α-Synuclein Activates Microglia by Inducing the Expressions of Matrix Metalloproteinases and the Subsequent Activation of Protease-Activated Receptor-1

Matrix metalloproteinases (MMPs) are zinc-dependent endopeptidases that degrade extracellular matrix and nonmatrix proteins. During development, MMPs play beneficial roles in neurogenesis and myelinogenesis and in axonal growth and guidance (1, 2). Within the adult brain, MMPs play critical roles in synaptic plasticity, angiogenesis, learning and memory, and myelin turnover. However, during various neuropathological conditions, the upregulation of MMPs is associated with breakdown of the blood–brain barrier, demyelination, neuronal death, and inflammation (1, 2). Furthermore, because MMP upregulation is usually observed during the early stage of CNS injuries, modulation in the expression and/or activities of MMPs is considered a promising therapeutic strategy (2).

A number of papers have reported on the role of MMPs in neurodegenerative diseases, such as Alzheimer’s disease, Parkinson’s disease (PD), and multiple sclerosis (3–5). Of these, Parkinson’s disease is the second most common neurodegenerative disease and affects ~1% of the population aged over 65 y (6). Several MMPs have been reported to be upregulated postmortem in the substantia nigra of the PD brain (7). Furthermore, it has been reported that MMP-9 is upregulated in 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-induced parkinsonism in mice and that its inhibition suppressed the death of dopaminergic neurons (4). In addition, MMP-3 was found to contribute to dopaminergic neuronal cell death by inducing intracellular apoptotic signaling pathways (8). These results suggest that MMPs play a crucial role in pathogenesis of PD.

α-Synuclein is a small, soluble protein (14 kDa) and a major component of Lewy bodies, which are the pathological hallmarks of PD and Lewy body dementia. Multiplication or missense mutations of the α-synuclein gene (A30P, A53T, and E46K) have been identified in familial cases of PD and have been found to promote aggregation of the protein and produce toxicity on dopaminergic neurons (9, 10). Furthermore, alterations of the ubiquitin–proteasome system, which are usually observed in PD, may hinder the degradation of aggregated α-synuclein and aggravate cell death (11, 12). Native unfolded α-synuclein protein also causes apoptosis by inducing the dissociation of 14-3-3 chaperone protein from BAD (a proapoptotic protein) (10). Furthermore, modifications of α-synuclein, such as nitration or oxidation, produce oxidative stress and induce dopaminergic neuronal cell death (13, 14). In addition, observations that nigral aggregates immunoreactive to α-synuclein are often surrounded by activated microglia indicate that neuroinflammation plays an important role in the pathogenesis of PD (15, 16).

Recently, several papers have reported that α-synuclein induces inflammatory reactions in microglia, astrocytes, and human monocytes in vitro and in vivo (17–20). Activated immune cells express various neurotoxic molecules, such as cytokines, PGE₂, and NO, and induce neuronal cell death. Furthermore, α-synuclein released from degenerating dopaminergic neurons has been reported to activate microglia (21, 22). In addition, a recent study reported that monomeric α-synuclein, but not its aggregated form, enhances microglial phagocytosis (23).

Although dozens of papers have reported that MMPs play an important role in neuronal cell death and neuroinflammation, the roles of MMPs derived from activated microglia remain unclear. To investigate the role of MMPs in α-synuclein–stimulated microglia, we identified MMPs and investigated their regulatory effects on the expression of various inflammatory/neurotoxic molecules and upstream signaling pathways. We found that MMPs are secreted from α-synuclein–stimulated microglia and in turn activate microglia by activating protease-activated receptor-1 (PAR-1) in an autocrine or paracrine manner.

Purified α-synuclein protein was obtained from r-Peptide (Athens, GA). The amount of endotoxin in purified α-synuclein was found to be <1 endotoxin unit/mg of protein. Aggregated α-synuclein was prepared by incubation of monomer α-synuclein with agitation at 37°C for 7 d by modification of a previous method (24). MMP inhibitors and cathepsin G were purchased from Calbiochem (La Jolla, CA). SCH-79797 dihydrochloride was obtained from Tocris Bioscience (Bristol, U.K.). Active recombinant MMP-3, -8, and -9 were purchased from BIOMOL (Plymouth Meeting, PA), and all of the enzymes and chemicals for RT-PCR were purchased from Promega (Madison, WI). Abs against MAPKs, α-synuclein, MMPs, and transcription factors (NF-κB and AP-1) were purchased from Cell Signaling Technology (Beverley, MA) or from Santa Cruz Biotechnology (Santa Cruz, CA). All of the other chemicals were obtained from Sigma-Aldrich (St. Louis, MO), unless otherwise stated.

