Relationship between alpha synuclein phosphorylation, proteasomal inhibition and cell death: relevance to Parkinson’s disease pathogenesis


Address correspondence and reprint requests to Dr J. M. Cooper, Department of Clinical Neurosciences, Institute of Neurology, University College London, Rowland Hill Street, London NW3 2 PF, UK. E-mail:


Alpha synuclein can be phosphorylated at serine129 (P-S129), and the presence of highly phosphorylated α-synuclein in Lewy bodies suggests changes to its phosphorylation status has an important pathological role. We demonstrate that the kinase(s) responsible for α-synuclein S129 phosphorylation is constitutively active in SH-SY5Y cells and involves casein kinase 2 activity. Increased oxidative stress or proteasomal inhibition caused significant elevation of P-S129 α-synuclein levels. Under these conditions, similar increases in P-S129 α-synuclein were found in both sodium dodecyl sulphate lysates and Triton extracts indicating the phosphorylated protein was soluble and did not lead to aggregation. The rate of S129 phosphorylation was increased in response to proteasomal inhibition indicating a higher activity of the relevant kinase. Cells expressing the phosphorylation mimic, S129D α-synuclein increased cell death and enhanced sensitivity to epoxomycin exposure. Proteasomal inhibition markedly decreased S129D α-synuclein turnover suggesting proteasomal inhibition leads to the accumulation of P-S129 α-synuclein through an increase in the kinase activity and a decrease in protein turnover resulting in increased cell death. We conclude that S129 phosphorylation is toxic to dopaminergic cells and both the levels of S129 phosphorylated protein and its toxicity are increased with proteasomal inhibition emphasising the interdependence of these pathways in Parkinson’s disease pathogenesis.

Abbreviations used

casein kinase






G-protein related kinase




okadaic acid


Parkinson’s disease




serine 129 phosphorylation




sodium dodecyl sulphate



While the underlying cause of Parkinson’s disease (PD) in the majority of patients is not understood, increasing age and a positive family history are risk factors for PD indicating the ageing process and genetic factors play important roles in pathogenesis. This is supported by a small but growing number of patients where mutations have been identified in specific genes (e.g. α-synuclein, LRRK2, parkin, DJ1 and PINK1, Schapira 2006). The loss of dopaminergic neurons and presence of Lewy bodies in surviving neurons of the substantia nigra are key pathological features of PD alongside an increase in oxidative damage and dysfunction of mitochondrial respiratory chain and ubiquitin-proteasome systems (Schapira 2008). This has led to the suggestion that PD may have many causes leading to a common end point. The seminal observation that α-synuclein protein aggregates were abundant in Lewy bodies in idiopathic PD patients made an important link between a genetic cause of the disease (α-synuclein mutations) and the pathology in sporadic patients and placed α-synuclein at the centre of PD pathology (Spillantini et al. 1997).

α-Synuclein is widely expressed throughout the brain with a cytosolic location, although it has also been associated with synaptic vesicles, the plasma membrane, lysosomes, lipid rafts and nuclei. While a definitive function for α-synuclein has yet to be assigned it has been extensively studied and a number of interesting features have emerged including its binding to lipid membranes, similarity to the 14-3-3 chaperone proteins, propensity to aggregate and its role in dopamine regulation (Bennett 2005). In relation to dopamine metabolism α-synuclein expression has been associated with the inhibition of tyrosine hydroxylase (Perez et al. 2002), regulation of dopamine uptake (Wersinger and Sidhu 2005) and the inhibition of dopamine release (Larsen et al. 2006) all of which may help explain the selective involvement of dopaminergic neurones in PD.

α-Synuclein has at least three experimentally proven phosphorylation sites (serine 87 and 129 and tyrosine125), but little is currently known about the regulation of its phosphorylation nor how this influences its function. In vitro analyses have suggested casein kinase-1 and -2 (CK1, CK2), G-protein coupled receptor kinase-2 and 5 (GRK2, GRK5) and polo-like kinase 2 are important in the phosphorylation of S129, and may decrease the association of α-synuclein for lipids and decrease its inhibition of phospholipase D2 (Pronin et al. 2000), but the physiological importance of these observations is not clear.

