Address correspondence and reprint requests to Bernard Knoops, Group of Cell Biology, Institut des Sciences de la Vie, Université catholique de Louvain, 4-5 Place Croix du Sud, bte L7.07.02, 1348 Louvain-la-Neuve, Belgium. E-mail: email@example.com
Peroxiredoxin-5 (PRDX5) is an antioxidant enzyme which differs from the other peroxiredoxins with regards to its enzymatic mechanism, its high affinity for organic peroxides and peroxynitrite and its wide subcellular distribution. In particular, the mitochondrial isoform of PRDX5 confers a remarkable cytoprotection toward oxidative stress to mammalian cells. Mitochondrial dysfunction and disruption of Ca2+ homeostasis are implicated in neurodegeneration. Growing evidence supports that endoplasmic reticulum (ER) could operate in tandem with mitochondria to regulate intracellular Ca2+ fluxes in neurodegenerative processes. Here, we overexpressed mitochondrial PRDX5 in SH-SY5Y cells to dissect the role of this enzyme in 1-methyl-4-phenylpyridinium (MPP)+-induced cell death. Our data show that mitochondria-dependent apoptosis triggered by MPP+, assessed by the measurement of caspase-9 activation and mitochondrial DNA damage, is prevented by mitochondrial PRDX5 overexpression. Moreover, PRDX5 overexpression blocks the increase in intracellular Ca2+, Ca2+-dependent activation of calpains and Bax cleavage. Finally, using Ca2+ channel inhibitors (Nimodipine, Dantrolene and 2-APB), we show that Ca2+ release arises essentially from ER stores through 1,4,5-inositol-trisphosphate receptors (IP3R). Altogether, our results suggest that the MPP+ mitochondrial pathway of apoptosis is regulated by mitochondrial PRDX5 in a process that could involve redox modulation of Ca2+ transporters via a crosstalk between mitochondria and ER.
Oxidative stress and impaired energy metabolism have been implicated in the pathogenesis of neurodegenerative disorders, such as Parkinson's disease (PD) (Zhou et al. 2008; Schapira and Jenner 2011; Surmeier et al. 2011). This makes mitochondria pivotal players in the commitment of neurons to programmed cell death as they are the major source of ATP and reactive oxygen species (ROS) in neurons. Indeed, reduced activity of mitochondrial complex I, as well as oxidative and nitrosative damage, has been observed in PD patients and is believed to contribute to the loss of dopaminergic neurons (Zhou et al. 2008; Chinta and Andersen 2011).
The only neurotoxin that has been clearly linked to a human form of parkinsonism is 1-methyl-4-phenylpyridinium (MPP+), the active metabolite of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) (Langston et al. 1983). Since this discovery, these neurotoxic species have been used extensively to model PD in a variety of mammalian species and cell lines (Collier et al. 2003; Xie et al. 2010a; Bezard and Przedborski 2011). MPP+ has been shown to inhibit the activity of mitochondrial complex I, causing an acute ATP deficiency with increased ROS and reactive nitrogen species (RNS) production (Chan et al. 1991; Cleeter et al. 1992; Xie et al. 2010b). Activation of caspase-9 and caspase-3 is also observed in several cell types, supporting a mitochondrial apoptotic pathway (Fall and Bennett 1999; Viswanath et al. 2001; Wang et al. 2011). Moreover, exposure to MPP+ increases intracellular Ca2+ concentrations (Chen et al. 1995; Arduino et al. 2009). Considerable attention is dedicated to the disruption of Ca2+ homeostasis during the process of neurodegeneration (Mattson 2007). In particular, it has been reported that ROS and RNS enhance Ca2+ release from endoplasmic reticulum (ER) stores through thiol redox modulation of ryanodine receptors (RyRs) and 1,4,5-inositol-trisphosphate receptors (IP3Rs) (Xia et al. 2000; Hidalgo 2005; Joseph et al. 2006), the two major classes of Ca2+ release channels in neurons (Gleichmann and Mattson 2011). Thus, converging evidence suggests that ER and mitochondria are interconnected in the control of Ca2+-dependent neuronal dysfunction (Csordas and Hajnoczky 2009; Zundorf and Reiser 2011).
