total counted cells.
MTT assay Cell mitochondrial function was measured using the MTT assay. H9c2 cells (5 ⫻ 104/well) were seeded in 24-well plates. At the end of the treatments, the culture medium was removed from each well and replaced with 300 μl of MTT stock solution, followed by 4-h incubation at 37°C. Then, 350 μl of dimethyl sulfoxide were added to each well to dissolve the formazan crystals. The plate was gently shaken for 10 min and read at 550 nm on a plate reader. Optical density was assumed as indicator of mitochondrial activity and, indirectly, cell viability. Mitochondrial membrane potential (Δψ) Mitochondrial membrane potential was assessed using tetramethylrhodamine methyl ester perchlorate (TMRM), a lipophilic potentiometric fluorescent dye that distributes between the mitochondria and cytosol in proportion to Δψ by virtue of its positive charge. At low concentrations, the fluorescence intensity depends on dye accumulation in mitochondria, which in turn is directly related to mitochondrial potential. For confocal microscope analysis, cells were cultured on glass coverslips and loaded for 20 min at 37°C with TMRM, dissolved in 0.1% dimethyl sulfoxide (DMSO) to a final concentration of 100 nM in the culture medium. The cells were fixed in 2% buffered paraformaldehyde for 10 min at room temperature and the TMRM fluorescence was analyzed under a confocal Leica TCS SP5 scanning microscope (Mannheim, Germany) equipped with a helium–neon laser source, using a 543-nm excitation wavelength, and with a Leica Plan Apo x63 oil immersion objective. Mitochondrial membrane potential was also quantified by flow cytometry. Single-cell suspensions were washed twice with PBS and incubated for 20 min at 37°C in the dark with TMRM dissolved in DMEM (100 nM). The cells were then washed, resuspended in PBS, and analyzed using a FACSCanto flow cytometer (Becton–Dickinson, San Jose, CA). Mitochondrial permeability transition pore opening Mitochondrial permeability, an index of mitochondrial dysfunction and early apoptosis, was measured by calcein fluorescence, as described [40]. The fluorescent probe calcein-(acetoxymethyl) AM freely enters the cells and emits fluorescence upon de-esterification. Coloading of cells with cobalt chloride, which cannot cross the mitochondrial membranes in living cells, quenches the fluorescence in the whole cell except mitochondria. During induction of mPTP, cobalt can enter mitochondria and quench calcein fluorescence, whose decrease can be taken as a measure of the extent of mPTP induction. H9c2 cells grown on glass coverslips were loaded with calcein-AM (3 μM) and cobalt chloride (1 mM) added to the culture medium for 20 min at 37°C. The cells were then washed in PBS, fixed in 2% buffered paraformaldehyde for 10 min at room temperature, and analyzed by a Leica TCS SP5 confocal laser scanning microscope equipped with an
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argon laser source, using 488-nm excitation wavelength, and with a Leica Plan Apo x63 oil immersion objective. Mitochondrial permeability was also monitored by flow cytometry: single-cell suspensions were incubated with calcein-AM (3 μM) and cobalt chloride (1 mM) for 20 min at 37°C, washed twice with PBS, and analyzed using a FACSCanto flow cytometer (Becton–Dickinson).