Primary microglial cells were cultured from the cerebral cortices of 1- to 2-d-old Sprague-Dawley rat pups, as described previously (25). In brief, cortices were triturated to single cells and cultured in minimal essential medium containing 10% FBS. After 10–14 d, microglial cells were separated by shaking the culture plate for 30 min at 200 rpm and were then plated in 48-well plates (5 × 10⁴ cells per well) and allowed to settle for 1 h. Nonadhering cells were removed by washing with the same medium before experiments. The purity of microglial was >95%, which was determined by isolectin B4 staining (data not shown). Stable SH-SY5Y human neuroblastoma cells overexpressing wild-type or A53T mutant α-synuclein were previously described (26) and maintained in DMEM supplemented with 10% heat-inactivated FBS, streptomycin (10 μg/ml), and penicillin (10 U/ml) in the presence of G418 (500 μg/ml).

Rat primary microglial cells (7 × 10⁶ cells on a 6-cm dish) were treated with α-synuclein in the presence of MMP or PAR-1 inhibitors, and total RNA was extracted from cells with TRIzol reagent (Invitrogen, Carlsbad, CA). For RT-PCR, total RNA (1 μg) was reverse-transcribed in a reaction mixture containing 1 U RNase inhibitor, 500 ng random primers, 3 mM MgCl₂, 0.5 mM dNTP, 1× RT buffer, and 10 U reverse transcriptase (Promega). The synthesized cDNA was used as a template for PCR, in which Go Taq polymerase (Promega) and the primers for MMPs, TNF-α, IL-1β, inducible NO synthase (iNOS), and GAPDH were used as shown in Table I. Analysis of the resulting PCR products on 1% agarose gels showed single-band amplification products of the expected sizes.

Microglial cells were seeded onto 12-mm coverslips 24 h prior to being treated with stimulants. The cells were then rinsed twice with PBS, fixed with 4% paraformaldehyde solution for 10 min at 4°C, rinsed with PBS, and incubated in permeabilization buffer (0.5% Triton X-100 and 1% BSA in PBS) for 2 h at room temperature. Primary Abs against MMP-3 or MMP-9 (1:200; Santa Cruz Biotechnology) were applied to the cells, which were then incubated at 4°C overnight. After three rinses with PBS, fluorescent dye-labeled secondary Abs or Hoechst 33258 fluorescent staining dyes were applied for 1 h at room temperature, and the cells were mounted on slides and observed under a fluorescence microscope (Zeiss, Oberkochen, Germany).

Primary microglia (2.5 × 10⁵ cells per well in a 48-well plate) were pretreated with MMP or PAR-1 inhibitors for 30 min and then stimulated with α-synuclein. Supernatants of cultured microglia were collected after 24 h of α-synuclein stimulation, and TNF-α and IL-1β concentrations were measured by ELISA using mAbs as recommended by the manufacturer (BD Pharmingen, San Diego, CA). Accumulated nitrite was measured in cell supernatants using Griess reagent (Promega).

Nuclear extracts from treated microglia were prepared as previously described (27). Double-stranded DNA oligonucleotides containing the NF-κB or AP-1 consensus sequences (Promega) were end-labeled using T4 polynucleotide kinase (NEB, Beverly, MA) in the presence of [γ-³²P]ATP. One microgram of nuclear proteins were incubated with ³²P-labeled NF-κB or AP-1 probe on ice for 30 min and resolved on 5% acrylamide gels, as previously described (27). For supershift assays, Abs against the p65 or p50 subunits of NF-κB (Santa Cruz Biotechnology) were coincubated with nuclear extract mixes for 30 min at 4°C before adding the radiolabeled probe.

To detect MAPK activation, cells were treated with MMP inhibitors before being stimulated with α-synuclein, and total cell lysates were prepared as previously described (27). To detect secreted α-synuclein, stable SH-SY5Y cell lines were cultured in DMEM containing 0.1% FBS for 24 h, and conditioned media were collected. The above-mentioned total cell lysates or conditioned media were separated by SDS-PAGE and transferred to polyvinylidene difluoride membranes. Membranes were blocked with 5% skimmed milk in 10 mM Tris-HCl containing 150 mM NaCl and 0.5% Tween 20 (TBST) and then incubated with primary Abs (1:1000) that recognized the phospho- or total forms of MAPKs or α-synuclein. After membranes were washed thoroughly with TBST, HRP-conjugated secondary Abs (1:3000 dilution in TBST; NEB) were applied and blots were developed using an ECL kit (Pierce, Rockford, IL)

