The present study examined whether the antidepressant paroxetine promotes the survival of nigrostriatal dopaminergic (DA) neurons in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) mouse model of Parkinson’s disease. MPTP induced degeneration of nigrostriatal DA neurons and glial activation as visualized by tyrosine hydroxylase, macrophage Ag complex-1, and/or glial fibrillary acidic protein immunoreactivity. Real-time PCR, Western blotting, and immunohistochemistry showed upregulation of proinflammatory cytokines, activation of microglial NADPH oxidase and astroglial myeloperoxidase, and subsequent reactive oxygen species production and oxidative DNA damage in the MPTP-treated substantia nigra. Treatment with paroxetine prevented degeneration of nigrostriatal DA neurons, increased striatal dopamine levels, and improved motor function. This neuroprotection afforded by paroxetine was associated with the suppression of astroglial myeloperoxidase expression and/or NADPH oxidase-derived reactive oxygen species production and reduced expression of proinflammatory cytokines, including IL-1β, TNF-α, and inducible NO synthase, by activated microglia. The present findings show that paroxetine may possess anti-inflammatory properties and inhibit glial activation-mediated oxidative stress, suggesting that paroxetine and its analogues may have therapeutic value in the treatment of aspects of Parkinson’s disease related to neuroinflammation.

Parkinson’s disease (PD) is a common neurodegenerative disease characterized by abnormal motor behavior, characterized by resting tremor, rigidity, and bradykinesia (1, 2). The neuropathological features of PD are progressive loss of dopaminergic (DA) neurons in the substantia nigra (SN) and depletion of dopamine in the striatum (STR), the site to which these neuronal nerve terminals project (3). Although PD is a sporadic disease of unknown pathogenesis, accumulating evidence suggests that glial activation-derived oxidative stress increases the risk of developing PD (4). In vivo and in vitro 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) models of PD show that key enzymes involved in the production of reactive oxygen species (ROS)/reactive nitrogen species (RNS), such as microglial NAPDH oxidase and inducible NO synthase (iNOS), and astroglial myeloperoxidase (MPO) are upregulated in damaged areas and contribute to DA neuronal death (58). Additionally, proinflammatory cytokines, such as TNF-α and IL-1β, are also increased and contribute to DA neuronal death in the MPTP model of PD (9).

Paroxetine, a common selective serotonin reuptake inhibitor, is widely prescribed as an antidepressant because it has fewer side effects and lower toxicity than earlier-generation selective serotonin reuptake inhibitors (10). The most frequent neuropsychiatric symptom in PD is depression (11), and several clinical reports have shown that paroxetine is safe and effective in treating PD-associated depression (12, 13). Antidepressants such as fluoxetine and imipramine have been shown to possess potent anti-inflammatory effects in the rat cerebral ischemia model of middle cerebral artery occlusion (14) and in a mouse model of Alzheimer’s disease (15). These studies demonstrated that fluoxetine and/or imipramine attenuated expression of TNF-α and IL-1β. Similarly, paroxetine was found to decrease the release of TNF-α in rat splenocytes (16) and inhibit infiltration of peripheral immune cells and production of the proinflammatory molecule substance P in an animal model of atopic dermatitis (17). However, little is known about the effects of paroxetine in the CNS, especially in the nigrostriatal DA system in the context of PD. Thus, the current study sought to determine whether paroxetine could prevent the degeneration of nigrostriatal DA neurons by inhibiting glial activation and, ultimately, reducing oxidative stress in the MPTP model of PD.

All of the experiments were done in accordance with approved animal protocol and guidelines established by Kyung Hee University. All of the experiments were conducted with 8- to 10-wk-old male C57BL/6 mice (22–24 g; Charles River Breeding Laboratory, Yokohama, Japan) in a room maintained at 20–22°C on a 12 h light/dark cycle with food and water available ad libitum. For MPTP intoxication, mice received four i.p. injections of MPTP-HCl (20 mg/kg; Sigma-Aldrich, St. Louis, MO) dissolved in PBS at 2 h intervals as previously described (18). For paroxetine treatment, mice received a single injection per day of paroxetine (10 mg/kg body weight, equivalent to 0.2–0.25 mg/day) into the peritoneum for 6 d, beginning at 12 h after last MPTP injection. Some mice were injected with paroxetine alone or vehicle as a control.

Animals were transcardially perfused with a saline solution containing 0.5% sodium nitrate and heparin (10 U/ml) and then fixed with 4% paraformaldehyde dissolved in 0.1 M phosphate buffer. Brains were dissected from the skull, postfixed overnight in buffered 4% paraformaldehyde at 4°C, stored in a 30% sucrose solution for 24–48 h at 4°C until they sank, frozen, and sectioned on a sliding microtome in 30-μm-thick coronal sections. All of the sections were collected in six separate series and processed for immunohistochemical staining as described previously (19). The primary Abs included those directed against tyrosine hydroxylase (TH; 1:2000; Pel-Freez, Brown Deer, WI), macrophage Ag complex-1 (MAC-1; 1:200; Serotec, Oxford, U.K.), ionized calcium-binding adaptor molecule 1 (Iba-1; 1:1000; Wako Chemicals, Osaka, Japan), CD68 (ED-1; 1:1000; Serotec), glial fibrillary acidic protein (GFAP; 1:5000; Neuromics, Edina, MN), MPO (1:500; Thermo Scientific, Fremont, CA), 8-hydroxy-2′-deoxyguanosine (8-OHdG; 1:200; JaICA, Shizuoka, Japan), iNOS (1:200, BD Biosciences, San Jose, CA), IL-1β (1:200, R&D Systems, Minneapolis, MN), and p47phox (1:200; Santa Cruz Biotechnology, Santa Cruz, CA). Stained cells were viewed and analyzed under a bright-field microscope (Nikon, Tokyo, Japan) or viewed with a confocal laser-scanning microscope (Olympus Optical, Tokyo, Japan). For Nissl staining, SN tissues immunostained with TH Ab were mounted on gelatin-coated slide, dried for 1 h at room temperature, and stained with 0.5% cresyl violet (Sigma-Aldrich). To validate immunohistochemical data, we performed immunohistochemistry with each isotype-matched control Ab (Supplemental Fig. 3, Supplemental Table I).

The unbiased stereological estimation of the total number of the TH-positive cells and Nissl-stained neurons in the SN was made using the optical fractionator method performed on an Olympus Computer Assisted Stereological Toolbox system, version 2.1.4 (Olympus, Ballerup, Denmark), as we described in detail (19, 20). The sections used for counting covered the entire SN from the rostral tip of the pars compacta back to the caudal end of the pars reticulate (anterioposterior, −2.06 to −4.16 mm from bregma) (21). The SN was delineated at a ×1.25 objective and generated a counting grid of 150 × 150 μm. An unbiased counting frame of known area (47.87 × 36.19 μm = 1733 μm2) superimposed on the image was placed randomly on the first counting area and systemically moved through all of the counting areas until the entire delineated area was sampled. Actual counting was performed using a ×100 oil immersion objective. The estimate of the total number of neurons was calculated according to the optical fractionator equation (22). More than 300 points over all of the sections of each specimen were analyzed.

As previously described (23), an average of 17 coronal sections of the striatum, starting from the rostral anterioposterior (+1.60 mm) to anterioposterior (0.00 mm), according to bregma of the brain atlas (21), were examined at a ×5 magnification using the Image-Pro Plus system (version 4.0; Media Cybernetics, Silver Spring, MD) on a computer attached to a light microscope (Zeiss Axioskop, Oberkochen, Germany), interfaced with a charge-coupled device video camera (MegaPlus model 1.4i; Kodak, New York, NY). To determine the density of the TH-immunoreactive staining in the striatum, a square frame of 700 × 700 μm was placed in the dorsal part of the striatum. A second square frame of 200 × 200 μm was placed in the region of the corpus callosum to measure background values. To control for variations in background illumination, the average of the background density readings from the corpus callosum was subtracted from the average of density readings of the striatum for each section. Then the average of all of the sections of each animal was calculated separately before data were statistically processed.

