PC12 cells (a cellular line of rat pheochromocytoma from Sigma-Aldrich Co., St. Louis, MO, USA. Cat. no. 88022401) were cultured in a 5% CO₂ atmosphere at 37°C in RPMI with 10 mM HEPES, 1.0 g/l glucose, 3.7 g/l NaHCO₃, 100 U/ml penicillin, 100 µg/ml streptomycin, 10% fetal calf serum and 15% horse serum. Once grown to 85% confluence, the cells were subcultured at an appropriate density according to each experimental procedure.

The cells were treated with a physiological concentration (60 µM) of DHA (purchased from Sigma-Aldrich Co.) 24 h prior to and during H₂O₂ exposure (for a further 24 h). For the control groups, medium or H₂O₂ alone were used where appropriate. DHA was dissolved in 99.5% ethanol to prepare a 500 mM stock solution; prior to use, DHA was complexed (1:2) to bovine serum albumin (BSA) and diluted in RPMI to the required concentration. The concentration of H₂O₂ used was 300 µM; since, in a previous series of dose-response experiments, described in detail in (14), we demonstrated that after 24 h, H₂O₂ resulted in a reduction of the viability of PC12 cells of 50% at this concentration. At the end of all the experiments, the cells were collected and stored at 37°C (unless otherwise stated) for the analytical experiments.

For the determination of cell viability and ROS production, the PC12 cells were plated in 96-well plates at a density of 10,000 cells/well and incubated with DHA 24 h prior to exposure to H₂O₂. Cell survival was evaluated by the 3-[(4,5-dimethylthiazol-2-yl)- 5,3-carboxymethoxyphenyl]- 2- (4- sulfophenyl)-2H tetrazolium, inner salt reduction assay (MTS). The MTS assay (Promega srl, Padova, Italy) is a sensitive measurement of the normal metabolic status of cells, which reflects early cellular redox changes. The intracellular soluble formazan produced by the cellular reduction of the MTS was determined by recording the absorbance of each 96-well plate using the automatic microplate photometer (BioTek™ Elx800 -Box 998; BioTek Instruments, Winooski, VT, USA) at a wavelength of 490 nm. The morphological features of the PC12 cells subjected to the different treatments were analyzed and photographed by phase-contrast microscopy (Nikon eclipse ts100 microscope; Nikon, Tokyo, Japan), at a magnification of x20.

The detection of ROS production was performed using the 2′,7′-dichlorofluorescein diacetate (DCFDA)-Cellular ROS Detection Assay kit (Abcam, Cambridge, UK). DCFDA is initially non-fluorescent and is converted by oxidation to DCF, a highly fluorescent compound. DCF was quantified using a CytoFluor Multiwell Plate Reader (Victor 3 Wallac 1420; Perkin Elmer Waltham, MA, USA), with 485 nm excitation and 538 nm emission filters. ROS production was expressed as the fluorescence intensity and presented as a percentage relative to the control.

