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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
J Neurosci. Author manuscript; available in PMC 2013 April 24.
Published in final edited form as:
PMCID: PMC3506191
NIHMSID: NIHMS417361

Systematic Mutagenesis of α-Synuclein Reveals Distinct Sequence Requirements for Physiological and Pathological Activities

Abstract

α-Synuclein is an abundant presynaptic protein that binds to phospholipids and synaptic vesicles. Physiologically, α-synuclein functions as a SNARE-protein chaperone that promotes SNARE-complex assembly for neurotransmitter release. Pathologically, α-synuclein mutations and α-synuclein overexpression cause Parkinson’s disease, and aggregates of α-synuclein are found as Lewy bodies in multiple neurodegenerative disorders (“synucleinopathies”). The relation of the physiological functions and pathological effects of α-synuclein remain unclear. As an initial avenue of addressing this question, we here systematically examined the effect of α-synuclein mutations on its physiological and pathological activities. We generated 26 α-synuclein mutants spanning the entire molecule, and analyzed them in comparison to wild-type α-synuclein in seven assays that range from biochemical studies with purified α-synuclein to expression of α-synuclein in cultured neurons to examination of the effects of virally expressed α-synuclein that was introduced into the mouse substrantia nigra by stereotactic injections. We found that both the N- and C-terminal sequences of α-synuclein were required for its physiological function as SNARE-complex chaperone, but that these sequences were not essential for its neuropathological effects. In contrast, point mutations in the region of α-synuclein referred to as non-amyloid β component (NAC; residues 61–95) as well as point mutations linked to Parkinson’s disease (A30P, E46K, and A53T) increased the neurotoxicity of α-synuclein but did not affect its physiological function. Thus, our data show that the physiological function of α-synuclein, although protective of neurodegeneration in some contexts, is fundamentally distinct from its neuropathological effects, thereby dissociating the two activities of α-synuclein.

INTRODUCTION

α-Synuclein is a small abundant neuronal protein that is natively unstructured, but folds into amphipathic α-helices in the presence of negatively charged lipids (Maroteaux et al., 1988; Perrin et al., 2000), binds to synaptobrevin-2/VAMP2 (Burré et al., 2010), and localizes to synaptic vesicles in nerve terminals (Iwai et al., 1995). In vitro and in cultured cells and neurons, α-synuclein promotes SNARE-complex assembly (Burré et al., 2010). Three synuclein-related genes are expressed in mammals that encode α-, β-, and γ-synuclein. α/β/γ-Synuclein triple knockout mice develop progressive neuropathology and motor impairments, die prematurely, and exhibit impaired SNARE-complex assembly, consistent with its function as a SNARE-complex chaperone (Burré et al., 2010; Greten-Harrison et al., 2010).

Aggregates of α-synuclein are found in age-dependent disorders called synucleinopathies, including Parkinson’s disease (PD), Alzheimer’s disease, multiple system atrophy, and dementia with Lewy bodies (Spillantini and Goedert, 2000; Masliah et al., 2001). Both point mutations in α-synuclein (A30P, E46K, A53T; Polymeropoulos et al., 1997; Kruger et al., 1998; Zarranz et al., 2004) and duplication or triplication of the α-synuclein gene (Singleton et al., 2003; Ibanez et al., 2004) produce PD. PD-linked α-synuclein mutations affect α-synuclein fibril formation in vitro (Conway et al., 1998; Narhi et al., 1999; Conway et al., 2000; Greenbaum et al., 2005; Fredenburg et al., 2007; Yonetani et al., 2009), and α-synuclein oligomers are toxic to neurons in vivo (Kayed et al., 2003; Lindersson et al., 2004; Tsika et al., 2010; Colla et al., 2012), suggesting that a toxic gain-of-function effect of α-synuclein may produce the neurodegeneration in PD and other synucleinopathies. At least in some instances, however, the physiological function of α-synuclein in promoting SNARE-complex assembly protects against neurodegeneration instead of promoting it (Chandra et al., 2005). Specifically, modest overexpression of α-synuclein rescues the lethal neurodegeneration caused by deletion of CSPα, a chaperone for the SNARE-protein SNAP-25 (Sharma et al., 2011b). α-Synuclein blocks neurodegeneration in CSPα KO mice by compensating for the decreased SNARE-complex assembly induced by the loss of SNAP-25 in these mice (Sharma et al., 2011a). Thus, the question arises whether α-synuclein performs independent physiological functions and pathological actions, or whether pathology induced by α-synuclein mutations or overexpression is related to a loss of its overall physiological function.

