DJ-1 deficiency caused a defect of PGD 2 expression by attenuation of L-PGDS

and is involved in increasing HO-1 mRNA expression through the DP2 receptor and phosphorylated AKT signaling pathway in the retinal pigment epithelium (Kuesap, et al., 2008;Satarug, et al., 2008), I investigated whether secreted levels of PGD₂ were altered in DJ-1 KO-ACM. Using a PGD₂ specific ELISA assay, I found thatPGD₂ concentration was 50% less in DJ-1 KO-ACM compared with WT-ACM (Fig. 17A). To confirm, I antagonized the PGD2 receptor in BV2 cells and investigated HO-1 induction by ACM treatment.

Consistent with other reports, I found that HO-1 was barely expressed after treatment with a DP2 receptor-specific antagonist in ACM treated BV2 cells (Fig. 17B). It has been reported that PGD2 induced AKT activation leading to HO-1 expression. Thus, I confirmed AKT phosphorylation following treatment with WT- and DJ-1 KO-ACM in BV2 cells.

Interestingly, I found a 50% reduction in pAKT expression in DJ-1 KO-ACM treated BV2 cells compared with WT-ACM treated BV2 cells (Fig. 17C). To confirm whether AKT phosphorylation is a critical signaling pathway for HO-1 induction, I inhibited the AKT phosphorylation in BV2 cells and investigated HO-1 induction by ACM treatment.

Incubation for 6 h, I found that HO-1 protein was barely expressed following treatment with PI3K inhibitors (Fig 17D).

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Figure 17. DJ-1 deficiency attenuated astrocyte PGD₂ production. Astrocytes conditioned media (ACM) was prepared from WT and DJ-1 KO astrocytes. (A) The amount of PGD₂ was measured by ELISA. (B) BV2 cells were incubated with ACM in the absence or presence of antagonists of PGD₂ receptors (CAY1047 and TM30089 for DP2; BWA868C for DP1, 10 mM each) for 6 h. HO-1 protein expression was analyzed by western blot (left panel), and quantitated (right panel). (C) BV2 cells were incubated with each ACM for 30 min. Levels of pAKT were analyzed with western blot (left panel), and quantitated (right panel). (D) BV2 cells were incubated with WT-ACM with or without PI3K inhibitors as indicated for 6 h. HO-1 expression was analyzed by western blot and quantitated. Actin was used as a loading control (A-D). Data shown are representative of three independent experiments. Data are means ± SEMs of three samples (*P < 0.05; **P <0.005).

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The Next arising question is how PGD₂ production is attenuated by DJ-1 deficiency in astrocytes. Since, L-PGDS is glutathione (GSH)-independent enzyme and the major enzyme producing PGD₂ in the CNS. (Urade and Eguchi, 2002;Urade and Hayaishi, 2000),I investigated whether L-PGDS expression was altered in DJ-1 KO astrocytes. Using qPCR with an L-PGDS specific primer, I found that L-PGDS mRNA expression was 50% less in DJ-1 KO astrocytes compared with WT astrocytes (Fig. 18A). In addition, I found that L-PGDS mRNA expression demonstrated a 1.8-fold increased following 3xFlag DJ-1 transfection in DJ-1 KO astrocytes (Fig. 18B). To investigate whether L-PGDS protein expression is regulated by the altered mRNA expression, I confirmed L-PGDS protein expression in WT and DJ-1 KO astrocytes. Using Western blot with L-PGDS specific antibody, I found that L-PGDS protein expression showed a 50–60% decrease in DJ-1 KO astrocytes compared with WT astrocytes (Fig. 18C). In addition, I found that L-PGDS protein expression showed a 1.8-fold increase following 3xFlag DJ-1 transfection in DJ-1 KO astrocytes (Fig. 18D) although other prostaglandin synthases did not lead a difference between WT and DJ-1 KO astrocytes (Fig. 19A, B). Interestingly, I also found that L-PGDS protein was barely expressed in DJ-1 KO total brain lysates compared with WT total brain lysates (Fig 18E)

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Figure 18. DJ-1 deficient astrocytes reduced L-PGDS expression. (A) L-PGDS mRNA expression was analyzed in WT and DJ-1 KO astrocytes by RT-PCR and qPCR. (B) L-PGDS mRNA expression was analyzed in control or DJ-1 transfected astrocytes by RT-PCR and qPCR. (C) L-PGDS protein expression was analyzed in WT and DJ-1 KO astrocytes by Western blot. (D) L-PGDS protein expression was analyzed in control or DJ-1 transfected astrocytes by Western blot. Actin was used as a loading control (A-E). Data shown are representative of at least three independent experiments. Values are means ± SEMs of three samples (*P < 0.05; **P <0.005).