The finding that α-synuclein in Lewy bodies was predominantly phosphorylated at serine129 in comparison to a low constitutive level of phosphorylation (Fujiwara et al. 2002; Anderson et al. 2006) suggests this modification maybe important in pathogenesis. The pathological significance of this is further supported by the increased phosphorylation at S129 in transgenic mice expressing human mutant A53T α-synuclein (Wakamatsu et al. 2007).

It is not currently known if α-synuclein aggregates are toxic or are a protective mechanism. Aggregation may remove more soluble α-synuclein protofibrils which are thought to have greater toxicity if they are stabilised for example by dopamine (Conway et al. 2001). α-synuclein aggregation is potentiated by the A53T pathological mutation and by exposure to a variety of stimuli observed in PD pathology (mitochondrial inhibition, oxidative stress and nitration) (Bennett 2005), suggesting potential pathways leading to Lewy body formation. The influence of S129 phosphorylation upon cell viability and α-synuclein aggregation has been addressed in Drosophila by expressing α-synuclein modified to prevent (S129A) or mimic (S129D) phosphorylation. While S129A α-synuclein expression decreased neuronal loss, S129D expression led to an accelerated loss of dopaminergic neurons (Chen and Feany 2005), which contrasted with α-synuclein aggregates which were more pronounced in the S129A than the S129D or wild-type (WT) α-synuclein expressing flies. This suggested S129 phosphorylation of α-synuclein did not lead to aggregation and in fact lack of phosphorylation may increase aggregation leading to decreased toxicity. This, however was not replicated in a recent rat model involving viral delivery of S129D α-synuclein (Gorbatyuk et al. 2008) or the decreased aggregates observed in SH-SY5Y cells expressing S129A α-synuclein but this may possibly relate to the co-expression of synphilin-1 (Smith et al. 2005).

The central role of α-synuclein in PD pathology and the suggestion that its phosphorylation may have a direct pathological role has led us to investigate what may influence the phosphorylation of α-synuclein and how this relates to the loss of dopaminergic neurons in PD.


Cell culture, plasmids and transfection

SH-SY5Y cells were grown under standard conditions (Korlipara et al. 2004). WT human α-synuclein cDNA with a 3′ haemagluttinin (HA) epitope (YPYDVPDYA) was cloned into pcDNA3.1 (Invitrogen, Paisley, UK), using site directed mutagenesis (Quikchange kit; Stratagene, Cedar Creek, TX, USA) codon 129 (tct) was mutated to gat to give S129D or to gct to give S129A. Following transfection (Superfect; Qiagen, Hilden, Germany) of each construct into normal SH-SY5Y cells, stable transfectants were selected with G418 (400 μg/mL, PAA labs, Pasching, Austria), clonal lines established and characterised.

Toxin treatment

Cells were grown to approximately 40–60% confluence, following the addition of epoxomycin (Epox) (14 nM, Merck, Nottingham, UK), rotenone (rot) (0.25 μM), MPP+ (1 mM) or paraquat (PQ) (150 μM), as specified in the results, cells were harvested by scraping. Where appropriate 50 μM 5,6-dichloro-1-β-d-ribofuranosylbenzimadazole (DRB; Biomol, Exeter, UK) was added for 30 min and/or 1 μM okadaic acid (OKA; Merck) was added for up to 30 min prior to harvesting. Reagents were supplied by Sigma-Aldrich (Dorset, UK) unless otherwise stated.

Sample preparation

Cells were either solubilised in sodium dodecyl sulphate (SDS) buffer (0.1% SDS, 20 mM Tris–HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM Na3VO4 and 50 μM NaF) and the DNA digested with DNase I (Promega, Southampton, UK), or the cells were extracted into Triton buffer (1% Triton X-100, 20 mM Tris–HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM Na3VO4 and 50 μM NaF), centrifuged at 21 000 g for 10 min and the supernatant collected. All preparations were performed in the presence of a protease inhibitor cocktail (Sigma-Aldrich).