Although ROS and RNS are considered destructive to cellular macromolecules in oxidative conditions such as those encountered in neurodegenerative disorders, several cell types use ROS/RNS in the modulation of biological processes, including immune response and apoptosis. Self-protection is therefore crucial and requires a tight regulation of ROS/RNS generation. Regarding the mitochondrial matrix, manganese superoxide dismutase (MnSOD or SOD2) almost instantly dismutates O2·− to H2O2 (Cadenas 2004). Reduction of hydroperoxides and ONOO- in the mitochondria is mainly achieved by two distinct protein families in mammals, glutathione peroxidases (GPXs) and peroxiredoxins (PRDXs) (Cox et al. 2010). Among the six mammalian PRDX members, PRDX3 and PRDX5 are addressed to mitochondria in human and rodents. Recent studies highlight the predominant role of PRDX3 and PRDX5 as peroxide scavengers in mitochondria (De Simoni et al. 2008; Cox et al. 2010; Drechsel and Patel 2010). In addition to their antioxidant function, compelling evidence shows that PRDX members may be also important actors in redox signaling (Neumann et al. 2009; Rhee and Woo 2011; Karplus and Poole 2012). Interestingly, this is also supported by several works that have investigated the neuronal demise in PD models or even PD patients (Qu et al. 2007; Fang et al. 2007; Lee et al. 2008).
PRDX5, the last isoform identified in the PRDX family, possesses unique properties (for review see Knoops et al. 2011). First, PRDX5 is the only mammalian member of the atypical 2-Cysteine (2-Cys) PRDX subfamily. Atypical 2-Cys catalytic mechanism involves the formation of an intramolecular disulfide bond as opposed to the other PRDXs which need a molecular partner to form the disulfide. Second, PRDX5 exhibits a remarkably wide subcellular localization. Indeed, PRDX5 is located in the mitochondria when expressed as a long form with its mitochondrial targeting sequence (L-PRDX5) but a short form (S-PRDX5) also leads to cytosolic, peroxisomal, and nuclear localization of this protein, both mature forms displaying an identical molecular weight (17 kDa) (for review see Knoops et al. 2011). Third, regarding substrate specificity, PRDX5 was shown to react more efficiently with ONOO- and organic peroxides compared with other mammalian PRDXs, suggesting that PRDX5 would be more devoted to the reduction of ONOO- and organic peroxides than the other PRDX isoforms (Dubuisson et al. 2004; Trujillo et al. 2007; Cox et al. 2010).
The protective role of human PRDX5 toward oxidative stress has been demonstrated in several cell lines. Overexpression of PRDX5 in human tendon cells has been shown to reduce apoptosis following H2O2 treatment (Yuan et al. 2004). PRDX5 also protects from apoptosis induced by anti-cancer drugs (Kropotov et al. 2006). Another study reported that PRDX5 prevents TNF-α and PDGF-dependent accumulation of ROS (Seo et al. 2000). Moreover, Chinese hamster ovary cells overexpressing PRDX5 showed reduced cell death and DNA damage after exposure to H2O2 or ter-butylhydroperoxide, with the most effective protection conferred by PRDX5 overexpression in mitochondria (Banmeyer et al. 2004, 2005). Conversely, PRDX5 gene silencing made SH-SY5Y neuroblastoma cells more vulnerable to oxidative stress caused by MPP+ (De Simoni et al. 2008). Studies in human and mouse have also highlighted PRDX5 protective effects under physiopathological conditions (Wang et al. 2001; Plaisant et al. 2003).
The precise role of mitochondrial PRDX5 in neurodegeneration is still unclear. Here, we used human dopaminergic neuroblastoma SH-SY5Y cells overexpressing mitochondrial PRDX5 to investigate in more detail the function of this enzyme in neuronal cell death induced by MPP+. We report that the neuroprotective function of mitochondrial PRDX5 could link mitochondrial redox signaling to the control of Ca2+ release from ER.