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Figure 19. Expression levels of prostaglandin synthases, H-PGDS, mPGES, PGIS, were not different in WT and DJ-1 KO astrocytes. Expression levels of H-PGDS, mPGES, PGIS, were analyzed by qPCR (A) and western blot and quantified (B). Actin was used as a loading control. Data shown here are representative of at least three independent experiments.

Values are means ± SEMs of three samples (*P < 0.05; **P <0.005).

J.

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Next question is whether attenuation of L-PGDS expression in astrocytes led a failure of anti-inflammatory function. Using L-PGDS specific siRNA, I confirmed L-PGDS mRNA knock down (KD) (Fig. 20A) and obtained L-PGDS KD-ACM. To confirm the failure of anti-inflammatory function following L-PGDS KD in astrocytes, I investigated the expression of nitrite and iNOS in BV2 cells after IFN-g treatment. Interestingly, I found that nitrite expression showed a 4-fold increase in L-PGDS KD incubated BV2 at 48 h after g treatment (Fig. 20B). In addition, I also found that iNOS expression showed a 1.8-fold increase in L-PGDS KD-ACM incubated BV2 at 12 h after IFN- g treatment (Fig. 20C).

Using Western blot with HO-1 specific antibody, I found that HO-1 expression showed a 1.5-fold increase in BV2 cells by NT-ACM treatment but not by L-PGDS KD-ACM treatment (Fig. 20D). Taken together, these results suggest that the reduction of anti-inflammatory function of DJ-1 deficient astrocytes was caused by attenuated PGD₂ secretion and L-PGDS expression.

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Figure 20. ACM from L-PGDS deficient astrocytes decreased anti-inflammatory effect.

(A) L-PGDS specific siRNA decreased L-PGDS expression. L-PGDS expression was analyzed by qPCR at the indicated times. (B-D) BV2 cells were incubated with IFN-g (20 ng/ml) in the presence of ACM prepared from NT-siRNA or L-PGDS-siRNA treated astrocytes. Nitrite levels were measured at 48 h (B). iNOS (C) and HO-1 (D) expression were analyzed with western blot and quantitated at 12 h and 6 h, respectively. Actin was used as a loading control. Data shown are representative of at least three independent experiments.

Values are means ± SEMs of three samples (*P < 0.05; **P <0.005).

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3. DJ-1 deficiency caused a defect of Sox9 protein expression through the regulation of ubiquitination.

It has been reported that Sox9 regulates L-PGDS mRNA expression (Wilhelm, et al., 2007) and that Sox9 is specifically expressed in astroytes in the adult brain (Pompolo and Harley, 2001). Thus, I examined whether Sox9 is specifically expressed in astrocytes and L-PGDS could be regulated by Sox9 expression in astrocytes as has been reviously reported.

Using Sox9 specific antibody, I found that Sox9 is expressed in astrocytes not in neurons using immunostaining (Fig. 21A) and Western blot (Fig. 21B). In addition, I found that L-PGDS mRNA and protein expression showed a 60% reduction in Sox9 siRNAs treated astrocytes compared with NT siRNA treated astrocytes (Fig. 21C, D).

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Figure 21. L-PGDS expression was regulated by Sox9. (A) Brain sections obtained from 8 w old mouse, and double-labeled with GFAP/Sox9 or NeuN/Sox9 antibodies. (B) Sox9 expression in purely cultured neurons and astrocytes was analyzed by Western blot. (C, D) Sox9 siRNAs decreased L-PGDS mRNA (C) and protein expression (D). Actin was used as a loading control. Data shown are representative of at least three independent experiments.

Values are means ± SEMs of three samples (*P < 0.05; **P <0.005). Scale bars, 10 mm (A)

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Next, I investigated whether Sox9 expression was affected in DJ-1 KO mice, both in the brain as a whole and astrocytes specifically. In the whole brain, using Western blot, Sox9 expression showed a 60% reduction in DJ-1 KO brain compared with WT brain (Fig. 22A).