SDS–polyacrylamide gel electrophoresis and western blot

The NuPAGE gel system (Invitrogen) was used for all protein separations using 4–12% polyacrylamide gels and 2 (N morpholino) ethane sulphonic acid (MES) buffer, samples were solubilised in lithium dodecyl sulphate (LDS) sample buffer under reducing condition. Separated proteins were transferred to polyvinylidene difluoride membrane (Immobilon; Millipore, Watford, UK) following blocking with milk powder (10%) in phosphate-buffered saline. The following antibodies were used; anti-HA (1/5000, CoVance, Princeton, NJ, USA) for the ectopic expression of α-synuclein, anti-phospho serine 129 α-synuclein (1/2500, Wako, Neuss, Germany); CK2 (1/300) and P-S209 CK2 (1/2000, Abcam, Cambridge, UK); anti-β-actin (1/3500, Abcam). Primary antibodies were detected with horseradish peroxidase conjugated secondary antibodies (against mouse 1/3500, or rabbit 1/2000; Dako, Glostrup, Denmark) and blots were visualised using ECL reagents and Hyperfilm (GE Health, Bucks, UK). Band intensities were analysed using alpha DigiDoc gel analysis software (Alpha Innotech, San Leandro, CA, USA).

Cell death

Cell death was assessed by lactate dehydrogenase release using the CytoTox96 kit (Promega). Cells were seeded at 40–60% confluence in 24 well plates and cell death was evaluated after 48 h in the presence or absence of Epox (14 nM), identical wells were used to calculate total and released lactate dehydrogenase activities.

Protein turnover

α-Synuclein turnover was evaluated by growing the cells in cycloheximide (25 μg/mL), and where stated Epox (14 nM) was also added. At various times up to 48 h, cells were harvested, solubilised in 0.1% SDS, 10 mM Tris pH 7.4, protease inhibitor cocktail and digested with DNase I. Cell lysates were stained with 4′-6-diamidino-2-phenyl indole (DAPI 4 μg/mL) and fluorescence (Synergy; Biotek, Potton, UK; Em 360/Ex 460 nm) used to calculate samples of equal cell number for separation on SDS–polyacrylamide gel electrophoresis and western blots.


Cells were seeded onto glass coverslips and stained immunocytochemically using anti HA antibody as previously described (Tabrizi et al. 2000). Images were taken using an Axiophot fluorescence microscope and KS400 software (Zeiss, Welwyn Garden City, UK) with standardised exposure times.

Protein analysis

Protein levels were determined using the bicinchoninic acid protocol (Pierce, Cramlington, UK) using bovine serum albumen as standard.

Statistical analysis

All values represent mean ± SEM, and statistical analyses were performed using Student’s t-test or anova followed by the Dunnett post hoc test using Instat (Graphpad, LaJolla, CA, USA).


SH-SY5Y cells expressing α-synuclein (C205 cells) demonstrated a constitutive level of serine 129 phosphorylation (P-S129) which increased kinetically over 30 min of phosphatase inhibition by OKA (Fig. 1a). The specificity of the P-S129 antibody for this site was demonstrated by the complete lack of signal from cells expressing α-synuclein where the serine at codon 129 was replaced by alanine which cannot be phosphorylated (Fig. 1b). The higher molecular mass bands (marked with asterix) represent non-specific phosphorylated protein but were also present in cells lacking exogenous expression of α-synuclein.

Figure 1.

 Analysis of α-synuclein phosphorylation at S129 (P-S129). (a) C205 SH-SY5Y cells expressing wild-type α-synuclein with a C-terminal HA tag. The antibody detecting HA cross-reacted with a single band at 17 kDa while the P-S129 antibody cross-reacted with a 17 kDa band representing P-S129 α-synuclein and larger bands (asterix) which represented unidentified phosphorylated proteins. The α-synuclein and non-specific signal increased with longer exposure to okadaic acid (1 μM). (b) Extracts of cells expressing α-synuclein mutated to alanine at codon 129 (S129A), the P-S129 antibody did not cross-react with the 17 kDa S129A α-synuclein protein. The HA signal represents exogenous α-synuclein expression in these experiments.

To investigate the impact of the biochemical features associated with PD pathology upon the constitutive level of P-S129 α-synuclein, C205 cells were treated for 24 h with: PQ (150 μM) to increase oxidative stress, rot (0.25 μM) or MPP+ (1 mM) to inhibit complex I of the mitochondrial respiratory chain or Epox (14 nM) to inhibit the proteasome. These concentrations were chosen to be in their effective concentration range but giving minimal cell death over 24 h. Phosphorylated S129 α-synuclein levels in SDS lysates were markedly increased (increased 420%; = 0.006) by Epox treatment, mildly increased (40%) by PQ treatment, but unaffected by complex I inhibition by MPP+ or rot (Fig.  2a,b). To investigate the relative solubility of the phosphorylated α-synuclein cells were solubilised in 1% Triton buffer and analysed by western blot. The increase in P-S129 signals in the Triton extracts broadly reflected those observed in SDS lysates although the increase in P-S129 signal was more pronounced following PQ treatment and slightly lower following Epox treatment (Fig. 2c,d). This indicated that the phosphorylated pool of α-synuclein was relatively soluble and was consistent with undetectable levels of P-S129 α-synuclein in the Triton insoluble fractions (data not shown). The toxic agents used had little impact upon the overall levels and solubility of α-synuclein expression when related to β-actin levels (Fig. S1), suggesting the changes in P-S129 levels were not related to any changes in total α-synuclein levels or their relative solubility in Triton buffer.