While the pathology caused by PD-linked α-synuclein mutants has been extensively compared to wild-type α-synuclein, few studies have performed systematic targeted mutagenesis experiments of α-synuclein to compare the consequences of various mutations for the neuropathogenic effects and physiological functions of α-synuclein. Here, we set out to fill this gap in our understanding, and to clarify whether pathology in synucleinopathies is caused by a loss- or a gain-of-function of α-synuclein. Towards this goal, we generated mutants of all sequence regions of human α-synuclein, and examined their properties using a variety of functional and pathological readouts. Our data suggest that the physiological function and neuropathogenic effects of α-synuclein are mediated by molecularly distinct processes.

MATERIALS and METHODS

α-Synuclein expression vectors

A c-myc epitope with a four amino acid linker was introduced into all expression vectors, resulting in the following N-terminal sequence: EQKLISEEDLGSGS. Introduction of stop codons or point mutations were accomplished by site-directed mutagenesis. All myc-tagged α-synuclein mutants were inserted into either pGEX-KG for bacterial expression (with an N-terminal TEV cleavage site right before the myc epitope tag, leaving an extra N-terminal glycine upon proteolytic removal of the GST moiety), pCMV5 for expression in HEK293T cells, FUW for lentiviral expression in neuronal culture, or L302 (containing an IRES-driven GFP reporter) for lentiviral expression in substantia nigra upon stereotactic injection.

Mice

Synuclein triple knockout mice, synaptobrevin-2 knockout mice, and wild-type mice maintained on C57BL/6 background were maintained and bred as described (Schoch et al., 2001; Burré et al., 2010; Xu et al., 2012). Mice of either sex were used for stereotactic injections or primary neuronal culture. The animal protocols used in this study, as well as the overall mouse husbandry practices were approved by the respective Institutional Animal Care and Use Committees at UT Southwestern Medical Center and Stanford University.

Primary mouse neuronal culture

Mouse hippocampal neurons were cultured from newborn mice essentially as described (Tang et al., 2006; Maximov et al., 2007). Brain regions were dissected in ice-cold Hank’s Balanced Salt Solution, dissociated by trypsinization (0.05% trypsin-EDTA, for10 min at 371C), triturated with a siliconized pipette, and plated (100 ml) onto a 12-mm coverslip (for immunofluorescence) or on 12-well plastic dishes, coated for at least 30 min with Matrigel (BD Biosciences). Plating medium (MEM (Gibco) supplemented with 5 g/l glucose, 0.2 g/l NaHCO3 (Sigma), 0.1 g/l transferrin (Calbiochem),0.25 g/l insulin (Sigma), 0.3 g/l L-glutamine (Gibco), and10% fetal bovine serum) was replaced with growth medium (MEM (Gibco) containing 5 g/l glucose, 0.2 g/l NaHCO3 (Sigma), 0.1 g/l transferrin (Calbiochem), 0.3 g/l L-glutamine (Gibco), 5% fetal bovine serum, 2% B-27 supplement (Gibco), and 2 mM cytosine arabinoside (Sigma)) 24–48 h after plating. Cultured neurons were transduced with recombinant lentiviruses and used for experiments as indicated.

Lentiviral vector production, transduction, and expression

Lentiviral vector (FUW containing myc-tagged α-synuclein mutants or empty vector), VSVG envelope glycoprotein and Δ8.9 HIV-1 packaging vectors were co-transfected in a 1:1:1 molar ratio into HEK293T cells (ATCC) in neuronal growth medium using Fugene-6 (Roche). Medium containing the viral particles was collected 48 h later and centrifuged for 10 min at 2000 rpm to remove any cellular debris. The supernatant containing virions was added to cultured neurons at 7 DIV, and the expression of the recombinant proteins was monitored at 21 DIV.