Intracellular accumulations of reactive oxygen species (ROS) were measured using a modification of previously described methods (28). In brief, microglial cells were stimulated with α-synuclein for 24 h and then stained with 50 μM 2′,7′-dichlorodihydrofluorescein-diacetate in HBSS buffer for 30 min at 37°C. Dichlorofluorescein (DCF) fluorescence intensities were measured using excitation and emission wavelengths of 485 and 535 nm, respectively, using a fluorescence plate reader (Molecular Devices, Sunnyvale, CA).

The interactions between MMPs and PAR-1 were investigated using a synthetic peptide containing extracellular residues of PAR-1 receptor (TR33-62; Peptron, Daejon, South Korea). The peptide had the sequence A³³TNATLDPRSFLLRNPNDKYEPFWEDEEKN⁶², which contains a thrombin cleavage site between R41 and S42 (29). Cleavage assays were performed in 5 mM Tris, 0.5% polyethylene glycol, and 145 mM NaCl at pH 7.4 and 37°C for 1 h and using 2 μM TR33-62 peptide and 0.5–1 nM enzyme (thrombin, MMPs, or α-synuclein). Reactions were stopped by adding perchloric acid to a final concentration of 0.2 M. To eliminate the possibility that cleavage was due to thrombin contamination, 5 nM hirudin was added to each reaction buffer. TR33-62 and its cleavage products were separated and analyzed using an ÄKTA fast protein liquid chromatography (FPLC) system (GE Healthcare, Piscataway, NJ) by measuring ultraviolet absorbance (215 nm). Samples of cleavage products were injected onto a column (C18 Vydac narrow bore column 2.1 mm × 250 mm, 5 μm [GRACE, Deerfield, IL]) using a syringe supplied with the FPLC system. The gradient was started at 0% acetonitrile (ACN) containing 0.1% trifluoroacetic acid, linearly increased to 100% ACN over 30 min, and then held at 100% ACN for 15 min. The ÄKTA FPLC unit was operated using UNICORN control software (GE Healthcare).

PAR-1 cleavage products were also analyzed by electrospray mass spectroscopy (Q-TOF Premier; Waters, Milford, MA). Nanoscale liquid chromatography separations of digested peptides were performed using a nanoAcquity system (Waters), equipped with a Symmetry C18 5 μm, 5 mm × 300 μm precolumn and a bridged ethyl hybrid C18 1.7 μm, 25 cm × 75 μm analytical reverse-phase column (Waters). Five microliter samples were initially transferred in an aqueous 0.1% formic acid solution to the precolumn at a flow rate of 10 μl/min for 3 min. The mobile phase A was composed of water containing 0.1% formic acid, and the mobile phase B was 0.1% formic acid in ACN. Peptides were separated using a 3–40% gradient of mobile phase B over 30 min at a flow rate of 300 nl/min, followed by a 10 min rinse with 90% mobile phase B. The mass spectrometer was operated in analysis v-mode and had resolving power of at least 10,000 full width at half maximum. All of the analyses were conducted in positive nanoelectrospray ion mode.

Unless otherwise stated, all of the experiments were performed in triplicate and repeated at least three times. Results are presented as means ± SDs, and comparisons between groups were performed using one-way ANOVA followed by the Student t test. Statistical significance was accepted for p values <0.05.

To determine whether α-synuclein induces microglial activation, rat primary microglia were treated with α-synuclein protein (1–10 μM) for 24 h. α-Synuclein was found to significantly induce proinflammatory cytokines and to increase the production of NO and ROS in a concentration-dependent manner (Fig. 1A). Furthermore, α-synuclein induced mRNA expression of proinflammatory cytokines (TNF-α and IL-1β) and iNOS (Fig. 1B), and the levels of cytokines and iNOS induced by 10 μM α-synuclein were comparable to those induced by LPS (10 ng/ml). Endotoxin is often contaminated in purified protein preparations. In the current study, however, a well-known endotoxin antagonist polymyxin B did not alter the microglial activation induced by α-synuclein, whereas it almost completely blocked LPS-induced NO, TNF-α, and ROS production. This finding implies that α-synuclein may be a genuine microglial activator (Supplemental Fig. 1). We further found that aggregated α-synuclein induced microglial activation. The production of TNF-α, IL-1β, and ROS induced by the monomer form of α-synuclein was higher than that induced by the aggregated form, whereas the level of NO was almost similar between two forms (Supplemental Fig. 2). α-Synuclein also induced the mRNA expression of MMP-1, -3, -8, and -9 (Fig. 1C), and the protein expression of MMP-3 and MMP-9 was confirmed by immunocytochemical analysis in the cytoplasm of α-synuclein–treated microglial cells (Fig. 1D).