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Assessment of caspase-3 activity H9c2 cells seeded on glass coverslips were incubated with FAM FLICA™ Caspase 3&7 assay kit (Immunochemistry Technologies, Bloomington, MN, USA) for 30 min, following the manufacturer’s instructions. After incubation, the cells were thoroughly washed and fixed in 2% buffered paraformaldehyde for 10 min at room temperature. Fluorescence was detected by a confocal Leica TCS SP5 scanning microscope equipped with an argon laser source, using 488-nm excitation wavelength, and a Leica Plan Apo x63 oil immersion objective. Caspase-3 activity was also quantified by flow cytometry: single-cell suspensions were incubated with FAMFLICA™ for 30 min at 37°C, washed twice with PBS, and analyzed using a FACSCanto flow cytometer (Becton– Dickinson). TUNEL assay H9c2 cells were grown on glass coverslip and subjected to the different treatments. Cell death was studied with TUNEL assay for apoptosis, performed using a KlenowFragELTM DNA fragmentation detection kit (Calbiochem,
Figure 3. Evaluation of H9c2 cell viability by trypan blue assay. Reoxygenation (H ⫹ R) causes a marked reduction of the amounts of viable cells. This effect was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R), while ZnII(Me2DO2A) had no effects (H ⫹ Zn ⫹ R). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
Figure 4. Evaluation of H9c2 cell mitochondrial integrity and function by MTT assay, which reveals the efficiency of the respiratory chain (A), and TMRM assay, which evaluates the mitochondrial membrane potential, by confocal microscopy (B) and FACS analysis (C). Reoxygenation (H ⫹ R) causes a marked mitochondrial impairment, which was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). ZnII(Me2DO2A) (H ⫹ Zn ⫹ R) had no effects (A). FACS analysis confirms the microscopic findings of TMRM fluorescence as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose. Scale bars: 20 μm. Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
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72 S. Nistri et al.
Figure 5. Evaluation of apoptosis in H9c2 cells by mPTP and caspase-3 activity assayed by confocal microscopy (A) and FACS analysis (B). Reoxygenation (H ⫹ R) causes the extinction of calcein-AM fluorescence in mitochondria by increased cobalt ion permeation, while this effect was dose-dependently inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Similarly, cytoplasmic fluorescence related to activated caspase-3 was increased by H ⫹ R and markedly decreased by MnII(Me2DO2A). Scale bars: 20 μm. FACS analysis confirms the microscopic findings as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose.
San Diego, CA, USA), as reported in the manufacturer’s instructions. Briefly, H9c2 cells were incubated with 15 mg/ml of proteinase K for 5 min at room temperature. After rinsing in PBS, the cells were immersed in the Klenow Labeling Reaction Mixture containing deoxynucleotidyl transferase and unlabelled and biotin-labeled deoxynucleotides, and incubated at 37° C for 90 min in a humid atmosphere. Then, the cells were incubated with peroxidase-conjugated streptavidin for 30 min at room temperature and the signal was revealed with 3,3′diaminobenzidine. Finally, nuclear counterstaining was achieved using methyl green. Apoptotic nuclei were recognized by the presence of dark brown staining, at variance with those of viable cells, which instead appeared pale brown or green. TUNEL-positive nuclei were counted in 10 microscopic fields for each cell preparation. TUNEL apoptotic index was then expressed as relative percentage of TUNEL-positive nuclei on the total number of methylgreen-stained nuclei. Determination of intracellular ROS and mitochondrial superoxide H9c2 cells seeded on glass coverslips were loaded with the ROS-sensitive fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA; Invitrogen, CA, USA; 2.5 μmol/l) or the mitochondrial O2•-specific fluorescent probe MitoSOX (Invitrogen; 3 μmol/l)—dissolved in 0.1% DMSO and Pluronic acid F-127 (0.01% w/v)—which were