Materials and methods
SH-SY5Y cells were from ECACC (European Collection of Cell Culture). Penicillin, streptomycin, and trypsin EDTA were purchased from Life Technologies, Inc. (Gent, Belgium). Specific polyclonal antibody against PRDX5 was obtained as described before (Wang et al. 2001). Rabbit anti-glutamate dehydrogenase was from Rockland (Gilbertsville, PA, USA). Rabbit polyclonal anti-nitrotyrosine was from Upstate (Millipore, 06-284, Overijse, Belgium). Mouse anti-Bax was from BD Pharmingen (BD Biosciences, Erembodegem, Belgium). MPP+, 2-aminoethoxy diphenyl borate (2-APB), 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl tetrazolium bromide (MTT) and rabbit anti-β-actin were purchased from Sigma-Aldrich (Diegem, Belgium). The bicinchoninic acid (BCA) protein assay reagent was from Pierce (ThermoFisher Scientific, Erembodegem, Belgium). ApoAlert Caspase Fluorescent Assay Kit was from BD Pharmingen. Fura-2 acetoxymethyl (AM) ester, 4,6-diamino-2-phenylindole (DAPI), CM-H2DCFDA (5-(and-6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate), and MitoTracker Red CMXROS were obtained from Molecular Probes (Life Technologies, Gent, Belgium). Cytotoxicity Detection Kit (LDH) was from Roche (Brussels, Belgium). Cell Titer-Glo Luminescent Cell Viability Assay (ATP assay) was from Promega (Leiden, The Netherlands). Fluorogenic calpain activity assay kit was provided by Calbiochem (Merck, Darmstadt, Germany). Horseradish peroxidase (HRP)-conjugated goat anti-rabbit IgG and HRP-conjugated rabbit anti-mouse IgG were from DAKO (Heverlee, Belgium). FITC-conjugated donkey anti-rabbit IgG was provided by Jackson ImmunoResearch (Suffolk, England).
Cell culture, stable transfection, and treatment
SH-SY5Y cells were cultured as described previously (De Simoni et al. 2008). pEF-BOS containing the full length human mitochondrial PRDX5 gene was obtained as described before (Banmeyer et al. 2004). This plasmid and its corresponding empty vector were transfected into SH-SY5Y cells using a Nucleofector Instrument (Amaxa Biosystems, Lonza, Braine-l'Alleud, Belgium) according to manufacturer's instructions. Clonal selection was started 3 days after transfection using complete medium containing 0.5 μg/mL puromycin. Individual clones were isolated by dilution to obtain stably transfected cell lines. Despite several PRDX5-overexpressing clones were obtained and were demonstrated to protect against MPP+-induced cell death (see Figure S1), only one was used for subsequent experiments. Cells were exposed to MPP+ for 16 h at the indicated concentrations. In some experiments, cells were pretreated with Nimodipine, Dantrolene, or 2-APB 1 h prior to MPP+ treatment.
In some experiments, mitochondria were loaded with 100 nM MitoTracker Red CMXROS before fixation. Cells were then fixed with 4% formaldehyde. The immunostaining procedure was described by Knoops et al. (1999). Cells were incubated with 1/200 rabbit anti-human PRDX5 polyclonal antibody or 1/50 rabbit anti-nitrotyrosine antibody and finally mounted in Mowiol containing 50 μg/mL DAPI.
Cytosolic and mitochondria enriched fractions were obtained from SH-SY5Y cells with the use of mitochondrial isolation kit according to manufacturer's instructions (Pierce).
Cell lysates were obtained as described previously (Banmeyer et al. 2004) with minor modifications. After treatment, cells were harvested by trypsinization, sonicated in phosphate-buffered saline (PBS) containing 1% of Triton X-100, and centrifuged at 200 g for 10 min. Forty micrograms of soluble proteins were separated on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a nitrocellulose membrane. The blots were probed with 1/4000 rabbit anti-human PRDX5 polyclonal antibody, 1/800 rabbit anti-β-actin, 1/2500 rabbit anti-glutamate dehydrogenase (GDH), or 1/250 mouse monoclonal anti-human Bax followed by HRP coupled secondary antibody and revealed by standard chemiluminescent procedure. Quantifications were performed by densitometric analysis of band intensity (Kodak 1D 3.5 software, Kodak, Zaventem, Belgium). Normalization of PRDX5 protein level was performed with β-actin or GDH.
The LDH assay was carried out as described previously with minor modifications (Banmeyer et al. 2004). Fifty microliters of cellular supernatants from treated and untreated cells was removed for LDH assay. The total LDH release was determined for each clone after cell lysis in 1% Triton X-100. Data were normalized to the value obtained with complete lysis of cells in Triton X-100 considered 100%.