Figure 2.

 Impact of oxidative stress, proteasome inhibition and complex I inhibition on S129 phosphorylation of α-synuclein. P-S129 relative to total ectopic α-synuclein (HA) levels in C205 SH-SY5Y cells following 24 h exposure to paraquat (PQ, 150 μM), epoxomycin (Epox, 14 nM), rotenone (rot, 0.25 μM) or MPP+ (1 mM). (a) Representative western blots and (b) P-S129 relative to HA signal in total protein extracts (SDS) from C205 SH-SY5Y cells; (c) representative western blots and (d) P-S129 relative to HA signal in 1% Triton X-100 extracts from C205 SH-SY5Y cells. Blots represent typical examples and values are the mean ± SEM of six analyses. Statistical comparisons for toxin treated samples versus untreated samples were performed using the Student’s t-test (*= 0.006, **< 0.0001).

The time course of the increased P-S129 levels with both PQ and Epox treatments were evaluated up to 48 h of treatment. The change in P-S129 levels up to 9 h of treatment were mild (Fig. S2) suggesting the response was a consequence of the gradual increase in the effects elicited by these agents, namely oxidative damage and decreased protein turnover. With PQ treatment there was a 200% increase in P-S129 levels after 24 h which increased by 260% (< 0.05) at 48 h (Fig. 3a,b). Epox treatment significantly increased the level of P-S129 α-synuclein by 326% (< 0.01) after 24 h which was maintained up to 48 h treatment (Fig. 3a,b).

Figure 3.

 Influence of time of exposure to paraquat and epoxomycin (Epox) upon P-S129 levels. (a) Western blot analyses for P-S129, HA or β-actin. C205 SH-SY5Y cells were exposed to paraquat (PQ, 150 μM) or Epox (14 nM) for up to 48 h before extraction in SDS (b) relative levels of P-S129: HA signals, mean ± SEM of four analyses. anova followed by Dunnett test were used to compare toxin treated versus untreated samples, *< 0.05; **< 0.01.

In the presence of the phosphatase inhibitor OKA we had demonstrated the kinase(s) responsible for the phosphorylation at S129 was active under normal conditions (Fig. 1). The rate of S129 phosphorylation was evaluated in the presence of OKA as a measure of the activity of the responsible kinase(s). We were therefore able to utilise this approach to determine if treatment with these agents influenced the level of S129 phosphorylation by influencing the responsible kinase activity. An increase in P-S129 signal with increasing time of OKA inhibition was evident for cells treated with all agents used (Fig. 4a–d, and additional blots shown in Fig. S3). This rate of phosphorylation was significantly decreased after 24 h of MPP+ treatment (< 0.05) and mildly decreased after 24 h of rot treatment (Fig. 4c,d) consistent with the levels of P-S129 seen earlier. There was a 98% increase (< 0.05) in the rate of S129 phosphorylation after 24 h of Epox treatment which was maintained at 36 h but no longer apparent at 48 h of treatment (Fig. 4b). In contrast there was no significant change in the rate of S129 phosphorylation with PQ treatment at either 24 or 48 h treatment (Fig. 4a). This suggests that inhibition of the proteasome enhanced S129 phosphorylation by increasing the responsible kinase activity, but PQ did not significantly influence the kinase activity at the time points analysed.

Figure 4.