Expression of α-synuclein mutants in substantia nigra

Lentiviral vector L302 containing an IRES-driven GFP reporter (containing myc-tagged α-synuclein mutants), VSVG, REV, and RRE were co-transfected in a 1:1:1:1 molar ratio into HEK293T cells as described above. Viral particles were concentrated by centrifugation at 50,000 gav for 90 min. The viral pellet was re-suspended in neuronal medium (at 1/50 of the pre-centrifugation volume) containing 4 mg/ml polybrene (Sigma), snap-frozen in liquid N2 and kept at −80°C.

CD1 mice (P40–45; Charles River Labs) were anesthetized by intraperitoneal injection of 125–250 mg/kg Avertin (tribromoethanol, Sigma). 1 μl of viral solution was delivered through a glass pipette at a flow rate of 0.15 μl/min unilaterally (left hemisphere), at following coordinates: anteroposterior = 2.6–3.3 mm posterior to bregma (determined based on lambda-to-bregma distance of each mouse), lateral = 1 mm from midline suture, ventral = 4.2 mm below brain surface. Following 10 days of recovery, analysis of mouse behavior was commenced.

Behavioral studies

Beam walking test was used to record deficits in balance and limb control (accuracy and strength in limb placement). Animal’s ability to navigate across a beam to return to its home cage was tested using a wood end owel (cylindrical beam 60 cm in length, 1 cm in diameter). The beam was steadily fixed on both ends 40 cm above the ground leading to a small cage filled with bedding from the animal’s homecage. Mice were placed onto one side of the beam and were left to cross the beam to reach the cage. Mice that escaped into the cage were picked up and placed on the opposite side again for a total of 3 trials with 1 min inter-trial intervals. Number of footslips for each mouse per trial was scored. Forceplate actometry, a sensitive and quantitative method (Fowler et al., 2001) was used to document changes in locomotor activity induced by injection of lentivirus expressing α-synuclein mutants in substantia nigra. From the force plate traces/coordinate records, low mobility bouts and spatial confinement per 6 min session were determined.

Immunofluorescence and immunohistochemistry

Cultured neurons infected with lentiviruses encoding myc-tagged α-synuclein, or transfected HEK293T cells were washed three times with PBS, and fixed for 20 min at room temperature in PBS containing 4% paraformaldehyde. Following three washes with PBS, the fixed cultures were permeabilized for 5 min (PBS containing 0.1% Triton X-100 (Sigma). Cells were washed three times with PBS and blocked for 20 min with 5% BSA in PBS. α-Synuclein expression was monitored using myc antibodies (Santa Cruz, monoclonal, 1:200), synapsin antibodies (E028, polyclonal, 1:1000) were used as synaptic marker, incubated each overnight at 4°C in 1% BSA in PBS, and followed by anti-rabbit Alexa 633 and anti-mouse Alexa 488 secondary antibody (1:500 each), both incubated for 1 h each in blocking solution. The coverslips were rinsed six times with PBS, mounted on slides in Vectashield aqueous mounting medium (Vector Labs) and stored at 4°C. Laser scanning confocal microscopy was performed to compare localization, with serial excitation at 488 and 633 nm on a Leica TCSSP-2 inverted microscope.

For immunohistochemical studies, anaesthetized mice were perfused with ice-cold 4% paraformaldehyde in PBS, followed by removal of the brain and overnight fixation in 4% paraformaldehyde in PBS (room temperature). Fixed brains were cryopreserved in 30% sucrose in PBS for 2 days and frozen in Tissue Tek OCT embedding medium (Sakura Finetechnical). Sagittal brain sections (20 mm) were cut at −20°C (Leica CM3050S cryostat), picked up on slides and heat adhered at 37°C for 30 min. For immunostaining, slides were incubated in blocking solution (3% BSA, 0.1% Triton X-100 in PBS) for 1 h followed by overnight incubation with primary antibodies (4°C). Slides were washed three times in PBS (5 min each) and incubated in blocking buffer containing Alexa Fluor 488-, 546-, or 633-coupled secondary antibodies (Molecular Probes) for 3 h at room temperature. Following six washes in PBS, slides were mounted with Vectashield hard-set mounting medium with DAPI (Vector) followed by fluorescence microscopy.