added to cell culture media for 15 min at 37°C. The cells were fixed in 2% buffered paraformaldehyde for 10 min at room temperature, and the H2DCFDA and MitoSOX fluorescence were analyzed using a Leica TCS SP5 confocal scanning microscope equipped with an argon laser source, using 488-nm and 543-nm excitation wavelength, respectively, and a Leica Plan Apo x63 oil immersion objective. ROS and mitochondrial O2• generation were also monitored by flow cytometry: briefly, single-cell suspensions were incubated with H2DCFDA (1 μmol/l) or MitoSOX (0.5 μmol/l) for 15 min at 37°C and immediately analyzed using a FACSCanto flow cytometer (Becton–Dickinson). Data were analyzed using FACSDiva software (Becton– Dickinson). Immunohistochemical localization and quantitation of nitrotyrosine NT, an index of protein nitrosylation by harmful ONOO⫺ generated during inflammation, was determined by immunocytochemistry as described previously [41]. Briefly, H9c2 cells were grown on glass coverslip and subjected to the different treatments. The cells were fixed with formaldehyde for 10 min and then incubated with rabbit polyclonal anti-NT antibody (Upstate Biotechnology, Buckingham, UK; 1:118) at 4°C overnight. Immune reaction was revealed by goat anti-rabbit IgG conjugated with biotin (1:200; Vector Lab, Burlingame, CA, USA) followed by incubation with
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the treatments, the cells were collected with TRIzol Reagent (Invitrogen) and DNA was isolated according to the manufacturer’s instructions. Then DNA was subjected to enzymatic digestion with 10 IU of P1 nuclease (Sigma-Aldrich) in 10 μL and incubated for 1 h at 37°C with 5 IU of alkaline phosphatase (Sigma-Aldrich) in 0.4 M of phosphate buffer, pH: 8.8. All of the procedures were performed in the dark under argon. The mixture was filtered using an Amicon Micropure-EZ filter (Millipore), and 50 μl of each sample was used for 8-OHdG determination. The values are expressed as ng 8-OHdG/ng total DNA.
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Statistical analysis The reported data are expressed as the mean ⫾ SEM of at least 3 independent experiments. Statistical comparison of differences between groups was carried out using one-way analysis of variance followed by Student–Newman–Keuls multiple comparison test. A p value of ⱕ 0.05 was considered significant. Calculations were done using GraphPad Prism 2.0 statistical program (GraphPad Software, San Diego, CA, USA).
Results MnII(Me2DO2A) preserves cardiac muscle cell viability impaired by H ⫹ R.
Figure 6. Evaluation of apoptosis in H9c2 cells by TUNEL assay. (A) Reoxygenation (H ⫹ R) causes a marked increase in the number of TUNEL-positive cell nuclei, which was inhibited by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Scale bar: 10 μm. (B) The percentage of TUNEL-positive apoptotic cells was increased by H ⫹ R and significantly decreased by MnII(Me2DO2A). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
avidin–biotin complex (Vector Lab; 1:200). Negative controls were carried out by omitting the primary antibodies. Densitometric analysis of the intensity of NT was performed in 20 regions of interest (ROI) of 100 μm2 taken from digitized images, 10 per experimental group, using Scion Image Beta 4.0.2. Determination of 8-hydroxy-2’-deoxyguanosine 8-Hydroxy-2’-deoxyguanosine (8-OHdG) levels, an indicator of oxidative DNA damage, were determined in H9c2 cells using the Highly Sensitive 8-OHdG Check (JaICA, Japan), according to the manufacturer’s instructions. After
Evaluation of H9c2 cell viability using trypan blue assay (Figure 3) showed that reoxygenation caused a marked reduction in the amounts of viable cells. This effect was inhibited by MnII(Me2DO2A) (1 and 10 μmol/l) added at reoxygenation, when high levels of O2• are produced. The MnII(Me2DO2A)-induced cytoprotection showed a dosedependent trend, as it was expected based on the mechanism of action of MnII(Me2DO2A) which involves its functional MnII center to scavenge O2•. Of note, ZnII(Me2DO2A), made with a similar organic scaffold but lacking MnII, substituted for MnII(Me2DO2A) showed no cytoprotective effect (Figure 3). MnII(Me2DO2A) preserves cardiac muscle cell mitochondrial function impaired by H ⫹ R. Parallel experiments to explore mitochondrial integrity and function were carried out with the MTT assay, which reveals the efficiency of the respiratory chain (Figure 4A), and the TMRM assay, which evaluates the mitochondrial membrane potential (Figure 4B). The results of these experiments have consistently shown that reoxygenation caused a marked impairment of mitochondrial function in H9c2 cells. This detrimental effect was significantly blunted by MnII(Me2DO2A) (10 μmol/l) added at reoxygenation. Replacement of MnII(Me2DO2A) with ZnII(Me2DO2A) had no protective effect against mitochondrial dysfunction (Figure 4A, B).