 Influence of toxins upon the rate of S129 phosphorylation of α-synuclein. Western blot analyses (left panels) demonstrating the influence of OKA (1 μM) upon P-S129 signal with time relative to HA signal in the presence of; (a) paraquat (PQ, 150 μM), (b) epoxomycin (Epox, 14 nM), (c) MPP+ (1 mM) or (d) rotenone (rot, 0.25 μM) for 24 h (black bars), 36 h (light grey bar) and 48 h (dark grey bars). Rate of change in P-S129: HA levels after exposure to toxins is shown in the right panels. Mean ± SEM of four analyses. anova followed by Dunnett test was used to compare PQ or Epox treatment versus no treatment (*< 0.05); and Student’s t-test was used to compare MPP+ or rotenone treatment versus no treatment (**= 0.005).

Both CK2 and GRK5 have both been implicated in S129 phosphorylation of α-synuclein (Takahashi et al. 2006, 2007). Using the CK2 inhibitor DRB with C205 cells we measured P-S129 levels and the kinetic increase in P-S129 in the presence of OKA to investigate the involvement of CK2. In comparison to untreated cells the level of P-S129 signal in the cells treated with DRB was decreased (by 30%) and the rate of increase in the presence of OKA was decreased (by 46%, = 0.02) (Fig. 5a–c). The decrease in P-S129 signal suggests CK2 was involved with the phosphorylation of this site, however the incomplete inhibition implies the involvement of additional kinases. To identify if Epox or PQ influenced CK2 activation, CK2 levels were determined using western blot analysis relative to β-actin levels. However, both PQ and Epox caused no detectable induction or accumulation of CK2 (Fig. 5d,e), and there was no evidence that exposure of these agents for up to 24 h influenced the level of CK2 phosphorylated at serine 209 (Fig. S2).

Figure 5.

 Role of CK2 in α-synuclein S129 phosphorylation and influence of various toxins on CK2 levels. (a) Western blot analysis of P-S129 levels in the presence of the CK2 inhibitor 5,6-dichloro-1-β-d-ribofuranosylbenzimadazole (DRB) (50 μM) and okadaic acid (OKA) treatment. (b) basal P-S129 and (c) the increase in P-S129 in the presence of OKA were decreased by DRB. (d) Western blot analyses CK2 and β-actin levels in C205 cells exposed to paraquat (PQ, 150 μM), epoxomycin (Epox, 14 nM), rotenone (rot, 0.25 μM) or MPP+ (1 mM) for 24 h and their quantitation (e). Mean ± SEM of five analyses. Student’s t-test comparing the effect of treatment versus no treatment *= 0.02.

SH-SY5Y cell lines expressing S129D α-synuclein to mimic phosphorylation at this site were used to investigate if phosphorylation of α-synuclein at S129 had any impact upon basal cell death or sensitivity to Epox. Cells expressing S129D demonstrated a higher degree of cell death under basal growth conditions (Fig. 6a, = 0.04) and increased sensitivity to exposure to Epox (Fig. 6b, < 0.0001), although there was no increased sensitivity to PQ (data not shown). Cells expressing S129A α-synuclein demonstrated cell death similar to the WT α-synuclein expressing cells confirming the specificity of the toxicity to the S129D alteration.

Figure 6.

 Influence of phosphorylation at S129 upon cell death. Cell death was analysed using lactate dehydrogenase (LDH) release in SH-SY5Y cell lines expressing wild-type (WT), S129D or S129A α-synuclein, or control cells transfected with empty pcDNA3.1. (a) Basal cell death over a 48 h period under standard growth conditions; (b) Cell death after 48 h exposure to epoxomycin (14 nM). Values represent mean ± SEM of six analyses (three clones for each expressed protein). Student’s t-test was used to compare cell death in cells expressing α-synuclein versus empty vector, *= 0.04, **< 0.0001.

The turnover of ectopically expressed α-synuclein was determined in SH-SY5Y cell lines expressing similar levels of WT or S129D α-synuclein, and the impact of proteasome inhibition by Epox was analysed. The turnover of both WT and S129D α-synuclein were similar however there was a marked decrease in S129D α-synuclein turnover when the proteasome was inhibited by Epox, which was not evident for WT α-synuclein (Fig. 7a). In agreement with this observation immunocytochemical analyses demonstrated an increase in S129D α-synuclein levels with Epox treatment which was not apparent in cells expressing WT α-synuclein (Fig. 7b).

Figure 7.