All quantitations of immunofluorescence images were done with the image processing and analysis software Image J (NIH, Bethesda). Synaptic co-localization in neuronal culture was assessed using Pearson’s coefficient, with translated pictures used for background subtraction. In mouse brains injected with α-synuclein expressing lentiviruses, total TH-positive neurons, NeuN-positive puncta, and DAPI puncta were counted in each section containing GFP fluorescence. NeuN puncta were normalized to total DAPI puncta in the same brain section. Image acquisition and thresholding parameters were kept constant across each experiment.

Immunoblotting and immunoprecipitation

Either transfected HEK293T cells or cultured neurons were solubilized in PBS (pH 7.4) containing 0.15% Triton X-100 and protease inhibitors (Roche). Following centrifugation at 16,000 gav for 20 min at 4°C, the clarified lysate was used for immunoblotting (after addition of 2x SDS sample buffer containing 100 mM DTT) or subjected to immunoprecipitation. Immunoprecipitation was performed with the indicated primary antibodies and 50 μl of a 50% slurry of protein-A Sepharose beads (Amersham) for 2 h at 4°C. Control immunoprecipitations were performed with pre-immune sera. Following five washes with 1 ml of the extraction buffer, bound proteins were eluted with 2x SDS sample buffer containing 100 mM DTT and boiled for 20 min at 100°C. Co-precipitated proteins were separated by SDS-PAGE, with 5% of the input in the indicated lane.

To retain α-synuclein truncations on the nitrocellulose membranes, membranes were dried for 1h at room temperature and fixed for 15 min at room temperature in 0.2% glutaraldehyde in PBS. Membranes were washed 3x with TBS-T and treated as above.

Quantitation of SNARE-complexes as high molecular mass bands

Whole brains or cortices were homogenized in ice-cold PBS, and immediately dissolved in 2x SDS sample buffer. The lysates were subjected to SDS-PAGE and immunoblotting with antibodies to SNAP-25 (SMI81), and polyclonal antibodies to synaptobrevin-2 (P939) and syntaxin-1 (438B). To measure total SNARE protein levels, samples were boiled for 20 min at 100°C. SDS-resistant SNARE-complexes were defined as the immunoreactive material above 40 kDa that was absent from boiled samples (Hayashi et al., 1994).

Recombinant α-synuclein expression

All proteins were expressed in bacteria (BL21 strain) as GST fusion proteins in modified pGEX-KG vectors (GE Healthcare), essentially as described (Burré et al., 2010). Bacteria were grown to OD 0.5 (measured at 600 nm), and protein expression was induced with 0.05 mM IPTG for 6h at room temperature. Bacteria were harvested by centrifugation for 20 min at 4,000 rpm and 4°C, and pellets were resuspended in solubilization buffer (PBS, 0.5 mg/ml lysozyme, 1 mM PMSF, 1 mM EDTA, DNase, and an EDTA-free protease inhibitor cocktail (Roche)). Cells were broken by sonication (3× 15 pulses, 50% output), and insoluble material was removed by centrifugation for 20 min at 7,000 gav and 4°C. Proteins were affinity-purified using glutathione sepharose bead (GE Healthcare) incubation overnight at 4°C, followed by TEV cleavage (10 U/mg protein) for 4h at 22°C.

Lipid binding

Liposome preparation: Liposomes were always prepared on the day of usage, essentially as described (Burré et al., 2010). Either 500 μg brain L-α-phosphatidylcholine (PC, Avanti Polar Lipids) or 345 μg PC and 155 μg brain L-α-phosphatidylserine (PS, Avanti Polar Lipids) were mixed with 2.5 μg 1-oleoyl-2-{6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl}-sn-glycero-3-phosphocholine (NBD, Avanti Polar Lipids) in a glass tube. The lipid mixture was dried under a nitrogen stream and for 2h in a speed vac. To form unilamellar small vesicles, dried lipids were solubilized in 500 μl 20 mM phosphate buffer pH 7.4, vortexed for 2 min and then sonicated for 3× 15 pulses at 1 sec intervals and 38% sonicator output.