An accelerating rotarod (five-lane accelerating rotarod; Ugo Basile, Comerio, Italy) was used to measure forelimb and hindlimb motor dysfunction (9), with some modifications. The rotarod unit consisted of a rotating spindle (diameter, 3 cm) where mice were challenged for speed. For training, mice were placed on the rotarod at 10 rpm for 20 min, seven consecutive days before MPTP treatment. Mice that stayed on the rod without falling during training were selected and randomly divided into experimental groups. After 7 d from last MPTP treatment, animals receiving various treatment regimes were placed in a separate compartment on the rotating rod and tested at 20 rpm for 20 min. The latency to fall was automatically recorded by magnetic trip plates.

To measure contents of dopamine in STR, we performed reverse-phase HPLC with electrochemical detector as previously described (24). Dissected striatal tissues were homogenized with 0.1 M perchloric acid and 0.1 mM EDTA buffer and centrifuged at 9000 rpm for 20 min. The supernatant was injected into an autosampler at 4°C (Waters 717 Plus Autosampler, Waters Division, Milford, MA) and eluted through μBondapak C18 column (3.9 × 300 mm ×10 μm; ESA, Chelmsford, MA) with mobile phase for catecholamine analysis (Chromosystems, Munich, Germany). The peaks of dopamine content were analyzed by ESA Coulochem II electrochemical detector and integrated using a commercially available program (Breeze; Waters). All of the samples were normalized for protein content as spectrophotometrically determined using the Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA).

Hydroethidine histochemistry was performed for in situ visualization of O2 and O2-derived oxidants (7). Three days after MPTP injection, hydroethidine (1 mg/ml in PBS containing 1% DMSO; Molecular Probes, Eugene, OR) was administered i.p. After 15 min, the animals were transcardially perfused with a saline solution containing 0.5% sodium nitrate and heparin (10 U/ml) and then fixed with 4% paraformaldehyde in 0.1 M phosphate buffer. After fixation, the brains were cut into 30-μm slices using a sliding microtome. Sections were mounted on gelatin-coated slides, and the oxidized hydroethidine product, ethidium, was examined by IX71 confocal microscopy (Olympus Optical, Tokyo, Japan). To determine specific cell type related to oxidant production, we performed double-fluorescence staining with HE and MAC-1 or HE and TH Ab (Supplemental Fig. 2).

Animals treated with or without paroxetine (10 mg/kg, i.p.) were humanely killed 24 h after injection of MPTP, and the bilateral SN regions were immediately isolated. In brief, tissues were homogenized and total RNA was extracting with RNAzol B (Tel-Test, Friendwood, TX). Reverse transcription was performed with SuperScript II Reverse Transcriptase (Life Technologies, Rockville, MD). The primer sequences used in this study were as follows: 5′-CTGCTGGTGGTGACAAGCACATTT-3′ (forward) and 5′- ATGTCATGAGCAAAGGCGCAGAAC-3′ (reverse) for iNOS; 5′-GCGACGTGGAACTGGCAGAAGAG-3′ (forward) and 5′-TGAGAGGGAGGCCATTTGGGAAC-3′ (reverse) for TNF-α; 5′-GCAACTGTTCCTGAACTCAACT-3′ (forward) and 5′-ATCTTTTGGGGTCCGTCAACT-3′ (reverse) for IL-1β; and 5′-TCAACAGCAACTCCCACTCTTCCA-3′ (forward) and 5′-ACCCTGTTGCTGTAGCCGTATTCA-3′ (reverse) for G3PDH. Real-time PCR was performed in a reaction volume of 10 μl including 2 μl 1:50 diluted reverse transcription product as a template, 5 μl SYBR Premix EX Taq (Takara, Shiga, Japan) and 10 pmol of each primer described above. The PCR amplifications were performed with 50 cycles of denaturation at 95°C for 5 s, annealing at 60°C for 10 s, and extension at 72°C for 20 s using LightCycler (Roche Applied Science, Indianapolis, IN). The ΔCT value was determined by subtracting average CT values of G3PDH from average CT values of IL-1β, TNF-α, and iNOS from PCR reactions (Supplemental Table II). To express the relative amounts of IL-1β, TNF-α, and iNOS, ΔΔCT values were calculated by subtracting the ΔCT value of the control group from the ΔCT values of each group. The ratios of expression levels of IL-1β, TNF-α, and iNOS were calculated as 2−(meanΔΔCt).

Animals treated with or without paroxetine (10 mg/kg, i.p.) were humanely killed 72 h after injection of MPTP, and the bilateral SN regions were immediately isolated. The production of TNF-α and IL-1β from mice SN tissues was determined by sandwich ELISA techniques. Tissues were homogenized in 200 μl ice-cold radioimmunoprecipitation assay buffer (60 mM NaCl, 0.1% SDS, 1% Nonidet P-40, 0.5% sodium deoxycholic acid, and 50 mM Tris [pH 8]) and centrifuged at 14,000 × g at 4°C for 20 min. Equal amounts of samples (100 μg) were placed in ELISA kit strips coated with the appropriate Ab. Sandwich ELISA was then performed according to the manufacturer’s instructions (BioSource International, Camarillo, CA). The detection limits of TNF-α and IL-1β were 5 and 25 pg/ml, respectively. We diluted TNF-α samples and IL-1β samples as 1:10, respectively.

For separating the cytosolic and membrane fraction, we dissected SN tissues from the animals at 3 d after injection of MPTP. As previously described (25, 26), SN samples were homogenized with using a glass homogenizer in ice-cold buffer consisting of the following: 20 mM HEPES, 250 mM sucrose, 10 mM KCl, 1.5 mM MgCl2, 2 mM EDTA, and protease inhibitor mixture (Sigma-Aldrich). Homogenates were centrifuged for 5 min at 500 × g at 4°C, and supernatants were collected and centrifuged for 20 min at 13,000 × g at 4°C. The pellets were further centrifuged for 1 h at 100,000 × g at 4°C, and the resulting supernatants and pellets were designated as the cytosolic and membrane fractions, respectively. Equal amounts of proteins (30 μg) were separated by 12% SDS-PAGE gels and transferred to polyvinylidene difluoride membranes (Millipore, Bedford, MA) using an electrophoretic transfer system (Bio-Rad). The membranes were immunoblotted with goat anti-actin (1:1000; Santa Cruz Biotechnology), rabbit anti-p47phox (1:1000; BD Biosciences), and mouse anti-Rac1 (1:500; Santa Cruz Biotechnology). To determine the relative degree of membrane purification, the membrane fraction was subjected to immunoblotting for calnexin, a membrane marker, using a rabbit anti-calnexin (1:1000; Stressgen, Victoria, BC, Canada). For semiquantitative analyses, the densities of bands on immunoblots were measured with a computer imaging device and accompanying software (Fujifilm, Tokyo, Japan).

All of the values are expressed as mean ± SEM. Statistical significance (p < 0.05 for all analyses) was assessed by ANOVA using Instat 3.05 (GraphPad, San Diego, CA), followed by Student–Newman–Keuls analyses.

In the brain, MPTP is converted to 1-methyl-4-phenyl-pyridinium (MPP+), an active toxic metabolite that is primarily responsible for MPTP neurotoxicity (27). Consistent with this, striatal MPP+ content correlates linearly with MPTP toxicity (28). Striatal levels of MPTP and MPP+ at specific time points after the last MPTP injection were measured and quantified by liquid chromatography with electrospray ionization mass spectrometry. MPP+ levels peaked within 30 min at 14.9 ± 2.4 μg/mg protein and then gradually declined, becoming almost negligible (0.2 ± 0.1 μg/mg protein) after 12 h. These data indicate that MPTP is completely converted to MPP+ and largely cleared within 12 h after injection. Because paroxetine was administered 12 h after the final injection of MPTP for all in vivo experiments, its effects could not be attributed to reduced metabolism of MPTP to MPP+ or uptake of MPP+ into DA neurons. At this time point, the extent of damage of the nigrostriatal DA system was assessed by TH immunostaining and the results are shown in Supplemental Fig. 1.