 Influence of epoxomycin on S129D α-synuclein turnover and accumulation. (a) The turnover of wild-type or S129D α-synuclein was assessed in the presence of cycloheximide over 48 h under normal growth conditions or with treatment with epoxomycin (14 nM). Protein from equivalent cell numbers were loaded and the relative change in α-synuclein in untreated (solid line) or epoxomycin treated (dotted line) cells was determined. (b) Immunocytochemical staining with anti HA antibodies of cells expressing WT or S129D α-synuclein in untreated cells or after 48 h epoxomycin treatment. All staining conditions and exposures were standardised.


The presence of α-synuclein phosphorylated at S129 in Lewy bodies of PD brains suggests it may play an important role in α-synuclein metabolism and the pathogenesis of PD (Fujiwara et al. 2002; Anderson et al. 2006), with S129 phosphorylation and aggregation being recently suggested to play an important role in the toxic α-synuclein pathway in oligodendrocytes (Kragh et al. 2009), however this is not a universally held view (Gorbatyuk et al. 2008; Paleologou et al. 2008). To understand its relevance it is important to determine both the cause of the increase in phosphorylation and any consequences it may have on cell function and survival. It is conceivable that changes to its phosphorylation status could represent a response to the other biochemical events reported in PD pathogenesis. Included in this are a number of biochemical abnormalities which are recurrent themes in both PD brain samples and genetic models of PD including mitochondrial complex I dysfunction, oxidative stress and proteasome dysfunction (Schapira 2006). α-synuclein has been shown to be partially phosphorylated at S129 under normal conditions and we have confirmed that SH-SY5Y cells over-expressing WT α-synuclein also demonstrate a low level of S129 phosphorylation suggesting this is a convenient model to investigate factors that influence S129 phosphorylation.

In this study we have clearly demonstrated proteasome inhibition by Epox and increased oxidative stress by PQ both increased P-S129 α-synuclein levels, whereas inhibition of complex I by MPP+ or rot had little influence upon phosphorylation. This difference in response suggests that an increase in S129 phosphorylation was not a non-specific response to cellular stress and is in agreement with acute experiments using similar agents (Waxman and Giasson 2008).

The increase in S129 phosphorylation was apparent after 24 h and up to 48 h of Epox or PQ treatment clearly indicating that prolonged inhibition of the proteasome and increased oxidative stress rather than the toxins themselves were responsible. The increase in phosphorylation could represent an increase in the activity of the responsible kinase, decreased phosphatase activity or a decrease in the degradation of the phosphorylated protein. While we were able to detect an increase in the rate of phosphorylation with Epox treatment this was not observed with PQ treatment suggesting different mechanisms were involved.

In vitro experimentation has suggested CK1, CK2, GRK5 and GRK2 are all capable of phosphorylating α-synuclein at S129 (Pronin et al. 2000). However, there are no clear data demonstrating which kinase is responsible under normal cellular conditions or the signalling pathway that may regulate it. In the presence of OKA we were able to indirectly measure the kinase activity responsible for S129 phosphorylation and demonstrated CK2 was at least partly involved with this phosphorylation. However, we did not detect any increase in CK2 levels in response to Epox treatment suggesting either the increased S129 phosphorylation was mediated by an increased activation of CK2 activity or an increase in another kinase responsible for this phosphorylation. Other studies have also implicated the role of CK2 in S129 phosphorylation however increased levels of S129 phosphorylation have not been associated with changes to the levels of specific kinases (including CK2, CK1, GRK2 or GRK5) following the acute incubation of the proteasomal inhibitor MG132 or in mice transgenic for A53T α-synuclein although changes to CK2 activity have been implied (Waxman and Giasson 2008). More recent data suggests the polo-like kinase 2 may play an important role in α-synuclein S129 phosphorylation (Inglis et al. 2009) and further research is required to pin point not only the kinase involved but also the pathway leading to its in vivo activation.

Increased oxidative stress leads to partial inhibition of the proteasome (Elkon et al. 2004) and therefore may lead to an increase in P-S129 levels in a similar mechanism induced by Epox. However, our inability to detect an increase in the rate of S129 phosphorylation with PQ treatment would suggest a different mechanism is involved. While models demonstrating increased S129 phosphorylation of α-synuclein have not influenced CK2 levels, iron overload has been associated with both increased S129 phosphorylation and increased CK2 levels (Takahashi et al. 2007) suggesting iron overload influences S129 phosphorylation via an alternative pathway.