Lipid binding assay: 3.5 μM α-synuclein were incubated with liposomes at 1:363 molar ratio for 2h at room temperature. In a centrifugation tube, 100 μl protein sample was mixed with 100 μl 20 mM phosphate buffer pH 7.4 and 200 μl 80% Accudenz reagent (Accurate Chemical & Scientific corporation) in the same buffer (40% final density), and were carefully overlaid with 200μl 35% and 200 μl 30% Accudenz reagent, and 30 μl buffer. To achieve separation of bound and non-bound proteins, gradients were centrifuged for 3h at 280,000 gav and 100 μl fractions were collected from the top to the bottom of the gradient. The distribution of the liposomes in the gradient was determined by measuring the fluorescence of the lipid derivative NBD in each fraction using a fluorescence plate reader (Mithras LB 940, Berthold Technologies; excitation at 464 nm, emission at 531 nm). For analysis of protein distribution within the gradient, 10 μl 5x SDS sample buffer containing 100 mM DTT were mixed with 100 μl protein sample, and 20 μl were separated by SDS-PAGE and immunoblotted for SNARE proteins and α-synuclein.

Protein quantitation

All quantitative immunoblotting experiments were performed with iodinated secondary antibodies as described (Rosahl et al., 1995). Samples were separated by SDS–PAGE, and transferred onto nitrocellulose membranes. Blots were blocked in Tris-buffered saline containing 0.1% Tween-20 (Sigma) and 3% fat-free milk for 30 min at room temperature. The blocked membrane was incubated in blocking buffer containing primary antibody overnight at 4°C, followed by 3 washes in blocking buffer. The washed membrane was incubated in blocking buffer containing either horseradish peroxidase (HRP)-conjugated secondary antibody (MP Biomedicals, 1:8000) for 1 h at room temperature, or 125I-labelled secondary antibody (Perkin-Elmer,1:1000) overnight at room temperature. HRP immunoblots were developed using enhanced chemiluminescence (GE Healthcare). 125I blots were exposed to a phosphorimager screen (Amersham) overnight and scanned using a Typhoon scanner (GE Healthcare), followed by quantitation with ImageQuant software (GE Healthcare).

Antibodies

Monoclonal: β-actin (Sigma, A1978, 1:5,000), myc (Santa Cruz, 1:200), NeuN (MAB377, Millipore), SNAP-25 (SMI81, Sternberger Monoclonals, 1:5,000; cl. 71.1, SYSY, 1:1,000), synaptobrevin-2 (cl. 69.1, SYSY, 1:1,000), α-synuclein (BD Transduction; 1:1,000), syntaxin-1 (cl. HPC-1, SYSY, 1:1,000). Polyclonal: myc (Sigma, 1:1,000), synapsin (E028, 1:1,000), syntaxin-1 (438B, 1:1,000). TH (AB112, Abcam, 1:400).

Quantitation of functional and pathological indices

Average function was calculated from data obtained for lipid binding (% of WT), synaptic targeting (% of WT), synaptobrevin-2 binding (100% or 0%), and SNARE-complex assembly (% of WT). Average pathology was calculated from data obtained for aggregation in transfected HEK cells (% of WT), beamwalk analysis (% of WT), spatial confinement and low mobility bouts obtained by force plate analysis (% of WT), and analysis of dopaminergic and total neuron loss in substantia nigra (% of WT).

Statistical analyses

Unless stated otherwise, co-immunoprecipitation experiments are shown as recovered protein (relative to the input), which was first normalized to the immunoprecipitated protein and then normalized to the control. All other data shown are means ± SEMs, and were statistically analyzed by Student’s unpaired two-tailed t test to compare the data groups. “n” refers to the number of different cultures or mice used in each group in separate experiments, or to the number of independent in vitro experiments.