Cellular ATP determination
For cellular ATP measurement, a commercially available kit luciferin-luciferase was used according to manufacturer's protocol. Briefly, after MPP+ treatment in 96-well plates, luciferin substrate and luciferase enzyme were added and bioluminescence was monitored on Fluoroskan. Whole cell ATP content was calculated based on a standard curve, normalized to protein content and expressed as percent of ATP of untreated control cells.
Detection of reactive oxygen species
Intracellular ROS were monitored by using fluorescent probe CM-H2DCFDA. After MPP+ treatment, cells were incubated with 10 μM CM-H2DCFDA in Krebs-Ringer Solution [118 mM NaCl, 5 mM HEPES, 4.7 mM KCl, 1.2 mM MgSO4.7H2O, 1.2 mM KH2PO4, 25 mM NaHCO3, 11 mM glucose, and 1.3 mM CaCl2.2H2O] for 30 min at 37°C. The fluorescence intensity of DCF was measured in a microplate reader (Fluoroskan Ascent Thermo FL, Labsystems, Brussels, Belgium) at λex 485 nm and λem 538 nm.
Image analysis and quantification of 3-nitrotyrosine formation
Cells were immunostained with a rabbit anti-nitrotyrosine antibody (1/50). Fluorescence microscopy was performed with a Reichert-Jung polyvar microscope (Van Hopplynus Instruments, Brussels, Belgium) coupled with a Nikon DS-U1 camera (Nikon Instruments, Amsterdam, The Netherlands). The pictures, obtained under identical exposure conditions, were converted from RGB (red, green, blue) to 8-bit grayscale images and subjected to densitometric analysis (Ferrini et al. 2001) using Zeiss KS400 3.0 software (Carl Zeiss Vision, Zaventem, Belgium). The pixel intensities of whole cells were turned to numerical values where 0 is referring to black and 255 to white. The measurement was performed on three randomly chosen fields (each containing ~30 cells) by slide, three slides by condition, and each condition was reproduced in three independent experiments. Values were normalized to untreated control group considered 100%.
Detection of oxidative mtDNA damage
This assay relies on the ability of certain lesions to hamper the polymerase progression on the template, resulting in the decrease of DNA amplification in the damaged templates compared with undamaged DNA. mtDNA PCR is a reliable assay to determine oxidative damage at mitochondrial genomic level (Banmeyer et al. 2012; Mao et al. 2005). After treatment, DNA was extracted and analyzed for the presence of lesions in mtDNA using long-run PCR to amplify a fragment of 8.2 kbp from mtDNA, located between CYTB and COX2 genes. The yield of long mtDNA PCR product was quantified by real-time PCR using primers hybridizing to the ND4 gene. High molecular weight DNA was isolated with the DNeasy Tissue Kit (Qiagen, Venlo, The Netherlands) according to manufacturer's instructions. Concentration of total cellular DNA was determined and long amplification of mtDNA was performed as described before (Banmeyer et al. 2005). The primer nucleotide sequences were forward primer 5′-ATG GCA CAT GCA GCG CAA GTA GGT CTA CAA G-3′ flanking the region of cytochrome c oxidase subunit II gene (COX2, GenBank accession number: EF488201) and reverse primer 5′-AGG ATT GTT GTG AAG TAT AGT ACG GAT GC-3′ flanking the region of cytochrome b (CYTB, GenBank accession number: EF488201). Quantitative real-time PCR was also performed as described by Banmeyer et al. (2005) with primers hybridizing on NADH dehydrogenase subunit 4 (ND4, GenBank accession number: EF488201): forward primer 5′-ACG CAG GCA CAT ACT TCC TAT TC-3′ and reverse primer 5′-CTT GGG CAG TGA GAG TGA GTA GTA GA-3′. The following Taqman probe labeled with 5′FAM and 3′TAMRA was used: 5′-CCC TTC CCC TAC TCA TCG CAC TAA TTT ACA CTC-3′. Results were expressed in lesions per large PCR fragment and also in relative PCR amplification (Banmeyer et al. 2005).
Assay for caspase-3, caspase-9 and calpain activities
Caspase-3 and caspase-9 activities were determined using ApoAlert Caspase Fluorescent Assay kit (BD Biosciences Pharmingen, Erembodegem, Belgium) and calpain activity using Fluorogenic Calpain Activity Assay kit according to manufacturer's instructions.