The cell cultures (1×10⁶) were washed twice in phosphate buffered saline (PBS, pH 7.4) and subsequently centrifuged at 1,890 × g for 10 min at 4°C, producing a cell pellet and the supernatant. Cell pellets were processed for the determination of the enzymatic activity of total glutathione peroxidase (GSH-Px), catalase and total superoxide dismutase (SOD), by adding 500 µl/10⁶ cells of a hypotonic buffer containing 15 mM KCl + 1 mM KH₂PO₄, pH 7.4. After vigorous vortexing for 60 sec, the cell suspensions were sonicated at 4°C for 3 cycles of 20 sec each with 20 sec intervals between each pulse of sonication. Subsequently, the samples were centrifuged at 20,690 × g for 15 min at 4°C to remove any cell debris. Enzymatic activities were performed spectrophotometrically (using an Agilent 89090A spectrophotometer; Agilent Technologies, Santa Clara, CA, USA) on cell lysates. Total GSH-Px activity was assayed as previously described (15). Briefly, 1 ml of reaction mixture had the following composition: 0.1 M Tris-HCl, pH 8.0, 4 mM EDTA, 2 mM reduced glutathione (GSH), 1 IU glutathione reductase, 0.3 mM NADPH and 10 µl of cell extract. Following 10 min of incubation at 37°C, the reaction began by the addition of 70 µM tert-butyl hydroperoxide (TBH) and monitored spectro-photometrically (Agilent 89090A) at 340 nm by following for 10 min NADPH oxidation; blank without TBH and additional blank without cell lysate were used. The millimolar extinction coefficient of 6.3×10^-3 of NADPH was used for the calculation. Catalase activity was examined as previously described (15); the blank reaction mixture (1 ml final volume) was composed by 10 mM KH₂PO₄ pH 7.4 and H₂O₂ 10 mM; the reaction began by the addition of 5 µl of cell extract and monitored spectrophotometrically (Agilent 89090A) at 240 nm by following H₂O₂ dismutation. The millimolar extinction coefficient of 3.94×10^-3 of H₂O₂ was used for the calculation. Total SOD activity was analyzed as previously described (16). The reaction mixture (1 ml final volume) was formed by 50 mM KH₂PO₄ pH 7.8, 10 µM oxidized cytochrome c, 1.25 mM xanthine, 0.006 µM xanthine oxidase and 4 mM sodium azide (to inactivate catalase). The reaction began by the addition of 20 µl of cell extract and monitored spectrophotometrically (Agilent 89090A) at 550 nm following the cytochrome c reduction. The millimolar extinction coefficient of 8.4x10^-3 of oxidized cytochrome c was used for the calculation. All enzymatic activities were normalized for the protein content determined by Bradford assay (17) and expressed as IU/mg of protein (1 IU=1 µmol substrated consumed/min). In a different set of experiments, cell pellets, obtained as described above, were treated for the determination of non-enzymatic antioxidant levels with a precipitating solution composed of CH₃CN 75% and KH₂PO₄ 25% (10 mM) at pH 7.4, and then centrifuged at 20,690 × g for 15 min at 4°C. The supernatants were collected and subjected to 2 chloroform washings in order to obtain an upper aqueous phase that was directly injected onto the HPLC. Each sample was analyzed to determine the concentration of GSH, ascorbate, malondialdehyde (MDA), nitrites and nitrates (NO₂ and NO₃) according to an ion-pairing HPLC method previously set up in our laboratories (18,19). All concentration values were normalized with respect to the protein amount and expressed as nmol/mg protein (17).

The cells were seeded at a density of 1×10⁶ cells/well on a 24-well plate and incubated with DHA 24 h prior to the exposure to H₂O₂ (300 µM for 24 h). The medium was then aspirated, and the cells were washed with PBS. After the cells were incubated at room temperature, for 15 min in a dark room with Annexin V-FITC (TACS Annexin V kit; Trevigen, Gaithersburg, MD, USA). The cells were then examined under a fluorescence microscope (Nikon Eclipse TE300 Inverted Fluorescence Microscope; Nikon) with a ×20 objective lens.

ΔΨm, an early marker of the induction of apoptosis, was assessed using the Mitolight™ Apoptosis Detection kit (APT142; Chemicon International, Temecula, CA, USA). The cells (4×10⁴ cells/well in a PLL-coated black-bottom 96-well plate) (treated as described above) were incubated with 100 µl of prediluted Mitolight™ dye solution for 15 min (37°C, 5% CO₂, 95% air). In healthy cells, the dye accumulates and aggregates in the mitochondria, giving off a bright red fluorescence (λem=585 nm). In apoptotic cells with an altered ΔΨm, the dye in monomeric form remains in the cytoplasm, fluorescing green (λem=530 nm) and thus providing a ready discrimination between apoptotic and non-apoptotic cells. The fluorescence intensities were measured with a Victor multilabel plate reader (PerkinElmer, Waltham, MA, USA) at an excitation wavelength 485 nm and an emission wavelength 530 nm to monitor apoptotic cells and 585 nm for healthy cells, respectively. The data were expressed as a fluorescence ratio (585/530) representative of the arbitrary ΔΨm. The cells were also observed using a Nikon Eclipse TE300 Inverted Fluorescence Microscope (Nikon) (data not shown).