RESULTS

Design and generation of α-synuclein mutants

α-Synuclein is a small protein (140 residues) composed of an N-terminal region (residues 1–95 accounting for ~2/3 of the molecule) that binds to membranes in an α-helical conformation, and a C-terminal region (residues 96–140) that remains unstructured upon membrane-binding, but is phosphorylated and binds to synaptobrevin-2 (Fig. 1A; Okochi et al., 2000; Fujiwara et al., 2002; Kahle et al., 2002; Anderson et al., 2006; Beyer, 2006; McFarland et al., 2008; Paleologou et al., 2008; Waxman and Giasson, 2008; Burré et al., 2010; Paleologou et al., 2010). The N-terminal region of α-synuclein contains seven imperfect 11-amino acid repeats with a KTKEGV consensus sequence. Upon membrane binding, the N-terminal region forms either two α-helices that are connected by a flexible linker or a single extended α-helix with a possible continuous switch between the two states under physiological conditions (Chandra et al., 2003; Bisaglia et al., 2005; Ulmer et al., 2005; Borbat et al., 2006; Bortolus et al., 2008; Georgieva et al., 2010; Lokappa and Ulmer, 2011).

Figure 1
α-Synuclein mutagenesis strategy and experimental scheme

To investigate the relationship between the physiological function of α-synuclein in SNARE-complex assembly and its pathological effects in neurons, we systematically mutated key residues and sequences of human α-synuclein (Fig. 1A). We generated two types of α-synuclein mutations, (i) point mutations that introduce proline substitutions at defined positions in the 11-amino acid repeats to impair α-helix formation or to replicate Parkinson’s disease mutations; and (ii) deletion mutations that remove parts of the overall protein (Fig. 1A). In this manner, we produced a total of 26 different α-synuclein mutants. All α-synuclein mutants were tagged with a myc epitope to allow accurate quantitation of the levels of the various mutant proteins (Fig. 1B). We then examined all α-synuclein mutants by seven assays that range from experiments using purified proteins in in vitro systems to experiments in cultured neurons and in vivo in mice, thus producing 189 principal measured parameters (Fig. 1C). These seven assays include analyses of the biochemical activities of α-synuclein (phospholipid binding, aggregation, and synaptobrevin-binding), the targeting of α-synuclein and its mutants to synapses, analysis of their effects on SNARE-complex assembly, and examination of the in vivo effects of lentivirally expressed α-synuclein and its mutants (Fig. 1C).

Lipid binding of α-synuclein is mediated by its two α-helices

α-Synuclein cycles between a cytosolic monomeric state, and a membrane-bound state which localizes α-synuclein to synaptic vesicles (Iwai et al., 1995). Thus, any change in membrane affinity may alter its function. To measure the ability of point and truncation mutants of α-synuclein to bind to phospholipid vesicles, we used purified myc-tagged α-synuclein proteins (Fig. 1B) and a previously described liposome flotation assay (Figs. 2A and 2B; Burré et al., 2010). When α-synuclein was assayed without liposomes or in the presence of phosphatidylcholine liposomes (that are not negatively charged), no shift of α-synuclein to the liposome-containing top gradient fractions was detected (Fig. 2C). Upon introduction of 30% phosphatidylserine into phosphatidylcholine liposomes, the majority of wild-type (WT) α-synuclein was present in the liposome fraction, unaffected by presence or absence of the N-terminal myc epitope-tag (Fig. 2D). All proline mutations except G41P reduced the ability of α-synuclein to bind to phospholipid vesicles, presumably due to breaking of one of the two alpha-helices (Chandra et al., 2003; Bussell and Eliezer, 2004; Ulmer and Bax, 2005), whereas the two PD mutants A53T and E46K did not impair lipid binding (Fig. 2D). Deletion of the unstructured C-terminus of α-synuclein had no effect on lipid binding, but deleting one of the two α-helices abolished lipid binding (Fig. 2E). Notably, deletion of 7 N-terminal helical residues also dramatically reduced lipid binding (Fig. 2E). These liposome binding results were quantified in multiple independent experiments, allowing an accurate determination of the percentage of total protein that was bound (Fig. 2F). The quantitations revealed that although the proline mutations impair phospholipid binding, they do not block it in the same manner as complete deletions of one of the two α-helices, suggesting that the proline mutations only decrease the affinity of α-synuclein for phospholipids, but do not abolish binding. These quantitations also indicate that proline mutations in all parts of the two α-helices have similar detrimental effects (except for the G41P mutation which appeared to be innocuous), suggesting that both α-helices of α-synuclein and all segments of these α-helices are equally important for phospholipid binding.