Using immunostaining, I also found that Sox9 expression showed a 50–60% reduction in GFAP positive astrocytes in DJ-1 KO brain compared with WT brain (Fig. 22B). In addition, I found that Sox9 expression showed a 50–70% decrease in DJ-1 KO astrocytes compared with WT astrocytes by Western blot (Fig. 22C) and immunostaining (Fig. 22D). To confirm whether Sox9 was regulated by DJ-1 expression, I trasfected DJ-1 KO astrocytes with 3xflag-DJ-1 and I found that Sox9 expression showed a 1.5-fold increase in DJ-1 transfected astrocytes (Fig. 22E). Next, I investigated whether mutated DJ-1 also regulates Sox9 expression. Interestingly, I found that Sox9 expression was not rescued by E64D and C106A mutated DJ-1. L166P mutated DJ-1 did not cause a significant difference in Sox9 expression when compared with control group (Fig. 22F).

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Figure 22. DJ-1 regulated Sox9 expression in astrocytes. (A) Total brain lysates were obtained from 10 week old WT and DJ-1 KO mice. (B) WT and DJ-1 KO brain sections were double-labeled with GFAP and Sox9 antibodies. Sox9 intensities were quantified by Image J. (C, D) Sox9 protein expression was analyzed in WT and DJ-1 KO astrocytes by Western blot (C) and immunostaining (D). (E, F) Sox9 protein expression was analyzed in astrocytes transfected with mock vector or 3xFlag-WT DJ-1 (E) or 3xFlag- DJ-1 mutants, L166P, E64D, or C106A (F), with western blot and quantified. Actin was used as a loading control. Data shown are representative of at least three independent experiments. Values are means ± SEMs of three samples (*P < 0.05; **P <0.005). Scale bars, 10 mm (B, D).

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To better understand the link between DJ-1 and Sox9 protein expression, I next examined how Sox9 protein expression was regulated by DJ-1. First, I analyzed transcriptional regulation, such as Sox9 mRNA expression in WT and DJ-1 KO astrocytes using qPCR.

However, we detected no evidence of change in mRNA expression (Fig. 23A). Next, I analyzed post-transcriptional regulation. After treatment with cycloheximide, I found that level of Sox9 protein reduced a 20% at 2 h and a 50% at 3 h when compared with control level of Sox9 protein in WT astrocytes but it showed an 80% reduction at 2 h and complete degradation at 3 hour when compared with control level of Sox9 protein in DJ-1 KO astrocytes (Fig. 23B). Interestingly, it has been reported that Sox9 protein is degraded via ubiqutination and the proteasomal pathway (Akiyama, et al., 2005). Next, I examined whether DJ-1 regulates proteasomal degradation of Sox9 by ubiquitination. After MG132 treatment, I found that level of Sox9 protein showed a 50% reduction in DJ-1 KO astrocytes compared with WT astrocytes but it showed similar amount by inhibition of proeasomal degradation in WT and DJ-1 KO astrocytes (Fig. 23C).

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Figure 23. DJ-1 regulated Sox9 protein stability. (A) Sox9 mRNA expression was analyzed in WT and DJ-1 KO astrocytes by qPCR. (B, C) Sox9 protein levels were analyzed with western blot in WT and DJ-1 KO astrocytes in absence and presence of cycloheximide (CHX, 10 mg/ml) (B) or MG 132 (10 mg/ml) (C). Actin was used as a loading control. Data shown are representative of at least three independent experiments. Values are means ± SEMs of three samples (*P < 0.05; **P <0.005).

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To investigate the ubiquitination of Sox9 in WT and DJ-1 KO astrocytes, I performed co-immunoprecipitation (co-IP) with Sox9 and ubiquitin specific antibodies. After co-IP in WT and DJ-1 KO astrocytes, I found that the Sox9 and ubiquitin complex showed a 1.8-fold increase in DJ-1 KO astrocytes compared with WT astrocytes (Fig.

24A). To confirm this finding in the difference between WT and DJ-1 KO astrocytes, I utilized in situ proximity ligation assay (PLA). After PLA, I found that the Sox9 and ubiquitin complex showed a 5-fold increase in DJ-1 KO astrocytes (Fig. 24B).

Furthermore, I found that the Sox9 and ubiquitin complex showed a 60–70%

reduction following DJ-1 transfection compared with control vector transfection in DJ-1 KO MEF cells (Fig. 24C). Taken together, these results suggest that DJ-1 is a negative regulator ubiquitination of Sox9, which is a transcription factor of L-PGDS.

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Figure 24. DJ-1 negatively regulated Sox9 ubiquitination. (A) Immunoprecipitation (IP) was carried out in lysates prepared from WT and DJ-1 KO astrocytes using antibodies specific for Sox9 (or ubiquitin). Immunoprecipitated proteins were analyzed with Western blot using antibodies specific for ubiquitin (or Sox9), and quantitated. IgG was used as a negative control. (B) Interaction between Sox9 and ubiquitin was analyzed using in situ PLA.