Total RNA was isolated using the SV Total RNA Isolation System (Promega srl). The RNA concentration was evaluated by spectrophotometric reading at 280 and 260 nm. Total RNA was used for first strand cDNA synthesis with the HyperScript, First strand Synthesis kit and Oligo-dT, as random primer (GeneAll Biotechnology Co. Ltd., Seoul, Korea). PCR was performed with about 150 ng of cDNA using PCRBIO Classic Taq (PCR Biosystems Ltd., London, UK). The following primer sequences were used for amplification: Actin β (β-actin) forward, 5′-TGCTATGTTGCCCTAGACTTCG-3′ and reverse, 5′-GTTGGCATAGAGGTCTTTACGG-3′ (240 bp); Bax forward, 5′-GCAGGGAGGATGGCTGGGGAG-3′ and reverse, 5′-TCCAGACAAGCAGCCGCTCACG-3′ (352 bp); Bcl2 forward, 5′-CACCCCTGGCATCTTCTCCT-3′ and reverse, 5′-GTTGACGCTCCCCACACACA-3′ (349 bp); NFE2L2 forward, 5′-GCCAGCTGAACTCCTTAGAC-3′ and reverse, 5′-GATTCGTGCACAGCAGCA-3′ (466 bp); and HO-1 forward, 5′-GAAACAAGCAGAACCCAGTC-3′ and reverse, 5′-AGAGGTCACCCAGGTAGCG-3′ (225 bp).

The experimental protocols, in accordance with the conditions reported in the literature (14,20) and slightly modified, were as follows: An initial denaturation for 3 min at 95°C; amplification for 21 (β-actin), 20 (NFE2L2), 30 (HO-1) and 40 cycles (Bax and Bcl2) of denaturation, 15 sec, 95°C, annealing: 15 sec, at 61°C (β-Actin), 58°C (NFE2L2), 55°C (HO-1), 60°C (Bax), 59°C (Bcl2) and elongation at 72°C for 1 min; and a final elongation for 10 min at 72°C. The PCR products were then analyzed by 1.5% agarose gel electrophoresis in TBE1X buffer. Image acquisition and product analysis were made using Bio-Rad imaging systems with Quantity One^® 1-D analysis software.

The detection of intracellular Bax, Bcl2 and NFE2L2 proteins was performed using colorimetric cell-based ELISA kits from Assay Biotechnology (Sunnyvale, CA, USA) for Bax (cat. no. CB5066) and for Bcl2 (cat. no. CB5068) and from LSBio/LifeSpan Biosciences, Inc. (Seattle, WA, USA) for NFE2L2 (cat. no. LS-F2192). In brief, the cells were seeded at 3×10⁴ cells/well in a 96-well dish and treated according to the experimental protocol; at the end of the experiment, the cells were fixed with 4% formaldehyde. Successively, quenching buffer, blocking buffer and finally primary antibodies (rabbit anti-Bax, rabbit anti-Bcl2, rabbit anti-NFE2L2 according to the experimental design, or mouse anti-GAPDH; the latter used as a positive internal control) were added followed by incubation overnight at 4°C. All primary antibodies were diluted 1:100 according to the Technical Manual (the antibodies were included with the kit). Following this overnight incubation, the secondary antibodies (included with the kit; anti-rabbit IgG for Bax, Bcl2 and NFE2L2; Anti-mouse IgG for GADPH) conjugated to peroxidase were added and subsequently the samples were read with a microplate reader (BioTek™ Elx800 -Box 998; BioTek Instruments) [optical density (OD) at 450 nm]. The positive controls were performed in the same plates with the target experiments. All values obtained, normalized to GAPDH OD450, were expressed as a percentage relative to the control (untreated cells).