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74 S. Nistri et al.
Figure 7. Evaluation of intracellular ROS and mitochondrial O2• production by H2DCFDA and MitoSOX assays. (A) By confocal microscopy, both these parameters were enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R) in a dose-related fashion. Scale bar: 20 μm. (B) FACS analysis confirms the microscopic findings as it shows that, compared with the control cells, H ⫹ R shifts the fluorescence peaks toward higher values (right), while this effect is markedly reduced by MnII(Me2DO2A), especially at the higher dose.
MnII(Me2DO2A) protects cardiac muscle cells from H ⫹ R-induced apoptosis The reduction of H ⫹ R-induced oxidative stress by MnII(Me2DO2A) resulted in a significant decrease in apoptotic cell death. Indeed, the occurrence of mPTP typical of early apoptosis (Figure 5A), the activation of caspase-3 (Figure 5B), and the percentage of TUNELpositive nuclei (Figure 6) were markedly increased in H9c2 cells exposed to H ⫹ R, while the addition of 1 and 10 μmol/l MnII(Me2DO2A) at reoxygenation significantly attenuated these changes (Figures 5 and 6). MnII(Me2DO2A) protects cardiac muscle cells from oxidative damage induced by H ⫹ R MnII(Me2DO2A) decreased H9c2 cell death by reducing the oxidative stress occurring at reoxygenation. In fact, determination of intracellular ROS and mitochondrial O2• by loading the cells with the fluorescent probes H2DCFDA and MitoSOX, respectively, showed that these oxidant species were markedly increased by H ⫹ R, whereas they were significantly reduced by 10 μmol/l and, to a lesser extent, 1 μmol/l of MnII(Me2DO2A) (Figure 7A, B). In keeping with these findings, the levels of immunoreactive NT, a marker of protein nitration which was enhanced upon H ⫹ R, were significantly reduced after the addition of 10 μmol/l of MnII(Me2DO2A)
at reoxygenation (Figure 8A, B). Similar findings were observed in the experiments performed to evaluate the degree of DNA oxidation (Figure 9): in this instance, 10 μmol/l of MnII(Me2DO2A) significantly reduced the levels of 8-OHdG in H9c2 cell lysates. MnII(Me2DO2A) has no toxic effects on cardiac muscle cells As shown by the MTT assay (Figure 10), MnII(Me2DO2A) had no toxic effect on H9c2 cells, even at 10–100-fold higher concentrations (100 μmol/l) and for longer exposure times (24 h) than those displaying significant biological effects. Similarly, ZnII(Me2DO2A) (0.1–100 μmol/l) was also innocuous for the cells (Figure 10). Discussion Formation of reactive oxygen and nitrogen species and the resulting nitroxidative stress are the most invoked pathogenic mechanisms of ischemia–reperfusion-dependent diseases [9–13] and justify the search for new drugs capable of limiting the overproduction of O2•, NO•, and ONOO⫺. In this context, previous studies have demonstrated that low-molecular-weight NO• scavengers and SOD-mimetic Mn-containing porphyrins can preserve the function of isolated rat myocardial mitochondria subjected
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Figure 9. Evaluation of oxidative stress by 8-OHdG assay. 8-OHdG, an index of DNA oxidation, was enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Significance of differences:°°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
we have shown that MnII(Me2DO2A) has lipophilic properties which allow it to easily cross the cell membranes [25]. This property suggests that this scavenger was able to attain O2•-neutralizing levels within the cardiac muscle cells and exert antioxidative effects close to the mitochondrial site of O2• generation, as suggested by the results of the MitoSOX and TMRM membrane potential assays. Moreover,
Figure 8. Evaluation of oxidative stress by NT detection. (A) Immunoreactive NT, an index of protein nitration, was enhanced upon H ⫹ R and significantly reduced by MnII(Me2DO2A) given at reoxygenation (H ⫹ Mn ⫹ R). Scale bar: 10 μm. (B) Densitometric analysis of NT immunostaining. Significance of differences: °°°p ⬍ 0.001 versus control; *p ⬍ 0.05 and ***p ⬍ 0.001 versus H ⫹ R.