The mice in each group received four i.p. injections of MPTP (20 mg/kg) or PBS as a control at 2 h intervals. Seven days later, brains were removed and sections were immunostained for TH to specifically detect DA neurons, and then SN tissues were processed for Nissl staining. Similar to a recent report (18), there was a considerable loss of TH-immunopositive (ip) cell bodies in the SN (Fig. 1E, 1F) and fibers in the STR (Fig. 2C) at 7 d in the MPTP-injected mice compared with those of the PBS-treated control mice (Figs. 1A, 1B, 2A). TH-ip cells in the SN and their nerve terminals in the STR were quantified by stereological counts and densitometric analyses, respectively. MPTP treatment decreased the number of TH-ip neurons by 63% in the SN (Fig. 1I) and reduced the OD of TH-ip fibers by 62% in the STR (Fig. 2E) compared with those of the PBS-treated mice. Moreover, PBS-treated SN had prominent Nissl substances (Fig. 1B) when compared with MPTP-treated SN, showing marked loss of Nissl substances with gliosis (Fig. 1F). The stereological counts of these substances revealed that MPTP decreased the number of Nissl-positive neurons by 61% in the SN compared with those of the PBS-treated SN (Fig. 1I).

FIGURE 1.

Paroxetineprevents MPTP-induced neurotoxicity in the SN of mouse brains. Animals receiving PBS as a control (A, B), paroxetine alone (C, D), MPTP (E, F), or MPTP and paroxetine (G, H) were sacrificed 7 d after last MPTP administration. The brain tissues were cut using a sliding microtome in 30-μm-thick coronal sections and immunostained with Ab to TH (brown) for DA neurons, and then Nissl staining (blue) was performed. Note the absence of TH expression in some Nissl-stained neurons, indicated by arrows. B, D, F, and H show higher magnifications of A, C, E, and G, respectively. I, Numbers of TH-ip and Nissl-stained neurons in the SN were counted. Four to 10 animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.01, significantly different from MPTP only. AD, Scale bars, 200 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine; SNpc, substantia nigra pars compacta; SNr, substantia nigra reticulata; VTA, ventral tegmental area.

FIGURE 1.

Paroxetineprevents MPTP-induced neurotoxicity in the SN of mouse brains. Animals receiving PBS as a control (A, B), paroxetine alone (C, D), MPTP (E, F), or MPTP and paroxetine (G, H) were sacrificed 7 d after last MPTP administration. The brain tissues were cut using a sliding microtome in 30-μm-thick coronal sections and immunostained with Ab to TH (brown) for DA neurons, and then Nissl staining (blue) was performed. Note the absence of TH expression in some Nissl-stained neurons, indicated by arrows. B, D, F, and H show higher magnifications of A, C, E, and G, respectively. I, Numbers of TH-ip and Nissl-stained neurons in the SN were counted. Four to 10 animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.01, significantly different from MPTP only. AD, Scale bars, 200 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine; SNpc, substantia nigra pars compacta; SNr, substantia nigra reticulata; VTA, ventral tegmental area.

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FIGURE 2.

Paroxetine protects neurotoxicity induced by MPTP in the STR in vivo. The STR tissues obtained from the same animals as used in Fig. 1 were immunostained with TH Ab for dopaminergic fibers: PBS (A), paroxetine only (B), MPTP (C), or MPTP and paroxetine (D). E, OD of TH-ip fiber in the STR. Four to six animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.001, significantly different from MPTP only. AD, Scale bars, 250 μm. F, After the rotarod test, striatal tissues were dissected from animals to determine the dopamine level in the striatum. Five to six animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.05, significantly different from MPTP only. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

FIGURE 2.

Paroxetine protects neurotoxicity induced by MPTP in the STR in vivo. The STR tissues obtained from the same animals as used in Fig. 1 were immunostained with TH Ab for dopaminergic fibers: PBS (A), paroxetine only (B), MPTP (C), or MPTP and paroxetine (D). E, OD of TH-ip fiber in the STR. Four to six animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.001, significantly different from MPTP only. AD, Scale bars, 250 μm. F, After the rotarod test, striatal tissues were dissected from animals to determine the dopamine level in the striatum. Five to six animals were used for each experimental group. *p < 0.001, significantly different from controls; **p < 0.05, significantly different from MPTP only. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

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To investigate whether paroxetine altered MPTP-induced neurotoxicity of nigrostriatal DA neurons, we administered paroxetine (10 mg/kg body weight) for 6 d, starting 12 h after the last MPTP injection. The results of TH immunohistochemistry showed that paroxetine treatment effectively reduced MPTP-induced loss of DA neurons in the SN (Fig. 1G, 1H) and their nerve terminals in the STR (Fig. 2D). When quantified and expressed as a percentage of TH-ip neurons or fibers in the control SN and STR, respectively, paroxetine was found to increase the number of TH-ip neurons by 26% (Fig. 1I; p < 0.01) and the OD of TH-ip fibers by 21% (Fig. 2E; p < 0.001). Consistent with these results, paroxetine was found to increase the number of Nissl-stained neurons by 25% in the SN (Fig. 1I; p < 0.01). Paroxetine alone had no effects on the number of neurons in the SN (Fig. 1C, 1D, 1I) or DA fibers in the STR (Fig. 2B, 2E).

We next determined whether paroxetine improves MPTP-induced motor deficits by testing performance on a rotarod apparatus with some modifications (9). Animals receiving various treatment regimes were evaluated 7 d after the last MPTP injection by measuring the latency to falling. MPTP treatment decreased sustained rotarod time to 11.29 ± 0.49 min, a 56% decrease compared with that of PBS treatment (Fig. 2F; p < 0.001). The behavioral dysfunction in MPTP-treated mice was partially recovered by paroxetine treatment, which increased the latency to falling to 14.07 ± 0.35 min (Fig. 2F; p < 0.05). Paroxetine alone had no effects on motor behavior.

After rotarod performance tests, the mice were sacrificed for biochemical assessments. In parallel with behavioral dysfunction, HPLC analysis revealed that MPTP reduced STR dopamine levels to 75% of those in PBS controls (Fig. 2F). By contrast, treatment of MPTP-injected mice with paroxetine increased dopamine levels by 22% compared with MPTP only (Fig. 2F; p < 0.05). Paroxetine alone had no effect on STR dopamine levels.

Several lines of evidence suggest that activated microglia play a critical role in DA neuronal cell death in the MPTP model (29). Thus, we next examined whether the neuroprotective effect of paroxetine resulted from inhibition of microglial activation in the SN. At 3 d after the last MPTP injection, brain tissues from mice treated with or without paroxetine were processed for immunostaining with MAC-1, Iba-1, and ED-1 Abs to detect activated microglia. In contrast with PBS controls, where relatively few positive microglia were observed in the SN (Fig. 3A), there were numerous robustly immunoreactive MAC-1–positive (activated) microglia in the MPTP-treated SN (Fig. 3B), consistent with a previous report (30). Paroxetine treatment dramatically reduced the number of MAC-1-ip microglia in the MPTP-treated SN (Fig. 3C), indicating that paroxetine suppressed MPTP-induced microglial activation.

FIGURE 3.

Paroxetine prevents microglial activation in the SN in vivo. Animals receiving PBS as a control (A, D, G), MPTP (B, E, H), or MPTP and paroxetine (C, F, I) were sacrificed 3 d after the last MPTP injection. The mouse brain tissues were removed and immunostained with Abs to MAC-1 (AC), Iba-1 (DF), and ED-1 (GI) to identify microglia. Insets show higher magnifications of AI, respectively. Dotted lines indicate SNpc. AI, Scale bars, 100 μm.