PLA spots were counted using Image J. (C) IP was carried out in lysates prepared from DJ-1 KO MEF cells transfected with control vector or 3xFlag-DJ-1 as described above in (A).

Actin was used as a loading control (A, C). Data shown are representative of at least three independent experiments. Values are means ± SEMs of three samples (*P < 0.05; **P

<0.005). Scale bars, 20 mm (B, upper panel), 10 mm (B, lower panel).

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IV. DISCUSSION

Almost all studies examining the progression of PD were focused on neuronal death, since it has been reported that PD is characterized by dopaminergic neuronal death in the SNpc. However, recently studies suggest that risk factors for PD progression are not only dopaminergic neuronal death but also dysfunction of non-neuronal cells. Based on this, I examined functional change in astrocytes caused by a PD-related gene, DJ-1 and the relationship between dysfunction of astrocytes and progression of PD.

PART A. DJ-1, (PARK7), plays critical roles in astrogliosis by regulating of the pSTAT3 signaling pathway for tissue repair after brain injury

DJ-1, a gene related to an early onset autosomal-recessive form of PD (Bonifati, et al., 2003a). Interestingly, however, animal models of PD based on mutations of DJ-1 did not show PD phenotypes such as dopaminergic neuronal death and Lewy body formation (Chen, et al., 2005;Kim, et al., 2005;Kitada, et al., 2009). For example, it has been reported that the number of dopamine neurons was not decreased in 6 month and 11 month old DJ-1 null mice although striatal dopaminergic neurons were affected by DJ-1 deficiency, leading to increased dopamine reuptake rates and elevated tissue dopamine content (Chen, et al., 2005).

In addition, DJ-1null mice did not demonstrate any anatomical or neuronal abnormalities, reduction in the number of dopaminergic neuron or change in the dopamine levels in the striatum and in the substantia nigra although dopaminergic neuronal loss was increased by 1-methyl-4-phenyl-1,2,4,5-tetrahydrophyridine (MPTP) treatment (Kim, et al., 2005). These

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studies suggest that genetic mutation and environmental insults are necessary for PD progression.

After an environmental insult, the damaged brain tissues begin a process of repair and regeneration, in order to restore brain structure and function (Compston, 1995;Lo, 2010;Okano, et al., 2007). Unfortunately, however, CNS axons do not spontaneously regenerate, unlike peripheral nervous system (PNS) axons which have the capacity to regenerate and recover after injury (Benfey and Aguayo, 1982;David and Aguayo, 1981;Richardson, et al., 1984;Richardson, et al., 1980). However, mature CNS neurons are able to regenerate if provided with sufficient necessary factors. For example, it has been reported that mature CNS neurons have the capacity to regenerate after transplantation of peripheral nerve graft (Richardson, et al., 1980). This study suggests that regulation of brain environment is necessary for CNS repair and regeneration. Recently, it has been reported that PD patients or PD animal models demonstrate altered release of repair-related factors. For example, nerve growth factor (NGF) expression was significantly decreased in the serum of patients with early-stage PD and PD animal models (Lorigados Pedre, et al., 2002), and brain derived neurotrophic factors (BDNF) expression was increased in the serum of patients with PD (Ventriglia, et al., 2013). In addition, angiogenesis marker, integrin a_vb₃ was highly expressed in the SNpc in patients with parkinsonian syndromes and in the putamen in patients with PD (Desai Bradaric, et al., 2012). These studies strongly suggest that a change in the levels of repair-related factors was observed in patients with PD and could be a risk factor for progression of PD. In this study, I found that DJ-1 KO causes defects in the repair and regeneration of TH positive axons (Fig 1, 11). This result is the first direct evidence of a

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possible defect in tissue repair in a familial form of PD and I suggest that it is one of risk factors for progression of PD.