To examine whether neuron cell-derived α-synuclein activates microglia, primary microglial cells were incubated with the conditioned media obtained from stable SH-SY5Y cells overexpressing wild-type or A53T mutant α-synuclein. The release of α-synuclein into conditioned media was confirmed by Western blotting (Fig. 2A). Conditioned media from stable clones harboring the α-synuclein gene significantly increased the production of NO, IL-1β, TNF-α, and ROS in primary microglia compared with that of the mock control with only pcDNA empty vector (Fig. 2B). The effects of the A53T clone were slightly higher than those of the wild type. These results confirm previous reports that α-synuclein secreted by neuronal cells via an exocytosis mechanism activates microglia (21, 22). The reason why the levels of NO and cytokines induced by α-synuclein in neuronal cell media are higher than those by the same concentration of recombinant protein (1 μM) may be due to some immunostimulatory factors (i.e., MMP-3, α-synuclein, and neuromelanin) released from neuronal cells in serum-deprived conditions (21). In support of this, neutralization of α-synuclein in media dramatically inhibited NO, ROS, and cytokines to lower levels than those in the mock control, possibly because the neutralizing Ab blocked even the basal levels of α-synuclein (Supplemental Fig. 3).

To investigate the roles of MMP-3 and MMP-9 expressed in α-synuclein–stimulated microglia, microglial cells were pretreated with MMP-3– or MMP-9–specific inhibitors before stimulation with α-synuclein. As shown in Fig. 3A, both the MMP-3 inhibitor (N-isobutyl-N-(4-methoxyphenylsulfonyl)-glycylhydroxamic acid [NNGH]) and the MMP-9 inhibitor (M9I) significantly suppressed α-synuclein–induced NO, TNF-α, and IL-1β production. Furthermore, MMP inhibitors significantly suppressed α-synuclein–induced ROS production (Fig. 3B). These results suggest that MMP-3 and MMP-9 play an important role in α-synuclein–induced microglial activation. In addition, it was interesting to note that the MMP-8 inhibitor (M8I) specifically suppressed TNF-α production. The IC₅₀ of the MMP-8 inhibitor was <1 μM, whereas the IC₅₀ values of the MMP-3 and MMP-9 inhibitors were ~10 and ~50 μM, respectively (Fig. 3C). In contrast, the efficacies of the MMP-8 inhibitor with respect to the suppression of other proinflammatory cytokines (e.g., IL-1β and IL-6) and iNOS were similar to those of the MMP-3 and MMP-9 inhibitors (data not shown). Microscopic examination of cell morphology and the MTT cell viability assay showed that the anti-inflammatory effects of MMP inhibitors are not a result of cell death (Fig. 3D, Supplemental Fig. 4).

To investigate the molecular mechanism underlying immunosuppression by MMP inhibition, we first examined the effects of the MMP-3 and MMP-9 inhibitors on the activities of NF-κB and AP-1, which are important transcription factors for the expression of various cytokines and iNOS (30). α-Synuclein was found to significantly increase the DNA binding activities of NF-κB and AP-1 (Fig. 4A, 4C), and this binding was suppressed by the MMP-3 and MMP-9 inhibitors (Fig. 4B, 4D). Next, we examined the effects of MMP inhibitors on MAPK activities, which are upstream signaling molecules of the inflammatory reactions of activated microglia. α-Synuclein was found to induce the phosphorylation of the three MAPKs within 30 min, and treatment with MMP-3 or MMP-9 inhibitor remarkably suppressed the phosphorylation of all three MAPKs (Fig. 5). These results suggest that NF-κB, AP-1, and MAPKs are involved in the anti-inflammatory mechanism of MMP inhibitors in α-synuclein–activated microglia.