FIGURE 3.

Paroxetine prevents microglial activation in the SN in vivo. Animals receiving PBS as a control (A, D, G), MPTP (B, E, H), or MPTP and paroxetine (C, F, I) were sacrificed 3 d after the last MPTP injection. The mouse brain tissues were removed and immunostained with Abs to MAC-1 (AC), Iba-1 (DF), and ED-1 (GI) to identify microglia. Insets show higher magnifications of AI, respectively. Dotted lines indicate SNpc. AI, Scale bars, 100 μm.

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Consistent with MAC-1 staining, Iba-1–positive microglia exhibited the typical ramified morphology of resting microglia in the PBS-treated control SN (Fig. 3D). In contrast, the majority of Iba-1–positive microglia displayed an activated morphology, including larger cell bodies with short, thick, or no processes in MPTP-treated SN (Fig. 3E). Treatment with paroxetine mitigated these effects of MPTP, dramatically decreasing the number of activated microglia in the MPTP-treated SN (Fig. 3F).

Microglia in the MPTP-treated SN appeared to reach a state similar to that of active phagocytes (Fig. 3H), as determined by ED-1 immunohistochemical staining, which labels phagocytic microglia, in particular, the presence of injured cells or debris (31). Few such ED-1-ip cells were observed in the paroxetine-treated SN (Fig. 3I), similar to the results of MAC-1 immunostaining. ED-1-ip cells were not detectable in the PBS-treated SN (Fig. 3G). Paroxetine alone had no effect on microglial activation (data not shown).

It has been shown that mice expressing a dominant-negative inhibitor of IL-1β–converting enzyme or those deficient in TNF-α or iNOS are resistant to MPTP-induced neurotoxicity (6, 32, 33). Thus, we examined whether paroxetine might modulate DA neuronal survival by affecting MPTP-induced expression of IL-1β, TNF-α, and/or iNOS in the SN. For this purpose, paroxetine was administered 12 h after the last MPTP injection and animals were sacrificed 12 h later. An analysis of dissected brain tissues by real-time PCR revealed that paroxetine administered after MPTP significantly inhibited MPTP-induced expression of proinflammatory cytokines, reducing IL-1β, TNF-α, and iNOS expression by 57, 60, and 76%, respectively (Fig. 4A), but had no effect when administered alone. Consistent with real-time PCR analyses, ELISA results showed that the levels of IL-1β and TNF-α protein were significantly increased at 3 d in the MPTP-treated SN compared with those of the PBS-treated SN (Fig. 4B). These MPTP-induced increases were significantly attenuated by treatment with paroxetine, which had no effect alone. To identify the cell types expressing IL-1β and iNOS protein, we performed double-immunofluorescent staining in SN sections obtained 3 d after MPTP injection using a combination of Abs against MAC-1 and IL-1β or iNOS. Simultaneous imaging of immunofluorescence in the same tissue sections revealed that IL-1β (Fig. 4C) and iNOS (Fig. 4D) immunoreactivity was localized within MAC-1-ip microglia.

FIGURE 4.

Paroxetine reduces the MPTP-induced inflammation in the SN of mouse brains. Animals were administered MPTP or vehicle in the presence or absence of paroxetine and sacrificed 1 d later for real-time PCR (A) or 3 d later for a sandwich ELISA in the SN (B). Paroxetine dramatically attenuated MPTP-induced expression of proinflammatory cytokines including IL-1β, TNF-α, and iNOS. Each bar represents the mean ± SEM of four to five animals per group. *p < 0.05, significantly different from control; **p < 0.05, significantly different from MPTP. Colocalization of IL-1β (E) or iNOS (I) immunoreactivity within activated microglia in the SN in vivo. The sections of mouse SN were prepared 3 d after MPTP injection and then immunostained simultaneously with MAC-1 (C, G; green) as a marker of microglia and IL-1β (D; red) or iNOS (H; red). To verify the colocalized images, double-immunofluorescence staining was performed with each isotype-matched control (F, J). Images from the same double-labeled tissue were merged (E, I). CJ, Scale bars, 50 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

FIGURE 4.

Paroxetine reduces the MPTP-induced inflammation in the SN of mouse brains. Animals were administered MPTP or vehicle in the presence or absence of paroxetine and sacrificed 1 d later for real-time PCR (A) or 3 d later for a sandwich ELISA in the SN (B). Paroxetine dramatically attenuated MPTP-induced expression of proinflammatory cytokines including IL-1β, TNF-α, and iNOS. Each bar represents the mean ± SEM of four to five animals per group. *p < 0.05, significantly different from control; **p < 0.05, significantly different from MPTP. Colocalization of IL-1β (E) or iNOS (I) immunoreactivity within activated microglia in the SN in vivo. The sections of mouse SN were prepared 3 d after MPTP injection and then immunostained simultaneously with MAC-1 (C, G; green) as a marker of microglia and IL-1β (D; red) or iNOS (H; red). To verify the colocalized images, double-immunofluorescence staining was performed with each isotype-matched control (F, J). Images from the same double-labeled tissue were merged (E, I). CJ, Scale bars, 50 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

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The production of ROS, such as hydrogen peroxide and superoxide, are increased in the midbrain of MPTP-treated mice, and oxidant originating from microglia is thought to mediate the loss of DA neurons in the SN (34). Thus, we examined whether paroxetine rescued nigral DA neurons by inhibiting MPTP-induced oxidant production. The fluorescent product of oxidized hydroethidine (i.e., ethidium) was significantly increased in the MPTP-injected SN 72 h after administration (Fig. 5B) compared with that of the PBS-injected SN (Fig. 5A). Paroxetine dramatically decreased MPTP-induced oxidant production in the SN in vivo (Fig. 5C) but had no effect alone (data not shown).

FIGURE 5.

Paroxetine inhibits MPTP-induced oxidant production, activation of NADPH oxidase, and oxidative DNA damage in the SN in vivo. AC, Animals receiving PBS as a control (A), MPTP (B), or MPTP and paroxetine (C) were sacrificed 3 d after the last MPTP injection. SN tissues were prepared for hydroethidine histochemistry to detect oxidant production. Dotted lines indicate SNpc. D, Translocation of cytosolic subunits (p47phox and Rac1) from the cytosol to the plasma membrane after MPTP injection, indicating activation of NADPH oxidase in the SN. Animals were decapitated after injection of MPTP, and SN tissues were isolated. Tissue lysates were fractionated and analyzed by immunoblot analysis with p47phox or Rac1 Ab. The membrane protein calnexin was used to normalize the data. E, The histogram shows quantification of p47phox and Rac1 levels expressed as the ratio of membrane fraction to total. The results represent the mean ± SEM of three to four separate experiments. *p < 0.01, compared with control; ** p < 0.05, compared with MPTP only. F, Localization of p47phox immunoreactivity in activated microglia in the SN treated with MPTP. The sections of SN were prepared 3 d after MPTP injection and then simultaneously stained with an Ab against p47phox and MAC-1 as a marker for microglia. GI, The SN tissues obtained from the same animals as used in AC were prepared for 8-OHdG histochemistry to detect oxidative DNA damage in the SN. Dotted lines indicate SNpc. Insets show higher magnifications of GI, respectively. Scale bars: AC, 150 μm; F, 50 μm; GI, 100 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

FIGURE 5.