The next question I addressed was how DJ-1 deficiency reduces repair and regeneration after brain damage. Astrogliosis is related to repair-related factors such as the regulation of supporting nutrients, angiogenesis, remyelination, secretion of neurotrophic factors and neurogenesis. Thus many studies have suggested that astrogliosis has important roles in regeneration and repair (do Carmo Cunha, et al., 2007;Liberto, et al., 2004;Triolo, et al., 2006;White, et al., 2008), although, various studies have suggested negative effects of astrogliosis as a result of scar formation. For example, reactive astrocytes induced glycolytic capacity via glucose uptake, pyruvate kinase activity and lactate dehydrogenase (LDH) in hypoxic conditions, in order to support the energy needs of neurons (Marrif and Juurlink, 1999). In addition, astrocytes store glycogen after stimulation by IGF-1 and use it in glucose-limited conditions to provide energy support to neighboring neurons (Dringen and Hamprecht, 1992). Astrocytes also regulate the outgrowth of DRG neurites and axon regeneration in mature white matter through the action of fibronectin. In slice cultures of the P35 rat brain, astrocyte-associated fibronectin was detected. Furthermore, DRG neurites and axon regeneration were dramatically decreased by administration of anti-fibronectin antibody (Tom, et al., 2004). Reactive astrocytes are also associated with neovascularization and angiogenesis. In a traumatic injury model such as a stab wound and neural grafting, VEGF mRNA and protein expression and its receptor, flt-1, was highly expressed in reactive astrocytes, and not endothelium (Krum and Rosenstein, 1998). In hypoxic conditions, reactive astrocytes also regulate angiogenesis via VEGF. For example, co-culture of

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astrocytes and microvascular endothelial cells resulted in in vitro angiogenesis with the formation of tube-like structures. In this model, reactive astrocytes highly expressed VEGF (Ment, et al., 1997). Remyelination is also regulated by reactive astrocyte-released CNTF.

CNTF promotes the proliferation of oligodendrocyte precursors via fibroblast growth factor-2 (FGF-factor-2) (Albrecht, et al., factor-2003). CNTF is located in normal astrocytes and has been found in white matter structures as well (Dallner, et al., 2002). CNTF is located in normal astrocytes and has been found in white matter structures as well. CNTF protein and mRNA expression was highly increased during the remyelination phase and its expression was detected within reactive astrocytes surrounding the injured area. In addition, CNTF increased FGF-2, a factor needed for oligodendrocyte precursor cell (OPC) proliferation, in reactive astrocytes during remyelination in the spinal cord (Albrecht, et al., 2003). These studies suggest that astrogliosis maintains the required microenvironment for brain repair. In this study, I found that DJ-1 KO causes defects in the progression of astrogliosis after ATP-induced injury (Fig 2–8). This suggests the possibility of a defect in astrogliosis is present in a familial form of PD and I suggest that it causes a defect in brain repair.

Insufficient astrogliosis could affect the release of neurotrophic factors that mediate brain repair and regeneration. It has been reported that reactive astrocytes express neurotrophic factors necessary for protection and repair after brain injury. In particular, GDNF is expressed by human astrocytes; it also has been reported that GDNF is expressed in reactive astrocytes (Moretto, et al., 1996;Nakagawa and Schwartz, 2004). GDNF is a well-known glial derived neurotrophic factor and is a major supporting factor for dopaminergic neuronal protection in SNpc. GDNF supports the viability of postnatal nigral dopamine

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neurons by inhibiting apoptotic death (Burke, et al., 1998). In addition, GDNF induces dopaminergic nerve fiber sprouting in the margins of striatal wounds (Batchelor, et al., 1999), which is reduced by inhibition of GDNF via antisense oligonucleotides (Batchelor, et al., 2000). Furthermore, many studies suggest that GDNF regulates the protection and function of dopaminergic neurons. For example, GDNF injection induced significant improvement in the symptomatology and pathophysiological features of MPTP-induced parkinsonism in monkeys (Gash, et al., 1996). Adding GDNF via a lentiviral vector also reversed functional deficits monkeys with in MPTP-induced PD (Kordower, et al., 2000). Interestingly, GDNF induction was observed in patients with severe PD although GDNF receptor molecules, GFRa1 and cRET, were not changed (Backman, et al., 2006). These studies suggest that GDNF expression is necessary for neural protection and repair in the injured brain and PD.

In this study, I found that DJ-1 KO causes defects in GDNF expression in reactive astrocytes after ATP-induced injury (Fig 9, 10). This result represents a functional defect of insufficient astrogliosis. Thus, I suggest that functional defects of astrogliosis lead to delayed brain repair and are one of risk factors for progression of PD.

Intermediate filaments (IFs) play a critical role in the progression of astrogliosis.

After brain injury, IFs are highly expressed in reactive astrocytes and promote numerous features of morphological change such as hypertrophy of the cell body, and thickening and extension of cellular processes. For example, GFAP and vimentin double-KO showed insufficient features of reactive astrocytes such as low hypertrophy, process shortening, and reduction of glial scar formation, when compared with WT astrocytes after brain injury (Wilhelmsson, et al., 2004). Interestingly, it has been reported that brain damage was

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increased by a lack of GFAP and other IFs. For example, more GFAP-null mice died than

increased by a lack of GFAP and other IFs. For example, more GFAP-null mice died than