On the basis of the above results, we hypothesized that MMPs secreted by α-synuclein–stimulated microglia might activate microglia in an autocrine or paracrine manner and that this activation is blocked by MMP inhibitors. During our search for cell surface receptors involved in MMP-mediated microglial activation, we noted that PAR-1 had been reported to mediate MMP-1–induced endothelial activation and tumorigenesis and thrombin-induced microglial activation (31, 32). Thus, to test the possible involvement of PAR-1 in α-synuclein–induced microglial activation, we examined the effect of the broad spectrum PAR-1 inhibitor cathepsin G and of the PAR-1 antagonist (SCH-79797) on α-synuclein–induced inflammatory responses. As shown in Fig. 6, both cathepsin G and SCH-79797 significantly inhibited the production of NO, TNF-α, IL-1β, and ROS induced by α-synuclein. The effects of PAR-1 inhibitors are not due to their cytotoxicities, as shown by morphological analysis and MTT assay data (Fig. 6C, Supplemental Fig. 4). Furthermore, cathepsin G and SCH-79797 both inhibited the mRNA expression of TNF-α, IL-1β, and several MMPs (Fig. 6D). Overall, the extent of inhibition by cathepsin G was much higher than that by SCH-79797, suggesting that other PAR family receptors (e.g., PAR-2 or -3) in addition to PAR-1 participate in α-synuclein–induced microglial activation.

During the process of PAR-1 activation, the N-terminal extracellular domain of PAR-1 is cleaved by proteases and the remaining domain acts as a tethered ligand and generates intracellular signals (33). To address the possibility that MMPs secreted from microglia might activate microglia via a feedback mechanism, we performed in vitro PAR-1 cleavage assays using the catalytic forms of MMPs described in 1Materials and Methods. PAR-1 peptide containing its N-terminal 33 aas encompassing its conventional cleavage sites was synthesized and incubated with MMPs or α-synuclein protein. Thrombin was used as a positive control, because it digests the peptide bond between R41 and S42. Cleavage products were analyzed by FPLC and mass spectrometry (MS).

As shown in Fig. 7A, top panel, the PAR-1 intact form was eluted as a major peak by FPLC and was found to have an m/z (4+) of 903.4775 by MS (Fig. 7C, top panel). After the peptide was treated with thrombin, the major intact peak decreased and two new peaks were generated (Fig. 7A, middle panel), which were found by MS to have m/z values of 479.7804 (2+) and 668.3423 (4+), respectively (Fig. 7C). These results indicated that the first new peak (b) was ATNATLDPR and that the second new peak (c) was SFLLRNPNDKYEPFWEDEEKN. In the presence of hirudin (a specific thrombin inhibitor) the cleavage of PAR-1 peptide by thrombin was almost completely inhibited (Fig. 7A, bottom panel).

As shown in Fig. 7B, top panel, α-synuclein did not cleave PAR-1 peptide. However, MMP-8 generated two new peptides, which corresponded to those produced by thrombin (Fig. 7B, middle panel). To eliminate the possibility of thrombin contamination, all of the reactions were performed in the presence of hirudin, a specific thrombin inhibitor. To confirm that the cleavage reaction was specifically induced by MMP-8, M8I (the MMP-8 inhibitor) was added to the reaction mixture, and as expected, no meaningful products were produced, thus indicating that PAR-1 is a substrate of MMP-8 (Fig. 7B, bottom panel). Fig. 7D summarizes the enzyme activities of each protein tested by comparing the areas of the three FPLC peaks corresponding to intact peptide (a), the smaller fragment (ATNATLDRP) (b), and the larger fragment (SFLLRNPNDKYEPFWEDEEKN) (c). PAR-1 cleavage capacities of the MMPs were ranked as MMP-8 > MMP-3 > MMP-9. The specificities of the PAR-1 cleavage reactions were confirmed using MMP-specific inhibitors, as shown in Fig. 7D, right panel.

Previous studies have demonstrated that overexpression of α-synuclein plays a pivotal role in progression of PD and have suggested that it acts by activating microglia (34–37). The present study demonstrates that extracellularly applied α-synuclein induces microglial activation, proinflammatory/neurotoxic gene expression, and inflammatory signaling pathways. In addition, it was found that α-synuclein induces MMP-1, -3, -8, and -9 in rat primary microglia. Furthermore, experiments with MMP-specific inhibitors revealed that these MMPs play important roles during α-synuclein–induced microglial activation and that MMP-3, MMP-8, and MMP-9 inhibitors suppress iNOS and the expression of proinflammatory cytokine genes. MMP inhibitors were also found to block the activities of upstream signaling molecules, such as ROS, MAPKs, and NF-κB/AP-1. However, we were unable to evaluate the role of MMP-1 because of the lack of a commercially available MMP-1–specific inhibitor. Our findings indicate that MMPs are secreted by α-synuclein–stimulated microglia and that these MMPs in turn activate microglia by activating PAR-1 in an autocrine or paracrine manner.