Paroxetine inhibits MPTP-induced oxidant production, activation of NADPH oxidase, and oxidative DNA damage in the SN in vivo. AC, Animals receiving PBS as a control (A), MPTP (B), or MPTP and paroxetine (C) were sacrificed 3 d after the last MPTP injection. SN tissues were prepared for hydroethidine histochemistry to detect oxidant production. Dotted lines indicate SNpc. D, Translocation of cytosolic subunits (p47phox and Rac1) from the cytosol to the plasma membrane after MPTP injection, indicating activation of NADPH oxidase in the SN. Animals were decapitated after injection of MPTP, and SN tissues were isolated. Tissue lysates were fractionated and analyzed by immunoblot analysis with p47phox or Rac1 Ab. The membrane protein calnexin was used to normalize the data. E, The histogram shows quantification of p47phox and Rac1 levels expressed as the ratio of membrane fraction to total. The results represent the mean ± SEM of three to four separate experiments. *p < 0.01, compared with control; ** p < 0.05, compared with MPTP only. F, Localization of p47phox immunoreactivity in activated microglia in the SN treated with MPTP. The sections of SN were prepared 3 d after MPTP injection and then simultaneously stained with an Ab against p47phox and MAC-1 as a marker for microglia. GI, The SN tissues obtained from the same animals as used in AC were prepared for 8-OHdG histochemistry to detect oxidative DNA damage in the SN. Dotted lines indicate SNpc. Insets show higher magnifications of GI, respectively. Scale bars: AC, 150 μm; F, 50 μm; GI, 100 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

Close modal

NADPH oxidase is composed of the cytosolic components p47phox, p67phox, and Rac1 and the membrane components gp91phox and p22phox (35, 36). Translocation of NADPH oxidase subunits from the cytosol to the plasma membrane in activated microglia produces ROS, which ultimately contribute to DA neuronal death in the MPTP mouse model of PD (7). To investigate whether paroxetine modulates the activity of NADPH oxidase, we performed Western blotting after separating cell lysates into membrane and cytosolic components. Western blot analyses showed that the levels of p47phox and Rac1 in the MPTP-treated SN were significantly increased in membrane fractions 3 d after MPTP injection compared with those of the PBS-treated SN controls (Fig. 5D, 5E), indicating increased translocation of these subunits. Administration of paroxetine after MPTP injection significantly decreased translocation of p47phox and Rac1 from the cytosol to the membrane in the SN (Fig. 5D, 5E). Paroxetine alone had no effects on the level of p47phox or Rac1 in either cytosolic or membrane fractions. These results confirm that MPTP-induced activation of NADPH oxidase is inhibited by paroxetine. Moreover, the p47phox-ip cells present in the SN 3 d after MPTP injection corresponded to activated microglia (Fig. 5F) but not astrocytes or neurons (data not shown).

In MPTP mice, DA neuronal cell death is accompanied by increased levels of 8-OHdG, a marker of oxidative nucleic acid damage (37). To determine whether paroxetine prevented MPTP-induced oxidative damage to DNA in the SN, we immunostained for 8-OHdG. Our results revealed that the levels of 8-OHdG were dramatically increased in the SN 3 d after MPTP injection (Fig. 5H) compared with those of the PBS-treated SN (Fig. 5G). This MPTP-induced oxidative DNA damage was dramatically inhibited by paroxetine (Fig. 5I), which had no effect alone (data not shown).

It has been shown that MPO is upregulated in astrocytes in the SN of PD patients and MPTP-treated mice; furthermore, mice deficient in MPO are resistant to MPTP neurotoxicity (5). Thus, we next examined whether paroxetine inhibited MPTP-induced astroglial activation and expression of MPO in the SN, which would be predicted to enhance neuronal survival. To ascertain this, we immunostained sections adjacent to those used for MAC-1 immunostaining for GFAP and MPO 3 d after MPTP injection. In agreement with previous studies (6, 30), resting astrocytes (Fig. 6A; small somas and thin dendrites) were transformed into cells with enlarged bodies and thick dendrites in MPTP-treated mice (Fig. 6B). Treatment with paroxetine (10 mg/kg, i.p.) after MPTP injection dramatically decreased the number of activated astrocytes in the MPTP-treated mice (Fig. 6C). There was also an increase in immunostaining for MPO-ip cells in the MPTP-treated SN (Fig. 6E) compared with that of the control SN (Fig. 6D), an increase that was significantly attenuated by paroxetine (Fig. 6F). Quantification of MPO-ip cells in the SN by stereological cell counts confirmed these results, showing that the number MPO-ip cells was 263-fold higher in the MPTP-treated SN than that in the SN of PBS-treated controls (Fig. 6G). Paroxetine reduced the number of MPTP-induced MPO-ip cells by 75% (Fig. 6G). As controls, vehicle (PBS) and paroxetine alone had no effects. Additional immunofluorescence staining revealed that MPO immunoreactivity was localized within astrocytes 3 d after MPTP treatment (Fig. 6H).

FIGURE 6.

Paroxetine suppresses MPTP-induced astroglial activation and expression of MPO in the SN in vivo. The SN tissues obtained from the same animals as used in Fig. 5 were immunostained with GFAP Ab for astrocyte (AC) and MPO Ab for MPO immunoreactivity (DF). Animals receiving PBS as a control (A, D), MPTP (B, E), or MPTP and paroxetine (C, F) were sacrificed 3 d after the last MPTP injection. Insets show higher magnifications of AC, respectively. Dotted lines indicate SNpc. G, Number of MPO-ip cells in the SN were counted. Four to five animals were used for each experimental group. *p < 0.01, significantly different from controls; **p < 0.05, significantly different from MPTP only. H, Localization of MPO immunoreactivity in activated astrocytes in the SN treated with MPTP. The SN tissues obtained from the same animals as used in B were simultaneously stained with an Ab against MPO and GFAP as a maker for astrocyte. Scale bars: AF, 200 μm; G, 50 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

FIGURE 6.

Paroxetine suppresses MPTP-induced astroglial activation and expression of MPO in the SN in vivo. The SN tissues obtained from the same animals as used in Fig. 5 were immunostained with GFAP Ab for astrocyte (AC) and MPO Ab for MPO immunoreactivity (DF). Animals receiving PBS as a control (A, D), MPTP (B, E), or MPTP and paroxetine (C, F) were sacrificed 3 d after the last MPTP injection. Insets show higher magnifications of AC, respectively. Dotted lines indicate SNpc. G, Number of MPO-ip cells in the SN were counted. Four to five animals were used for each experimental group. *p < 0.01, significantly different from controls; **p < 0.05, significantly different from MPTP only. H, Localization of MPO immunoreactivity in activated astrocytes in the SN treated with MPTP. The SN tissues obtained from the same animals as used in B were simultaneously stained with an Ab against MPO and GFAP as a maker for astrocyte. Scale bars: AF, 200 μm; G, 50 μm. C, control; M, MPTP; MP, MPTP and paroxetine; P, paroxetine.

Close modal

In this study, we demonstrated that paroxetine protects nigrostriatal DA neurons from MPTP neurotoxicity in vivo by inhibiting brain inflammation and the resultant oxidative stress. We showed that paroxetine suppressed MPTP-induced ROS generation and reduced oxidative damage to nucleic acids by inhibiting microglia-derived NADPH oxidase and astrocyte-derived MPO, leading to survival of nigrostriatal DA neurons, recovery of striatal dopamine depletion in vivo, and reversal of motor dysfunction. Additionally, paroxetine attenuated the expression of proinflammatory cytokines and iNOS within activated microglia. To our knowledge, this is the first study to show that paroxetine prevents nigrostriatal DA neuronal death through blockade of glial activation in the MPTP model of PD.

Glial cells play an important role in supporting neurons in the CNS. However, in the presence of adverse stimuli, they may contribute to chronic, damaging inflammation and, ultimately, to neuronal cell death. Microglia are intrinsic immune effector cells that are dramatically activated in response to neuronal damage (38). Activated microglia produce several potentially neurotoxic substances, including ROS and/or proinflammatory cytokines (29).