to hypoxia–reoxygenation and can protect the heart from ischemia–reperfusion injury in in vivo models [42,43]. On a concurrent line of evidence, potentiation of O2• decomposition capability of H9c2 rat cardiomyocytes by transfection with Cu/Zn SOD was found to increase their resistance to hypoxia–reoxygenation damage [19]. The present cell culture model is intended to study the possible protection afforded by O2• scavenging on cardiac muscle cells subjected to hypoxia–reoxygenation-induced nitroxidative stress. In the noted experimental conditions, the O2• scavenger MnII(Me2DO2A), added at reoxygenation at low, micromolar concentrations, effectively prevented mitochondrial O2•generation, intracellular ROS generation, protein nitroxidation, and oxidative DNA damage, thereby reducing apoptotic cell death, improving mitochondrial function, and increasing cell viability. In a previous study,
Figure 10. Evaluation of the toxicity of MnII(Me2DO2A) and ZnII(Me2DO2A) on H9c2 cells by the MTT assay. Neither compounds show cytotoxic effects at any dose and exposure time assayed.
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76 S. Nistri et al. MnII forms a highly stable complex at physiological pH with the polyamine–polycarboxylate scaffold. Other metal cations present in the cellular environment, including CaII, MgII, and KI, give remarkably less stable complexes than the functional MnII ion. The high stability of the MnII(Me2DO2A) prevents both de-metallation of the complex upon MnII complexation by cellular chelating agents and transmetallation reactions, due to complexation of the ligand to other metals present in the cellular medium. As a matter of fact, MnII(Me2DO2A) does not release MnII even in the presence of excess amount of CaII and other metal ions, and of broad variations of pH (at pH: 6, MnII release is less than 5%) [25]. These properties are particularly important in view of a possible pharmacological extension of MnII(Me2DO2A) to the protection of ischemic–reperfused heart in vivo: in fact, the injured myocardium undergoes an overload of CaII [44], which may potentially compete with the active MnII center of the compound, and prominent acidosis due to metabolic impairment. Moreover, MnII(Me2DO2A) is less susceptible to inactivation by oxidative stress conditions than endogenously or exogenously administered SOD [6,19]. The possible molecular mechanism of O2• scavenging by MnII(Me2DO2A) may consist of a catalytic cycle involving oxidation of MnII to MnIII by O2• and then reduction of the resulting MnIII complex by another O2• to form the initial MnII compound [25,28]. Interestingly, MnII in aqueous solution is oxidized via a single-electron process with redox potential higher than that of natural Mn-SODs. This suggests that, in the cellular environment, MnIII(Me2DO2A) reduction may occur upon reaction with O2• as well as other cytoplasmic reductants. To confirm these assumptions, the antioxidative effects of MnII(Me2DO2A) were completely lost when inactive ZnII was substituted for MnII. Taken together, these features allow MnII(Me2DO2A) to behave as an efficient O2• scavenger. Of note, MnII pentaazamacrocyclic complexes have been shown to react with ONOO⫺ and dismute NO•, albeit at lower rates than with O2• [45]. Whether MnII(Me2DO2A) may also be able to remove harmful ONOO⫺ and excess NO•, thereby increasing its antioxidant properties, remains a tantalizing matter for further investigation.