DAPI nuclear staining
Following MPP+ treatment, cells were fixed and stained with DAPI as described above. Condensed and fragmented nuclei were considered apoptotic.
Ca2+ imaging experiments
For [Ca2+]i measurement, cells were cultured on 15-mm round glass coverslips. After 16 h MPP+ exposure, cells were loaded with 5 μM Fura-2-AM in assay buffer identical to Krebs-Ringer Solution described above. Rinsed coverslips were then mounted in a heated (37°C) and perfused 100 μL microscope chamber (Warner Instrument Corporation, Hamden, CT, USA). Loaded cells were excited successively at 340 and 380 nm (excitation lights obtained from a Xenon lamp coupled to a monochromator) for 30 ms, and emitted fluorescence was monitored at 510 nm using a CCD camera coupled to an inverted Olympus IX70 microscope (T.I.L.L. Photonics, Martinsried, Germany). The intracellular free [Ca2+] was calculated with the use of the following formula: [Ca2+] (nM) = KdQ (R−Rmin)/(Rmax−R) where R represents the fluorescence intensity ratio Fλ340 nm/Fλ380 nm. Q is the ratio of Fmin to Fmax at λ380 nm and Kd is the Ca2+ dissociation constant of the indicator assigned as 224 nM (Garavito-Aguilar et al. 2004). The minimal fluorescence (Rmin) was determined by perfusing cells with a Ca2+ free solution in which CaCl2 was omitted and 200 μM EGTA and 5 μM ionomycin were added. Five millimolar CaCl2 was added to assay buffer to saturate Fura-2 with calcium and to calculate the maximal fluorescence (Rmax). Autofluorescence was measured after the addition of 20 mM MnCl2. Fluorescence intensities from single cells excited at the two wavelengths were recorded separately and corrected for the background using the software TILLvisION v3.3 (T.I.L.L. Photonics, Martinsried, Germany).
Inactivation of endogenous PRDX5 expression by small hairpin RNA (shRNA)
The plasmid pSILENCER used for expressing a shRNA designed to interfere with human PRDX5 was obtained as described previously (De Simoni et al. 2008). The disruption of endogenous PRDX5 expression was confirmed seven days post-transfection by immunofluorescence (Fig. 5) or western blotting (see De Simoni et al. 2008).
Cell viability was assessed by MTT as described before with minor modifications (Banmeyer et al. 2004). After treatment, medium from 96 well plates was removed, plates were rinsed with PBS, and 50 μL of MTT solution (1 mg/mL in complete medium) was added. After incubation (1.5 h at 37°C), medium was removed and formazan crystals formed in metabolically active cells were dissolved in dimethylsulfoxide (DMSO) (50 μL/wells). The absorbance of formazan was read at 570 nm. Data were normalized to untreated group considered as 100%.
All measurements result from replicates within an experiment, and each experiment was repeated at least three times. Means and standard errors (SEM) were calculated either from the replicates or from the independent experiments (as indicated). Statistical analysis was performed using Student's t-test or two-way anova followed by Bonferroni post-hoc test (as indicated). Significance versus control is represented by *p <0.05, **p <0.01, ***p <0.001.
Overexpression of mitochondrial PRDX5 in SH-SY5Y cells
The construct coding for human mitochondrial PRDX5 in pEF-BOS plasmid was previously described by Banmeyer et al. (2004). Stable SH-SY5Y cell clones containing pEF-BOS plasmid or mito-PRDX5-pEF-BOS plasmid were obtained by nucleofection and puromycin selection. These clones were named control and mito-PRDX5, respectively. PRDX5 overexpression was confirmed by immunofluorescence (Fig. 1a) and expression level was estimated by western blotting (Fig. 1b–d). Immunoblot of total cell lysates revealed a twofold overexpression of PRDX5 in the mito-PRDX5 clone (Fig. 1b). Subcellular fractionation confirmed that the increased levels of PRDX5 were mainly localized into mitochondria (Fig. 1c).