ROS, such as O2 and O2-derived oxidant molecules, may impose an oxidative stress on DA neurons and induce and/or exacerbate neurotoxicity (39). Several studies have demonstrated evidence of oxidative stress in PD patients and in the MPTP model of PD, including oxidative modifications to nucleic acids (40, 41). Importantly, these ROS can be produced by microglia-derived NADPH oxidase and are capable of causing oxidative stress in the MPTP model of PD (29). Several studies, including ours, have demonstrated that activation of microglial NADPH oxidase participates in DA neuronal death in vivo and in vitro (7, 8, 25). The results of the current study show that MPTP activated NADPH oxidase, as demonstrated by the translocation of cytosolic components of NADPH oxidase, p47phox and Rac1. NADPH oxidase activation resulted in increased O2 and O2-derived oxidants and DNA damage, as visualized by hydroethidine staining and 8-OHdG immunostaining in the SN, respectively. Treatment with paroxetine not only inhibited microglial NADPH oxidase activation but also mitigated ROS production and nucleic acid oxidation. These results verify that paroxetine inhibited MPTP-induced activation of NADPH oxidase and oxidative damage, thereby resulting in neuroprotection in the MPTP model.

Accompanying oxidative stress, other microglial-derived proinflammatory cytokines or cytotoxic factors may be involved in nigrostriatal DA neuronal death. It has been shown that iNOS (42) and proinflammatory cytokines, such as TNF-α and IL-1β (43), are increased in the brains of PD patients. Moreover, NO, generated by iNOS, may also be involved in the pathogenesis of PD (27), although the presence of iNOS in human microglia remains controversial (44). The neurotoxic effects of NO are attributed to its reaction with O2 to form peroxynitrite, which can cause oxidative damage to proteins in the MPTP model (6). TNF-α and IL-1β, originating from activated glia, may trigger intracellular death-related signaling pathways or participate in the induction of iNOS expression in the MPTP model (45). The present data showed that MPTP increased the expression of iNOS, TNF-α, and IL-1β mRNA in the SN. Treatment with paroxetine inhibited the expression of these three molecules within activated microglia after MPTP injection, an effect that likely accounts, at least in part, for the observed paroxetine-induced neuroprotection. These results collectively suggest that paroxetine has anti-inflammatory properties that contribute to its neuroprotective effects. Similar to our results, a recent report showed that minocycline prevents MPTP neurotoxicity through relatively small changes in protein levels of IL-1β and TNF-α (46). Although genetic ablation of TNF-α decreased microglial activation and prevented disruption of blood–brain barriers, survival of DA neurons was not increased in the SN. Thus, biological relevance of these molecules may be varied depending on experimental conditions. Although we did not provide direct evidence, the present results combined with other observations carefully suggest that the biological relevance of these two molecules is due to each molecule and/or synergic effects combined with each other.

Although the role of astroglial activation in PD is disputed, several studies using the MPTP model support the idea that astroglial activation can contribute to the degeneration of nigrostriatal DA neurons via generation of neurotoxic substances (47, 48). One such substance expressed in activated astrocytes is MPO, which can use NO2 to generate RNS and cause DA neuronal death in the MPTP model (5). This increased expression of astroglial MPO mediates neurotoxicity on DA neurons under two possible mechanisms. First, MPO is known as the key enzyme for production of cytotoxic ROS/RNS (49). These astroglial MPO-derived oxidants can be released and then give deleterious effects on adjacent neurons (5). This was supported by evidence that increased levels of 3-chlorotyrosine– and HOCl-modified proteins and MPO-derived oxidative stress markers were observed in MPTP-treated SN (5). Another possible mechanism is the cross talk with other immune cells through MPO secretion (50). MPO can be secreted and then activate neutrophils through binding with CD11b/CD18 integrins. Because microglia also express CD11b/CD18 integrins, MPO secretion may participate in DA neuronal death in the MPTP model of PD via microglial activation. Although the cytokine-like effect of MPO is still unknown, this enzyme may play an important role in the signaling pathway of microglial activation, resulting in neuronal death. This is consistent with our present data showing that MPO was expressed in activated astrocytes in the MPTP-treated SN, as assessed by double-label immunostaining. Additional experiments showed that paroxetine attenuated MPO immunoreactivity in the SN. Taken together, our results suggest that paroxetine rescues DA neuronal death via suppression of MPO expression in activated astrocytes.

The most prominent biochemical change in the denervated STR caused by MPTP injection is a reduction in the levels of striatal dopamine (18). This biochemical deficit in mice yields a characteristic motor dysfunction that leads to increased latency to fall (deteriorated balance) on the accelerating rotarod (9). This notion is consistent with the present demonstration that motor performance on a rotarod is decreased with the loss of striatal TH terminals and, by extension, the consequent reduction of DA levels in the MPTP-treated STR (51). Paroxetine ameliorated these MPTP-induced motor deficits; accompanying this behavioral recovery was an increase in DA levels in the denervated STR. These behavioral and in vivo biochemical effects of paroxetine on the lesioned nigrostriatal DA system, taken together with our demonstration of inhibitory effects of paroxetine on glial activation-mediated oxidative stress, suggest that paroxetine and its analogues may have therapeutic value in the treatment PD symptoms related to neuroinflammation.

Finally, the most frequent neuropsychiatric symptom in PD is depression, which is observed in up to 46% of patients with PD (11). In this context, several clinical reports have concluded that paroxetine is a safe and effective drug for use in treating the depression associated with PD (12, 13). However, paroxetine has also been found to exacerbate parkinsonism symptoms associated with depression (52). Although our results suggest that paroxetine is beneficial in the treatment of aspects of PD related to glial activation, these conflicting clinical data suggest that the use of paroxetine in PD patients warrants caution (53).

We thank Dr. Noh and Dr. Yoon in Ajou University for providing paroxetine and valuable comments on this manuscript.

Disclosures The authors have no financial conflicts of interest.

This work was supported by the Basic Science Research Program through the National Research Foundation of Korea funded by the Ministry of Education, Science, and Technology (Grant 20090063274).

The online version of this article contains supplemental material.

Abbreviations used in this paper:

8-OHdG

8-hydroxy-2′-deoxyguanosine

C

control

DA

dopaminergic

ED-1

CD68

GFAP

glial fibrillary acidic protein

Iba-1

ionized calcium-binding adaptor molecule 1

iNOS

inducible NO synthase

ip

immunopositive

M

MPTP

MAC-1

macrophage Ag complex-1

MPO

myeloperoxidase

MP

MPTP and paroxetine

MPP+

1-methyl-4-phenyl-pyridinium

MPTP

1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine

P

paroxetine

PD

Parkinson’s disease

ROS

reactive oxygen species

RNS

reactive nitrogen species

SN

substantia nigra

SNpc

substantia nigra pars compacta

SNr

substantia nigra reticulata

STR

striatum

TH

tyrosine hydroxylase

VTA

ventral tegmental area.