Conclusions Our study suggests that MnII(Me2DO2A) is a promising member of a new class of SOD-mimetic antioxidant drugs, capable of reducing O2•-mediated cell injury which occurs at reoxygenation after prolonged hypoxia. Further studies on animal models of ischemia and reperfusion are required to corroborate this assumption. In a drug-developing perspective, MnII(Me2DO2A) deserves to be further investigated and exploited to reduce ischemia–reperfusion organ damage in acute vascular diseases, as well as to supplement the incubation fluid of explanted organs and extend their viability in view of transplantation.
Acknowledgments The authors gratefully acknowledge Dr. Eng. Moreno Naldoni, CEO of General Project Ltd., for kind gift of MnII(Me2DO2A). Declaration of interest The authors declare that they have no financial, consulting, and personal relationships with other people or organizations that could bias the present work. This work was supported by research funds from the University of Florence issued to Silvia Nistri. Dr. Barbara Valtancoli is recipient of a research grant from the Ente Cassa di Risparmio di Firenze. References [1] Gerschman R, Gilbert DL, Nye SW, Dwyer P, Fenn WO. Oxygen poisoning and x-irradiation: a mechanism in common. Science 1954;119:623–626. [2] Murphy E, Steenbergen C. Mechanisms underlying acute protection from cardiac ischemia-reperfusion injury. Physiol Rev 2008;88:581–609. [3] McCord JM, Fridovich I. Superoxide dismutase: an enzymatic function for erythrocuprein (hemocuprein). J Biol Chem 1969;244:6049–6055. [4] Fridovich I. Superoxide dismutases. An adaptation to a paramagnetic gas. J Biol Chem 1989;264:7761–7764. [5] Johnson F, Giulivi C. Superoxide dismutases and their impact upon human health. Mol Aspects Med 2005;26:340–352. [6] Finkel T. Radical medicine: treating ageing to cure disease. Nat Rev Mol Cell Biol 2005;6:971–976. [7] Wang ZQ, Porreca F, Cuzzocrea S, Galen K, Lightfoot R, Masini E, et al. A newly identified role for superoxide in inflammatory pain. J Pharmacol Exp Ther 2004;309: 869–878. [8] Salvemini D, Wang ZQ, Zweier JL, Samouilov A, Macarthur H, Misko TP, et al. A nonpeptidyl mimic of superoxide dismutase with therapeutic activity in rats. Science 1999; 286:304–306. [9] Jaeschke H, Woolbright BL. Current strategies to minimize hepatic ischemia-reperfusion injury by targeting reactive oxygen species. Transplant Rev (Orlando) 2012;26:103–114. [10] Murphy MP. How mitochondria produce reactive oxygen species. Biochem J 2009;417:1–13. [11] Poyton RO, Ball KA, Castello PR. Mitochondrial generation of free radicals and hypoxic signaling. Trends Endocrinol Metab 2009;20:332–340. [12] Castello PR, Woo DK, Ball K, Wojcik J, Liu L, Poyton RO. Oxygen-regulated isoforms of cytochrome c oxidase have differential effects on its nitric oxide production and on hypoxic signaling. Proc Natl Acad Sci USA 2008;105:8203–8208. [13] Galkin A, Higgs A, Moncada S. Nitric oxide and hypoxia. Essays Biochem 2007;3:29–42. [14] Kokura S, Yoshida N, Yoshikawa T. Anoxia/reoxygenationinduced leukocyte-endothelial cell interactions. Free Radic Biol Med 2002;33:427–432. [15] Bencini A, Failli P, Valtancoli B, Bani D. Low molecular weight compounds with transition metals as free radical scavengers and novel therapeutic agents. Cardiovasc Hematol Agents Med Chem 2010;8:128–146. [16] Ferrer-Sueta G, Radi R. Chemical biology of peroxynitrite: kinetics, diffusion, and radicals. ACS Chem Biol 2009;4: 161–177.
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