While α-synuclein is predominantly degraded by chaperone-mediated autophagy (Cuervo et al. 2004) there is evidence in cell free systems that various modifications to the protein including oxidation, nitration and the A53T mutation, prevent the degradation of α-synuclein by the chaperone-mediated autophagy pathway (Martinez-Vicente et al. 2008). Consequently, specific modifications to α-synuclein can lead to an increased dependency upon the proteasome for its degradation. Using S129D α-synuclein as a phospho mimic we demonstrated that under normal conditions it had a similar turnover to WT α-synuclein. However, under conditions of proteasomal inhibition there was a decrease in the rate of S129D α-synuclein degradation which was not observed for the WT protein in agreement with the role of the proteasome in the degradation of the phosphorylated protein in agreement with cell free data (Martinez-Vicente et al. 2008). This suggests that the increase in P-S129 levels with Epox treatment not only reflect the increased rate of phosphorylation but also the decrease in turnover of the phosphorylated protein which would help explain the higher levels of P-S129 α-synuclein after 48 h Epox treatment even in the absence of higher phosphorylation rates at this time point.

The increased P-S129 α-synuclein levels associated with Epox or PQ treatments were similarly soluble in triton and SDS lysates suggesting that they were not associated with insoluble aggregates over the duration of these experiments. This was supported by the lack of any obvious aggregates upon immunocytochemistry, implying phosphorylation of α-synuclein at S129 did not pre-dispose it to aggregate which is in agreement with the absence of increased α-synuclein aggregation in flies expressing S129D α-synuclein (Chen and Feany 2005), but contrasts with the increased insoluble S129 phosphorylated α-synuclein in A53T α-synuclein transgenic mice (Waxman and Giasson 2008). This suggests the pre-ponderance of S129 phosphorylated α-synuclein in Lewy bodies is not due to an inherent increase in aggregation of this form of the protein and may be in response to the presence of additional factors.

While Epox treatment leads to increased cell death it is not possible to identify what impact, if any, an increase in P-S129 α-synuclein has upon cell viability. However, using cells expressing the phospho mimic S129D α-synuclein we identified a greater cell death under normal growth conditions suggesting it is mildly toxic which is consistent with the accelerated loss of dopaminergic neurons in flies expressing S129D α-synuclein (Chen and Feany 2005), although this was not replicated in a recent rat model involving viral delivery of S129D α-synuclein (Gorbatyuk et al. 2008). Following exposure to Epox S129D expressing cells demonstrated an enhanced sensitivity suggesting the toxicity of phosphorylated α-synuclein is potentiated by proteasomal inhibition. This agrees with the dependence of S129D α-synuclein degradation upon the proteasome and implies the accumulation and toxicity of phosphorylated α-synuclein may involve the unfolded protein response (Sugeno et al. 2008). From this data we can speculate that the accumulation of S129 phosphorylated α-synuclein in Lewy bodies may be an attempt by neurones to sequester this toxic species in an environment where the proteasome is compromised.

In keeping with this study the substitution of aspartate or glutamate for serine have frequently been used to mimic phosphorylation of both proteins in general and α-synuclein specifically (Shioi et al. 2002; Chen and Feany 2005). These changes differ from phosphorylation in several features including an inability to be inter-converted been the phosphorylated and non-phosphorylated species but also in that they may have a different structural influence upon the protein which has been suggested for α-synuclein (Paleologou et al. 2008). However, there is also evidence aspartate modification is able to functionally replace serine phosphorylation (Shioi et al. 2002). While this caveat needs to be considered when interpreting data using S129D α-synuclein, the use of these mimetics provide invaluable cell and in vivo data for comparison with in vitro experimentation.

Our results support the notion that the pathogenesis of neuronal death in PD involves a complex network of interacting events. At the level of α-synuclein S129 phosphorylation, this is increased by reduced activity of the proteasome, a feature observed in the PD brain (McNaught and Jenner 2001) and involves an increase in kinase activity and a decreased turnover of the phosphorylated protein. S129 phosphorylation is toxic to dopaminergic cells and under conditions of impaired proteasomal function, the increase in phosphorylation and decrease in turnover lead to greater toxicity. These results emphasise the interdependence of the pathogenetic pathways in PD to dopaminergic cell loss and the importance of S129 phosphorylation and proteasomal inhibition as a potential trigger and amplifier.


This project was funded by the Peter Samuels Trust and Parkinson’s Disease Society (UK).