1
Dauer
W.
,
Przedborski
S.
.
2003
.
Parkinson’s disease: mechanisms and models.
Neuron
39
:
889
909
.
2
Olanow
C. W.
,
Tatton
W. G.
.
1999
.
Etiology and pathogenesis of Parkinson’s disease.
Annu. Rev. Neurosci.
22
:
123
144
.
3
Savitt
J. M.
,
Dawson
V. L.
,
Dawson
T. M.
.
2006
.
Diagnosis and treatment of Parkinson disease: molecules to medicine.
J. Clin. Invest.
116
:
1744
1754
.
4
Hirsch
E. C.
,
Hunot
S.
.
2009
.
Neuroinflammation in Parkinson’s disease: a target for neuroprotection?
Lancet Neurol.
8
:
382
397
.
5
Choi
D. K.
,
Pennathur
S.
,
Perier
C.
,
Tieu
K.
,
Teismann
P.
,
Wu
D. C.
,
Jackson-Lewis
V.
,
Vila
M.
,
Vonsattel
J. P.
,
Heinecke
J. W.
,
Przedborski
S.
.
2005
.
Ablation of the inflammatory enzyme myeloperoxidase mitigates features of Parkinson’s disease in mice.
J. Neurosci.
25
:
6594
6600
.
6
Liberatore
G. T.
,
Jackson-Lewis
V.
,
Vukosavic
S.
,
Mandir
A. S.
,
Vila
M.
,
McAuliffe
W. G.
,
Dawson
V. L.
,
Dawson
T. M.
,
Przedborski
S.
.
1999
.
Inducible nitric oxide synthase stimulates dopaminergic neurodegeneration in the MPTP model of Parkinson disease.
Nat. Med.
5
:
1403
1409
.
7
Wu
D. C.
,
Teismann
P.
,
Tieu
K.
,
Vila
M.
,
Jackson-Lewis
V.
,
Ischiropoulos
H.
,
Przedborski
S.
.
2003
.
NADPH oxidase mediates oxidative stress in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine model of Parkinson’s disease.
Proc. Natl. Acad. Sci. USA
100
:
6145
6150
.
8
Gao
H. M.
,
Liu
B.
,
Zhang
W.
,
Hong
J. S.
.
2003
.
Critical role of microglial NADPH oxidase-derived free radicals in the in vitro MPTP model of Parkinson’s disease.
FASEB J.
17
:
1954
1956
.
9
Moon
M.
,
Kim
H. G.
,
Hwang
L.
,
Seo
J. H.
,
Kim
S.
,
Hwang
S.
,
Kim
S.
,
Lee
D.
,
Chung
H.
,
Oh
M. S.
, et al
.
2009
.
Neuroprotective effect of ghrelin in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine mouse model of Parkinson’s disease by blocking microglial activation.
Neurotox. Res.
15
:
332
347
.
10
Anderson
I. M.
2000
.
Selective serotonin reuptake inhibitors versus tricyclic antidepressants: a meta-analysis of efficacy and tolerability.
J. Affect. Disord.
58
:
19
36
.
11
Tom
T.
,
Cummings
J. L.
.
1998
.
Depression in Parkinson’s disease. Pharmacological characteristics and treatment.
Drugs Aging
12
:
55
74
.
12
Ceravolo
R.
,
Nuti
A.
,
Piccinni
A.
,
Dell’Agnello
G.
,
Bellini
G.
,
Gambaccini
G.
,
Dell’Osso
L.
,
Murri
L.
,
Bonuccelli
U.
.
2000
.
Paroxetine in Parkinson’s disease: effects on motor and depressive symptoms.
Neurology
55
:
1216
1218
.
13
Tesei
S.
,
Antonini
A.
,
Canesi
M.
,
Zecchinelli
A.
,
Mariani
C. B.
,
Pezzoli
G.
.
2000
.
Tolerability of paroxetine in Parkinson’s disease: a prospective study.
Mov. Disord.
15
:
986
989
.
14
Lim
C. M.
,
Kim
S. W.
,
Park
J. Y.
,
Kim
C.
,
Yoon
S. H.
,
Lee
J. K.
.
2009
.
Fluoxetine affords robust neuroprotection in the postischemic brain via its anti-inflammatory effect.
J. Neurosci. Res.
87
:
1037
1045
.
15
Chavant
F.
,
Deguil
J.
,
Pain
S.
,
Ingrand
I.
,
Milin
S.
,
Fauconneau
B.
,
Pérault-Pochat
M. C.
,
Lafay-Chebassier
C.
.
2010
.
Imipramine, in part through tumor necrosis factor alpha inhibition, prevents cognitive decline and beta-amyloid accumulation in a mouse model of Alzheimer's disease.
J. Pharmacol. Exp. Ther.
332
:
505
514
.
16
Taler
M.
,
Bar
M.
,
Korob
I.
,
Lomnitski
L.
,
Baharav
E.
,
Grunbaum-Novak
N.
,
Weizman
A.
,
Gil-Ad
I.
.
2008
.
Evidence for an inhibitory immunomodulatory effect of selected antidepressants on rat splenocytes: possible relevance to depression and hyperactive-immune disorders.
Int. Immunopharmacol.
8
:
526
533
.
17
Jiang
J.
,
Kuhara
T.
,
Ueki
R.
,
Zheng
Y.
,
Suto
H.
,
Ikeda
S.
,
Ogawa
H.
.
2007
.
Inhibitory effects of paroxetine on the development of atopic dermatitis-like lesions in NC/Nga mice.
J. Dermatol. Sci.
47
:
244
247
.
18
Jackson-Lewis
V.
,
Przedborski
S.
.
2007
.
Protocol for the MPTP mouse model of Parkinson’s disease.
Nat. Protoc.
2
:
141
151
.
19
Kim
S. Y.
,
Choi
K. C.
,
Chang
M. S.
,
Kim
M. H.
,
Kim
S. Y.
,
Na
Y. S.
,
Lee
J. E.
,
Jin
B. K.
,
Lee
B. H.
,
Baik
J. H.
.
2006
.
The dopamine D2 receptor regulates the development of dopaminergic neurons via extracellular signal-regulated kinase and Nurr1 activation.
J. Neurosci.
26
:
4567
4576
.
20
Kim
S. R.
,
Lee
D. Y.
,
Chung
E. S.
,
Oh
U. T.
,
Kim
S. U.
,
Jin
B. K.
.
2005
.
Transient receptor potential vanilloid subtype 1 mediates cell death of mesencephalic dopaminergic neurons in vivo and in vitro.
J. Neurosci.
25
:
662
671
.
21
Paxinos
G.
,
Franklin
K.
.
2001
.
The Mouse Brain in Stereotaxic Coordinates.
Academic Press
,
San Diego, CA
.
22
West
M. J.
,
Slomianka
L.
,
Gundersen
H. J.
.
1991
.
Unbiased stereological estimation of the total number of neurons in thesubdivisions of the rat hippocampus using the optical fractionator.
Anat. Rec.
231
:
482
497
.
23
Ferger
B.
,
Leng
A.
,
Mura
A.
,
Hengerer
B.
,
Feldon
J.
.
2004
.
Genetic ablation of tumor necrosis factor-α (TNF-α) and pharmacological inhibition of TNF-synthesis attenuates MPTP toxicity in mouse striatum.
J. Neurochem.
89
:
822
833
.
24
Ryu
M. Y.
,
Lee
M. A.
,
Ahn
Y. H.
,
Kim
K. S.
,
Yoon
S. H.
,
Snyder
E. Y.
,
Cho
K. G.
,
Kim
S. U.
.
2005
.
Brain transplantation of neural stem cells cotransduced with tyrosine hydroxylase and GTP cyclohydrolase 1 in Parkinsonian rats.
Cell Transplant.
14
:
193
202
.
25
Choi
S. H.
,
Lee
D. Y.
,
Chung
E. S.
,
Hong
Y. B.
,
Kim
S. U.
,
Jin
B. K.
.
2005
.
Inhibition of thrombin-induced microglial activation and NADPH oxidase by minocycline protects dopaminergic neurons in the substantia nigra in vivo.
J. Neurochem.
95
:
1755
1765
.
26
Park
K. W.
,
Jin
B. K.
.
2008
.
Thrombin-induced oxidative stress contributes to the death of hippocampal neurons: role of neuronal NADPH oxidase.
J. Neurosci. Res.
86
:
1053
1063
.
27
Przedborski
S.
,
Jackson-Lewis
V.
,
Djaldetti
R.
,
Liberatore
G.
,
Vila
M.
,
Vukosavic
S.
,
Almer
G.
.
2000
.