For the intracellular detection of HO-1, the PC12 cells were examined using a kit sandwich ELISA (Abcam, Cambridge, UK, cat. no. ab213968). The ELISA was performed according to the manufacturer’s protocol. Briefly, 1x10⁶ cells, after treating, were lysed (using 1 ml extraction reagent, provided with the kit, supplemented with protease inhibitors) and centrifuged at 10,000 × g, at 4°C for 10 min. The collected supernatants, following protein measurement and normalization, were plated into plates precoated (with monoclonal anti-rat HO-1 antibody, ready to use). HO-1 immobilization was detected with a specific polyclonal antibody and successively with a secondary antibody (Rat HO-1, ready to use) conjugated to the peroxidase. The intensity of the color was measured on a microplate reader (BioTek™ Elx800 -Box 998; BioTek Instruments) at 450 nm. To generate the standard curve, provided rat HO-1 standard (5 µg/ml) was stepwise diluted (25, 12, 5, 6, 5, 3, 13, 1, 56, 0, 78, 0, 39 and 0 ng/ml) and the readers were plotted (using a linear scale) and thus: Concentrations (ng/ml) on the x-axis and absorbance measurements on the y-axis. HO-1 concentrations from the samples were quantified by interpolating the absorbance readings from a standard curve. The data were expressed as a percentage relative to the control (untreated cells).

Each experiment was repeated at least 3 times, separately. All the results are presented as the means ± SEM of (n) replicates per experimental group. Data were subsequently analyzed by one-way ANOVA, followed by the post hoc Newman-Keuls test for comparisons between group means, using a Prism™ computer program (GraphPad, San Diego, CA, USA). Differences were considered statistically significant at P<0.05.

In preliminary experiments, the protective effects of DHA against H₂O₂-induced oxidative stress were investigated in the pheochromocytoma cell line, PC12, by MTS assay. The cells were pre-treated with physiological concentrations of DHA (60 µM) for 24 h and then treated with H₂O₂ for a further 24 h, with the exception of the control groups. A significant and consistent decrease (approximately 40%) in cell viability was observed following 24 h of exposure to H₂O₂. Under these conditions, pre-treatment with DHA was able to antagonize the effects of H₂O₂. DHA alone did not affect cellular viability in the same experimental paradigm (Fig. 1A).

The cell viability data were also confirmed by subsequent morphological experiments conducted in the same experimental paradigm (Fig. 1B). The number of cells was markedly reduced and many of these had a rounded appearance following exposure to 300 µM H₂O₂ (Fig. 1B, panel c), compared to the controls and to cells treated with DHA alone (Fig. 1B, panels a and b); pre-treatment with DHA exerted potent protective effects against H₂O₂-induced oxidative damage (Fig. 1B, panel D).

A second series of experiments were performed to ascertain whether the protective effects observed post-DHA treatment were associated with the reduction of ROS production and oxidative damage. Under these conditions, pre-treatment (24 h) with DHA was able to reduce ROS production induced by H₂O₂, while it was not able to alter the intracellular ROS levels under baseline conditions (Fig. 1C).

Additional experiments were performed to investigate the biochemical mechanisms underlying the antioxidant effects of DHA: Enzymatic (SOD, catalase and GSH-Px) and non-enzymatic (GSH and ascorbic acid) antioxidants, an indicator of lipid peroxidation (MDA) and nitrates and nitrites were measured in the presence or absence of DHA under basal conditions and in response to oxidative stress. The mean concentration levels of enzymatic antioxidants were markedly lower in the cells exposed to H₂O₂ than in the controls (Table I). DHA pre-treatment increased the intracellular levels of SOD and GSH‑Px in a significant manner under basal conditions and following H₂O₂-induced oxidative stress (Table I). The catalase levels were increased by DHA pre‑treatment under oxidative stress conditions compared to H₂O₂ exposure alone, although the difference was not statistically significant. The catalase basal intracellular concentrations remained unaltered (P>0.05) in the presence of DHA (Table I).

In addition, Table II shows the effects of DHA pretreatment on GSH, ascorbic acid, MDA, nitrates and nitrites under basal and oxidative conditions. It was observed that: i) DHA significantly increased the GSH levels both under baseline conditions and in the presence of H₂O₂; ii) DHA increased the intracellular concentrations of ascorbic acid and decreased those of MDA in a statistically significant manner only in the presence of H₂O₂; iii) the intracellular levels of nitrates and nitrites remained unaltered following pre‑treatment with DHA under both experimental conditions. The results obtained above support the hypothesis that DHA exerts antioxidant effects on H₂O₂-exposed PC-12 cells by modulating the levels of antioxidant enzymes and non‑enzymatic scavengers, which regulate the levels of ROS.