Mitochondrial PRDX5 prevents mitochondria-dependent apoptosis induced by MPP+
To assess the effects of mito-PRDX5 overexpression toward MPP+ neurotoxicity, we treated control and mito-PRDX5 cells with increasing concentrations of MPP+ for 16 h. The incubation period and the concentrations used were determined according to previous studies using SH-SY5Y cells (Fall and Bennett 1999; De Simoni et al. 2008; Sorensen et al. 2009; Xie et al. 2011) and according to preliminary results (Fig. 2a and Figure S2). First, cell death was estimated using a LDH release assay and by measuring ATP concentration (Fig. 2a and b). Compared with control clone, overexpression of PRDX5 in mitochondria resulted in a marked reduction of LDH release (Fig. 2a) as well as the stabilization of ATP levels to 78% at 10 mM MPP+ (Fig. 2b). To test the antioxidant function of mitochondrial PRDX5, ROS and RNS production by MPP+ was also determined. Our results show that DCF fluorescence was reduced in mito-PRDX5 cells compared with control cells (Fig. 2c). Moreover, higher levels of 3-nitrotyrosine were detected in control cells compared with mito-PRDX5 cells at 5 and 10 mM MPP+ (Fig. 2d). Although these differences were not significant according to the conservative Bonferroni statistical post-hoc test, two-way anova resulted in a significant clone effect (p =0.0164). Note that only 3-nitrotyrosine levels are unchanged in mito-PRDX5 cells under basal conditions (0 mM MPP+). This could be explained by the fact that peroxynitrite precursors ·NO and O2·− only reach significant levels following exposure to MPP+. Indeed, in non-treated cells, this peroxynitrite formation is not favored because of the rapid dismutation of O2·− accelerated through catalysis by superoxide dismutase 2 and because of the diffusion/consumption of ·NO (for review see Radi 2004).
To investigate the role of mito-PRDX5 in the mitochondrial pathway of cell death triggered by MPP+, we performed different experiments. MtDNA being a critical target of ROS and RNS (Ott et al. 2007), we evaluated the frequency of mtDNA lesions in control and mito-PRDX5 cells by a two-step PCR (Banmeyer et al. 2005). The data presented in Fig. 3a reveal a genoprotective role of mitochondrial PRDX5 toward mtDNA in SH-SH5Y cells exposed to MPP+. Nuclear condensation/fragmentation occurring in apoptosis was also examined (Fig. 3b). The percentage of apoptotic nuclei increased with MPP+ concentration and mito-PRDX5 overexpression strongly reduced this rise of apoptosis. Then, we measured the activities of caspase-3 and caspase-9, the two major caspases associated with mitochondrial apoptotic pathway (Yuan et al. 2003). Caspase-3 and caspase-9 were activated following MPP+ treatment in control cells, in a dose-dependent manner for caspase-3 (Fig. 3c) and with a peak at 5 mM MPP+ for caspase-9 (Fig. 3d). Caspase-3 and caspase-9 activation was inhibited by overexpression of mitochondrial PRDX5.
Mitochondrial PRDX5 inhibits Ca2+-dependent apoptosis induced by MPP+
Besides the mitochondrial pathway of apoptosis, it has also been demonstrated that calpains, which are Ca2+-dependent cysteine proteases, are central in MPP+-induced cell death (Harbison et al. 2011). Under our experimental conditions, the intracellular increase in Ca2+ after MPP+ treatment was abolished when mitochondrial PRDX5 was overexpressed (Fig. 4a). Accordingly, while calpain activity was upregulated up to seven times in control cells after a treatment of 10 mM MPP+, overexpression of mito-PRDX5 maintained this activity at a basal level (Fig. 4b). Moreover, the specific cleavage of the pro-apoptotic protein Bax (21 kDa) to truncated Bax (tBax, 18 kDa), which has been attributed to calpain activation (Wood et al. 1998), was examined. This cleavage has been shown to enhance apoptogenic properties of Bax (Wood and Newcomb 2000; Toyota et al. 2003). We detected Bax profile by western blotting analysis of whole cell lysate from MPP+ treated cells (Fig. 5a). In absence of MPP+, the only band observed in control and mito-PRDX5 cells was the 21 kDa protein. Treatment with 10 mM MPP+ resulted in a predominant 18 kDa fragment of Bax (tBax). In contrast, the proteolytic cleavage of Bax taking place during MPP+ treatment was strikingly reduced by mitochondrial PRDX5 overexpression. This was further investigated using RNA interference. As shown in Fig. 5b, transient transfection of a shRNA directed to PRDX5 transcripts in mito-PRDX5 cells gave rise to a decrease of overall PRDX5 expression level. The decreased expression of PRDX5 was sufficient to restore the calpain-dependent cleavage of Bax after MPP+ treatment (Fig. 5a).