The parkinsonian toxin MPTP: action and mechanism.
Restor. Neurol. Neurosci.
16
:
135
142
.
28
Giovanni
A.
,
Sieber
B. A.
,
Heikkila
R. E.
,
Sonsalla
P. K.
.
1991
.
Correlation between the neostriatal content of the 1-methyl-4-phenylpyridinium species and dopaminergic neurotoxicity following 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine administration to several strains of mice.
J. Pharmacol. Exp. Ther.
257
:
691
697
.
29
Block
M. L.
,
Zecca
L.
,
Hong
J. S.
.
2007
.
Microglia-mediated neurotoxicity: uncovering the molecular mechanisms.
Nat. Rev. Neurosci.
8
:
57
69
.
30
Wu
D. C.
,
Jackson-Lewis
V.
,
Vila
M.
,
Tieu
K.
,
Teismann
P.
,
Vadseth
C.
,
Choi
D. K.
,
Ischiropoulos
H.
,
Przedborski
S.
.
2002
.
Blockade of microglial activation is neuroprotective in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine mouse model of Parkinson disease.
J. Neurosci.
22
:
1763
1771
.
31
Henkel
J. S.
,
Beers
D. R.
,
Siklós
L.
,
Appel
S. H.
.
2006
.
The chemokine MCP-1 and the dendritic and myeloid cells it attracts are increased in the mSOD1 mouse model of ALS.
Mol. Cell. Neurosci.
31
:
427
437
.
32
Klevenyi
P.
,
Andreassen
O.
,
Ferrante
R. J.
,
Schleicher
J. R.
 Jr.
,
Friedlander
R. M.
,
Beal
M. F.
.
1999
.
Transgenic mice expressing a dominant negative mutant interleukin-1β converting enzyme show resistance to MPTP neurotoxicity.
Neuroreport
10
:
635
638
.
33
Sriram
K.
,
Matheson
J. M.
,
Benkovic
S. A.
,
Miller
D. B.
,
Luster
M. I.
,
O’Callaghan
J. P.
.
2002
.
Mice deficient in TNF receptors are protected against dopaminergic neurotoxicity: implications for Parkinson’s disease.
FASEB J.
16
:
1474
1476
.
34
Zhou
C.
,
Huang
Y.
,
Przedborski
S.
.
2008
.
Oxidative stress in Parkinson’s disease: a mechanism of pathogenic and therapeutic significance.
Ann. N. Y. Acad. Sci.
1147
:
93
104
.
35
Babior
B. M.
1999
.
NADPH oxidase: an update.
Blood
93
:
1464
1476
.
36
Cross
A. R.
,
Segal
A. W.
.
2004
.
The NADPH oxidase of professional phagocytes—prototype of the NOX electron transport chain systems.
Biochim. Biophys. Acta
1657
:
1
22
.
37
Yamaguchi
H.
,
Kajitani
K.
,
Dan
Y.
,
Furuichi
M.
,
Ohno
M.
,
Sakumi
K.
,
Kang
D.
,
Nakabeppu
Y.
.
2006
.
MTH1, an oxidized purine nucleoside triphosphatase, protects the dopamine neurons from oxidative damage in nucleic acids caused by 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine.
Cell Death Differ.
13
:
551
563
.
38
Block
M. L.
,
Hong
J. S.
.
2005
.
Microglia and inflammation-mediated neurodegeneration: multiple triggers with a common mechanism.
Prog. Neurobiol.
76
:
77
98
.
39
Miller
R. L.
,
James-Kracke
M.
,
Sun
G. Y.
,
Sun
A. Y.
.
2009
.
Oxidative and inflammatory pathways in Parkinson’s disease.
Neurochem. Res.
34
:
55
65
.
40
Oyagi
A.
,
Oida
Y.
,
Hara
H.
,
Izuta
H.
,
Shimazawa
M.
,
Matsunaga
N.
,
Adachi
T.
,
Hara
H.
.
2008
.
Protective effects of SUN N8075, a novel agent with antioxidant properties, in in vitro and in vivo models of Parkinson’s disease.
Brain Res.
1214
:
169
176
.
41
Seet
R. C.
,
Lee
C. Y.
,
Lim
E. C.
,
Tan
J. J.
,
Quek
A. M.
,
Chong
W. L.
,
Looi
W. F.
,
Huang
S. H.
,
Wang
H.
,
Chan
Y. H.
,
Halliwell
B.
.
2009
.
Oxidative damage in Parkinson disease: measurement using accurate biomarkers.
Free Radic. Biol, Med.
48
:
560
566
.
42
Aquilano
K.
,
Baldelli
S.
,
Rotilio
G.
,
Ciriolo
M. R.
.
2008
.
Role of nitric oxide synthases in Parkinson’s disease: a review on the antioxidant and anti-inflammatory activity of polyphenols.
Neurochem. Res.
33
:
2416
2426
.
43
Nagatsu
T.
,
Mogi
M.
,
Ichinose
H.
,
Togari
A.
.
2000
.
Cytokines in Parkinson’s disease.
J. Neural Transm.
(
Suppl.58
)
143
151
.
44
Heneka
M. T.
,
Wiesinger
H.
,
Dumitrescu-Ozimek
L.
,
Riederer
P.
,
Feinstein
D. L.
,
Klockgether
T.
.
2001
.
Neuronal and glial coexpression of argininosuccinate synthetase and inducible nitric oxide synthase in Alzheimer disease.
J. Neuropathol. Exp. Neurol.
60
:
906
916
.
45
Teismann
P.
,
Tieu
K.
,
Cohen
O.
,
Choi
D. K.
,
Wu
D. C.
,
Marks
D.
,
Vila
M.
,
Jackson-Lewis
V.
,
Przedborski
S.
.
2003
.
Pathogenic role of glial cells in Parkinson’s disease.
Mov. Disord.
18
:
121
129
.
46
Zhao
C.
,
Ling
Z.
,
Newman
M. B.
,
Bhatia
A.
,
Carvey
P. M.
.
2007
.
TNF-alpha knockout and minocycline treatment attenuates blood-brain barrier leakage in MPTP-treated mice.
Neurobiol. Dis.
26
:
36
46
.
47
Oki
C.
,
Watanabe
Y.
,
Yokoyama
H.
,
Shimoda
T.
,
Kato
H.
,
Araki
T.
.
2008
.
Delayed treatment with arundic acid reduces the MPTP-induced neurotoxicity in mice.
Cell. Mol. Neurobiol.
28
:
417
430
.
48
Bolin
L. M.
,
Strycharska-Orczyk
I.
,
Murray
R.
,
Langston
J. W.
,
Di Monte
D.
.
2002
.
Increased vulnerability of dopaminergic neurons in MPTP-lesioned interleukin-6 deficient mice.
J. Neurochem.
83
:
167
175
.
49
Hampton
M. B.
,
Kettle
A. J.
,
Winterbourn
C. C.
.
1998
.
Inside the neutrophil phagosome: oxidants, myeloperoxidase, and bacterial killing.
Blood
92
:
3007
3017
.
50
Lau
D.
,
Mollnau
H.
,
Eiserich
J. P.
,
Freeman
B. A.
,
Daiber
A.
,
Gehling
U. M.
,
Brümmer
J.
,
Rudolph
V.
,
Münzel
T.
,
Heitzer
T.
, et al
.
2005
.
Myeloperoxidase mediates neutrophil activation by association with CD11b/CD18 integrins.
Proc. Natl. Acad. Sci. USA
102
:
431
436
.
51
Petzinger
G. M.
,
Walsh
J. P.
,
Akopian
G.
,
Hogg
E.
,
Abernathy
A.
,
Arevalo
P.
,
Turnquist
P.
,
Vucković
M.
,
Fisher
B. E.
,
Togasaki
D. M.
,
Jakowec
M. W.
.
2007
.
Effects of treadmill exercise on dopaminergic transmission in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-lesioned mouse model of basal ganglia injury.
J. Neurosci.
27
:
5291
5300
.
52
Jiménez-Jiménez
F. J.
,
Tejeiro
J.
,
Martínez-Junquera
G.
,
Cabrera-Valdivia
F.
,
Alarcón
J.
,
García-Albea
E.
.
1994
.
Parkinsonism exacerbated by paroxetine.
Neurology
44
:
2406
.
53
Leo
R. J.
1996
.
Movement disorders associated with the serotonin selective reuptake inhibitors.
J. Clin. Psychiatry
57
:
449
454
.