Figure 2
Binding of α-synuclein mutants to phospholipid membranes

Effect of α-synuclein mutations on α-synuclein localization

α-Synuclein is highly enriched in presynaptic terminals (Iwai et al., 1995), consistent with its role in promoting SNARE-complex assembly (Burré et al., 2010). Immunolocalization studies in cultured neurons from wild-type mice revealed a high degree of α-synuclein co-localization with the synaptic vesicle marker synapsin, measured using Pearson’s coefficient (Figs. 3A and 3B). The α-synuclein co-localization was almost as high as that of the synaptic vesicle protein synaptobrevin-2. In cultured neurons from synaptobrevin-2 knockout (KO) mice (Schoch et al., 2001), the synaptic localization of α-synuclein was significantly decreased (Figs. 3A and 3B). However, no change in synaptobrevin-2 localization was detected in neurons from α/β/γ-synuclein triple KO mice. Together, these experiments not only support the preferential localization of α-synuclein in presynaptic terminals adjacent to synapsin, but also suggest that this localization depends, at least in part, on the binding of α-synuclein to synaptobrevin-2 (Burré et al., 2010), whereas the localization of synaptobrevin-2 is independent of synucleins.

Figure 3
Synaptic targeting of α-synuclein mutants

We then asked which α-synuclein sequences may be necessary for its presynaptic localization. To address this question, we expressed wild-type α-synuclein and the 26 α-synuclein mutants in cultured cortical mouse neurons using lentiviral infection at 7 days in vitro (DIV7), and analyzed the localization of the expressed α-synuclein proteins by immunocytochemistry at DIV21 (Figs. 3C and 3D). These experiments were independently repeated multiple times and quantified by image analysis in a blinded manner, allowing a numerical description of the degree of presynaptic targeting of each α-synuclein mutant (Fig. 3E).

Exogenous wild-type myc-tagged α-synuclein exhibited the same degree of co-localization with endogenous synapsin as endogenous α-synuclein (Figs. 3B and 3E). Surprisingly, most proline mutants which had a profound effect on lipid binding, at best produced a marginal decrease in the presynaptic targeting of α-synuclein. Only the PD mutant A30P showed a significant (~25%) reduction of presynaptic targeting (Figs. 3CE), suggesting an additional defect besides the impaired lipid binding. Most truncation mutants, however, significantly decreased the presynaptic localization of α-synuclein, demonstrating that both α-helices of α-synuclein are essential for α-synuclein targeting to synaptic vesicles (Figs. 3D and 3E).

The C-terminal α-synuclein sequence is essential for synaptobrevin-2 binding and for promoting SNARE-complex assembly

To test which α-synuclein mutations impair binding to synaptobrevin-2, we transfected HEK293T cells with expression vectors encoding synaptobrevin-2 and wild-type or mutant α-synucleins, and analyzed their interaction using co-immunoprecipitation with antibodies to the myc epitope-tag on the transfected α-synucleins (Fig. 4). No point mutation impaired the co-immunoprecipitation of synaptobrevin-2 with α-synuclein (Fig. 4A). In contrast but in agreement with previous studies (Burré et al., 2010), truncation mutants lacking the C-terminal α-synuclein region did not bind to synaptobrevin-2, whereas truncation mutants of the N-terminal region had no effect (Fig. 4B).