PC12 cell morphology was observed by fluorescence microscopy using an Annexin V staining assay to detect apoptotic cells following pre-treatment with DHA in the presence or absence of oxidative damage. As shown in Fig. 2A, the apoptosis of the cells pre-treated with DHA was visibly decreased compared with that of the cells exposed to H₂O₂ alone. The process of apoptosis is strongly associated with the loss of ΔΨm in different cells. Indeed, the opening of the mitochondrial permeability transition pore induces the depolarization of the transmembrane potential, and the release of apoptotic factors and the loss of oxidative phosphorylation. Consequently, in this study, in order to determine whether DHA inhibits H₂O₂-induced apoptosis related to the mitochondrial pathway, PC12 cells were pre-treated with DHA and exposed to H₂O₂ to induce mitochondrial dysfunction. The ΔΨm of the cells was reduced with H₂O₂ alone (60% less than the control), but was enhanced follwoing pre-treatment (24 h) with DHA in a statistically significant manner (Fig. 2B). These results confirmed our hypothesis that DHA is able to inhibit H₂O₂-induced mitochondrial dysfunction in PC12 cells.

To confirm this hypothesis further, the subsequent experiments were aimed to investigate the effects of DHA pre-treatment on Bax and Bcl2 gene and protein expression levels. As shown (in Fig. 2C and D, left panel), the expression of the pro-apoptotic gene, Bax, was increased in the H₂O₂-exposed cells, while DHA pre-treatment markedly reduced its expression. On the contrary, the expression of anti-apoptotic gene, Bcl2, was decreased following H₂O₂ exposure, while the gene expression was reversed in the cells pre-treated with DHA (Fig. 2C and D, right panel). The same trend was also observed for the protein levels of Bax and Bcl2 in the same experimental paradigm (Fig. 2E).

To elucidate the plausible signal transduction pathways involved in the DHA-mediated protective effects, we examined the involvement of NFE2L2, the principal transcription factor that regulates the basal and inducible expression of a number of antioxidant genes. In order to provide direct evidence of the involvement of NFE2L2 activation, the PC12 cells were pre-treated with DHA for 24 h, followed by exposure to H₂O₂ for an additional 24 h. We observed in our experimental paradigm that H₂O₂ markedly downregulated NFE2L2 mRNA expression, which was antagonized by pre-treatment with DHA (Fig. 3A, left and middle panels). Notably, DHA was effective only in the presence of oxidative damage; in fact, pre-treatment with DHA alone did not stimulate NFE2L2 gene expression, but rather decreased it (Fig. 3A, left and middle panels). The results obtained were also confirmed by subsequent experiments conducted to determine the NFE2L2 protein intracellular levels. In particular, the NFE2L2 protein levels increased >3-fold following DHA pre-treatment both under basal conditions and following H₂O₂-induced oxidative damage (Fig. 3A, right panel).

The transcriptional activation of NFE2L2 evokes the activation of a series of phase II detoxifying enzymes, including HO-1, which is able to protect cells against oxidative damage. Therefore, to further investigate whether DHA-induced NFE2L2 activation modulate in turn HO-1, we examined the effects of DHA pre-treatment on the activation of the NFE2L2/HO-1 pathway in PC12 cells in same experimental paradigm. The intracellular mRNA expression and protein levels of HO-1 were markedly decreased to an estimated 50% following exposure to H₂O₂ in comparison to the untreated cells. The observed effect was markedly inhibited by DHA pre-treatment (24 h), which increased the mRNA expression and protein levels of HO-1 to values equal or higher than those observed in the untreated cells, respectively (Fig. 3; pan B). Under baseline conditions, the DHA alone significantly increased the HO-1 protein levels, while it had no effect on the mRNA expression of the enzyme (Fig. 3B).