PARs are G protein-coupled receptors that signal in response to extracellular proteases. After the N-terminal extracellular domain of PAR-1 has been digested by proteases, the remaining domain acts as a tethered ligand and generates intracellular signals (29). Previous studies have demonstrated that PAR-1 is a protease receptor located on the surfaces of microglia and that the binding of PAR-1 to its ligand activates microglia (38). To address the possible involvement of PAR-1 in α-synuclein–induced microglial activation, primary microglia were pretreated with PAR-1 inhibitors and then treated with α-synuclein. The inhibition of PAR-1 was found to significantly suppress the expression of iNOS, TNF-α, and IL-1β and ROS production. Interestingly, the effects of PAR-1 inhibitors were similar to those of MMP inhibitors, which led us to postulate that MMPs secreted from activated microglia might activate PAR-1 on cell surfaces of neighboring cells. This postulation is supported by the finding that PAR-1 is directly activated by interstitial collagenase MMP-1, which cleaves the PAR-1 exodomain at sites other than thrombin cleavage sites (39) (Fig. 7E).

In the current study, PAR-1 cleavage assays revealed that MMP-3, -8, and -9 cleaved and activated a PAR-1 peptide spanning the conventional PAR-1 cleavage site. However, α-synuclein did not cleave the PAR-1 peptide, as expected (Fig. 7). The efficacies of PAR-1 cleavage by MMPs reduced in the following order: MMP-8 > MMP-3 > MMP-9. To our knowledge, this is the first report of PAR-1 cleavage by MMP-3, MMP-8, or MMP-9. A recent report showed that thrombin (a ligand of PAR-1) induced CD95 expression via PAR-1 activation in N9 microglial cells (32). Interestingly, the sites of PAR-1 cleavage by MMP-3, -8, and -9 were the same as those of thrombin.

Although it is evident that PAR-1 is involved in α-synuclein–induced microglial activation, we cannot exclude the possible involvement of other PAR series receptors (e.g., PAR-2 or -3) because the inhibition of inflammatory reactions by the PAR-1 antagonist (SCH-79797) was rather incomplete as compared with that achieved by cathepsin G, a broad spectrum PAR inhibitor (40) (Fig. 6). In accordance with this possibility, we found that α-synuclein increased PAR-3 expression in microglia (Supplemental Fig. 5). Further studies are required to address whether PAR-3 is also involved in this inflammatory reaction. In contrast to the effect on PAR-3, α-synuclein did not affect PAR-1 expression at either mRNA or protein level, suggesting that α-synuclein activates microglia via posttranslational regulation of PAR-1 (i.e., PAR-1 cleavage) rather than via upregulating PAR-1 expression (Supplemental Fig. 5).

Of the pattern recognition receptors implicated in microglial activation, macrophage Ag complex-1 (Mac-1) and CD36 (class B scavenger receptor) have been recently reported to mediate α-synuclein–induced microglial activation (20, 41). Zhang et al. (41) reported that α-synuclein directly binds to Mac-1 receptors, which then activate NADPH oxidase to produce ROS. However, Su et al. (20) found that CD36 (a microglial scavenger receptor) mediates ROS production, ERK phosphorylation, and proinflammatory cytokine expression in α-synuclein–stimulated microglia. Thus, the mechanisms of microglial activation via Mac-1 and CD36 appear to be different from that of PAR-1, which is not directly activated by α-synuclein.

In the current study, we found that the PAR-1 antagonist SCH-79797 also inhibited LPS-induced inflammatory responses. The effects of SCH-79797 on NO, TNF-α, and ROS production in LPS-stimulated microglia were similar to those in α-synuclein–stimulated cells (data not shown). The results suggest that PAR-1 is a common receptor that mediates both the α-synuclein– and LPS-induced microglial activation probably via secreted MMPs.

In summary, the current study reveals the roles of MMPs during α-synuclein–induced microglial activation and the underlying molecular mechanisms involved. Recently, we reported that the inhibition of MMP-3 or MMP-9 also suppresses microglial activation in LPS-stimulated microglial cells (42). The results suggest that MMPs may play a role as common molecules that mediate inflammatory reactions and that PAR-1 is closely associated with the plausible mechanisms for MMP-mediated microglial activation. We conclude that modulation of the activities of MMPs and/or PAR-1 may provide new therapeutic strategies for the treatment of neurodegenerative diseases such as PD.

Disclosures The authors have no financial conflicts of interest.