Ca2+ release occurs through IP3R during MPP+ cell death
The intracellular Ca2+ increase occurring in neurodegenerative processes arises through altered regulation of Ca2+ channels present in the plasma membrane but also in mitochondria and ER (Zundorf and Reiser 2011; Gleichmann and Mattson 2011). To determine the source of Ca2+ entry following exposure to MPP+, we used different calcium channel inhibitors before MPP+ treatment and measured cell viability by MTT assay (Fig. 6). We used Nimodipine to inhibit L-type voltage-dependent calcium channels (VDCCs) in the plasma membrane (Li et al. 2009), Dantrolene to inhibit RyRs in the ER (Muehlschlegel and Sims 2009), and 2-APB (2-aminoethoxydiphenyl borate) as inhibitor of IP3Rs in the ER (Hagenston et al. 2009). Although Nimodipine (Fig. 6a) and Dantrolene (Fig. 6b) pretreatments did not rescue cells from apoptosis but rather worsened mortality following MPP+ exposure, 2-APB pretreatment led to a significant increase in cell viability compared with non-pretreated cells (Fig. 6c). Moreover, inhibition of IP3R Ca2+-channels by 2-APB in the ER nearly totally prevented the increase in [Ca2+]i provoked by MPP+ toxicity (Fig. 7a). Under these conditions, the number of apoptotic cells also dropped after pretreatment with 25 μM 2-APB (Fig. 7b). These results indicate that MPP+ cell death not only follows the mitochondrial pathway of apoptosis but also involves Ca2+ release from the ER through IP3R. Importantly, blocking this Ca2+-channel had a similar effect on [Ca2+]i and on the percentage of apoptotic cells as the overexpression of mitochondrial PRDX5 (Fig. 4a and 3b, respectively).
We previously reported that PRDX5 silencing in SH-SY5Y cells increased vulnerability to MPP+-induced neurodegeneration (De Simoni et al. 2008). In this study, we focused on the protective function of the mitochondrial form of PRDX5 in this process, to clarify the molecular events that link this mitochondrial enzyme to the control of Ca2+-dependent apoptosis induced by MPP+.
We first showed that overexpression of mitochondrial PRDX5 protects SH-SY5Y cells against the ROS/RNS-dependent neuronal death induced by MPP+. This confirms previous studies that have shown that PRDX5 is able to prevent oxidative cell death (Yuan et al. 2004; Banmeyer et al. 2004, 2005; De Simoni et al. 2008). In particular, the mitochondrial form of this enzyme has been demonstrated to provide the highest level of cytoprotection against H2O2 compared to its cytosolic, peroxisomal, and nuclear forms (Banmeyer et al. 2004). Here, we focused on a neuronal nitro-oxidative stress located at the mitochondria which could mimic the endogenous mitochondrial disturbance occurring in PD. It was recently shown that PRDX5 is barely expressed in dopaminergic neurons compared with other neuronal populations in mice (Goemaere and Knoops 2012). The low levels of PRDX5 in these neurons probably compromise redox homeostasis, especially when a mitochondrial nitro-oxidative stress occurs, like in PD. The overexpression study presented here highlights the importance of mitochondrial PRDX5 in the resistance of dopaminergic neurons to mitochondrial oxidative neurotoxicity.
This study also supports that ROS/RNS generation at the mitochondria is a primary event in the MPP+-induced cell death. Indeed, when this step is prevented by mitochondrial PRDX5 overexpression, the downstream molecular events leading to cell death are also inhibited. In particular, the protection conferred by mito-PRDX5 toward mtDNA damage and caspase-9 activation reinforces the idea of a mitochondrial oxidative stress and subsequent mitochondria-dependent pathway of apoptosis in this cellular model of PD. The fact that PRDX5 can reduce both ROS and RNS probably contributes to the strong effect we observed. Our results support that PRDX5 holds a special place among mitochondrial antioxidant enzymes, such as PRDX3, GPX1 and GPX4. Indeed, PRDX5 is able to reduce peroxides with high rate constants, especially with ONOO− (107 M−1s−1) and organic peroxides (106 M−1s−1), whereas its kinetic reaction with H2O2 is slower (105 M−1s−1). It has been proposed that this efficiency toward ONOO- and organic peroxides confer a specific role to mitochondrial PRDX5 (Cox et al. 2010; Knoops et al. 2011).
In addition to caspase activation, our results point to calpain-dependent cell death. This is in agreement with previous studies showing an increase of [Ca2+]i and calpain activation in response to MPTP/MPP+ treatment (Lee et al. 2006; Harbison et al. 2011). More importantly, calcium dysregulation and increased proteolytic activity of calpains have been reported in PD patients (Mouatt-Prigent et al. 1996; Crocker et al. 2003; Blandini et al. 2009). In our study, intracellular calcium increase and calpain activation by MPP+ were further confirmed by the specific calpain-dependent cleavage of Bax. These Ca2+-dependent events were all abolished by mitochondrial PRDX5 overexpression, meaning this enzyme acts upstream of calpain stimulation by Ca2+ and may play a role in the control of Ca2+ release in the cell. Ca2+ entry in neuronal cells depends either on entry from extracellular matrix through the plasma membrane or from intracellular stores in ER and mitochondria (Mattson 2007) (see Fig. 8). We showed that inhibition of the ER Ca2+ receptor IP3R by 2-APB protected neurons against apoptosis induced by MPP+ in a similar way to mito-PRDX5 overexpression. Interestingly, key calcium transporters in ER and mitochondria, including IP3R, are regulated by redox modifications (Csordas and Hajnoczky 2009). Furthermore, growing attention is dedicated to the communication between mitochondria and ER to control ROS and Ca2+ signaling pathways, in particular during neurodegeneration (Csordas and Hajnoczky 2009; Arduino et al. 2009; Gleichmann and Mattson 2011). In light of these studies, one can reasonably propose that PRDX5 mitochondrial overexpression in our MPP+ model allows the removal of mitochondrial ROS/RNS which are necessary to induce redox activation of IP3R and subsequent Ca2+ release in the cytosol to pursue apoptosis (Fig. 8). Although 2-APB has been widely used as an IP3R inhibitor, several reports have demonstrated that this compound is not totally IP3R specific as it could also modulate store-operated Ca2+ channels (SOCs), SERCA pumps and transient receptor potential (TRP) channels (Peppiatt et al. 2003; Hagenston et al. 2009) (Fig. 8). Nevertheless, SOCs are only activated following [Ca2+]ER depletion (Gleichmann and Mattson 2011), SERCA pumps are responsible of refilling ER with Ca2+ and TRP channels are not inhibited but seem to be rather activated by 2-APB (Hu et al. 2004). It is therefore likely that interference of intracellular Ca2+ increase caused by 2-APB in our study is essentially because of IP3R inhibition. Although 2-APB provoked a drastic decrease in [Ca2+]i in this study, it would also be of interest to investigate the effect of mitochondrial PRDX5 overexpression on mitochondrial Ca2+ release, in particular through the mitochondrial permeability transition pore (PTP), which opens during neurodegenerative processes (Gleichmann and Mattson 2011) (see Figure S3 and discussion in Supporting Information).
Employing MPP+, which in vivo causes dopaminergic loss resembling the neuropathology of PD, this study demonstrates that the potent protective function of mitochondrial PRDX5 occurs early during neurodegeneration, that is, before oxidative-dependent dysfunction of mitochondria. Our results also support that PRDX5 function can be viewed beyond its antioxidant properties stricto sensu. First, scavenging of peroxides and ONOO− by PRDX5 counters direct nitro-oxidative damage to macromolecules such as mtDNA and thus also possibly to mitochondrial pore proteins that allows cytochrome c release and caspase activation. Moreover, PRDX5 may modulate redox signaling between mitochondria and ER-located IP3R Ca2+ transporter to control Ca2+-dependent activation of calpains. This latter function of PRDX5 reveals a novel molecular link between ROS/RNS generation and Ca2+ dysregulation in neurodegenerative processes.
The authors have no conflict of interest to declare. The authors thank Mr Marc Pirson for helpful reading and corrections to the manuscript. This study is supported by the « Fonds pour la formation à la Recherche dans l'Industrie et dans l'Agriculture » (F.R.I.A.), by the « Fonds National de la Recherche Scientifique » (F.N.R.S), by the « Communauté française de Belgique-Actions de Recherche Concertées » (10/15-026), and by the DIANE research program of the Walloon region.