Figure 4
Synaptobrevin-2 binding by α-synuclein mutants

We next probed the ability of various α-synuclein mutants to promote SNARE-complex assembly similar to wild-type α-synuclein. To analyze the effect of α-synuclein mutants on SNARE-complex assembly, we expressed the various α-synuclein proteins by lentiviral delivery in cultured neurons obtained from triple α/β/γ-synuclein KO mice. We then assayed SNARE-complex assembly using two independent methods, immunoblotting of SDS-resistant SNARE-complexes on SDS-polyacrylamide gels (Hayashi et al., 1994), and co-immunoprecipitation of the SNARE proteins synaptobrevin-2 and SNAP-25 with antibodies to the SNARE-protein syntaxin-1 (Figs. 5 and and6).6). For both assays, we used quantitative immunoblotting with 125I-labeled secondary antibodies to measure the amounts of proteins.

Figure 5
Effects of α-synuclein mutants on SDS-resistant SNARE-complex assembly
Figure 6
Effects of α-synuclein mutants on SNARE-complex assembly measured by SNARE protein co-immunoprecipitation

Both assays produced the same results. Whereas wild-type α-synuclein potently promoted SNARE-complex assembly as reported previously (Burré et al., 2010), all truncation mutants except for the deletion of the N-terminal 8 α-synuclein residues were unable to promote SNARE-complex assembly (Figs. 5 and and6).6). Thus, both phospholipid binding mediated by the α-helices and synaptobrevin-2 binding mediated by the C-terminal α-synuclein sequence (Figs. 2 and and4)4) are required for the ability of α-synuclein to promote SNARE-complex assembly. In contrast to the truncation mutants, none of the point mutants of α-synuclein, including the PD mutants E46K and A53T, but except for the A30P mutation, impaired the ability of α-synuclein to promote SNARE-complex assembly. Thus, partial impairment of phospholipid binding – as observed with most of these point mutants (Fig. 2) – does not block the function of α-synuclein to promote SNARE-complex assembly. The A30P mutation is unique among the α-synuclein point mutants in that it did impair SNARE-complex assembly, possibly due to impaired synaptic targeting (Fig. 3).

Effect of α-synuclein mutations on α-synuclein aggregation

α-Synuclein aggregation into Lewy bodies is a hallmark of PD and other neurodegenerative disorders (Spillantini and Goedert, 2000; Masliah et al., 2001), raising the question how various α-synuclein mutations may alter α-synuclein aggregation. To assess the aggregation of wild-type and mutant α-synuclein in an intracellular environment, we over-expressed wild-type and mutant α-synucleins in HEK293T cells and measured amount of protein produced using quantitative immunoblotting (Figs. 7A and 7B) as well as the α-synuclein aggregates formed using immunostaining with antibodies to the myc epitope-tag on the expressed α-synuclein proteins (Figs. 7C-E).

Figure 7
Aggregation propensity of α-synuclein mutants

All α-synuclein proteins were expressed at similar levels except for the truncations that included only the single α-helix (Figs. 7A and 7B). As expected, all three PD mutations of α-synuclein increased the propensity of α-synuclein to aggregate (Figs. 7D and 7E). Moreover, consistent with previous results (Liu et al., 2005; Lewis et al., 2010), C-terminal truncations enhanced α-synuclein aggregation (Figs. 7C-E). However, most proline substitutions had no effect on α-synuclein aggregation except those falling in the so-called ‘non-amyloid β component’ (NAC) region, which is consistent with the known involvement of this region in synuclein aggregation (Giasson et al., 2001; Uversky and Fink, 2002; Du et al., 2003; Waxman et al., 2009).

Wild-type and mutant α-synucleins impair nigrostriatal function in vivo

Does either the loss-of-function or the aggregation propensity of various α-synuclein mutants correlate with their pathological potential in vivo? To monitor effects of α-synuclein mutants on dopaminergic cell survival and mouse motility, we stereotactically injected control lentivirus and lentiviruses expressing wild-type and mutant α-synucleins unilaterally into substantia nigra of wild-type mice. We performed these injections on postnatal day 40–45, and monitored motor coordination in the injected mice every 5 days, starting 10 days after injection.

Beamwalk analysis, used to measure balance and motor function, showed that mice injected with α-synuclein lentivirus performed worse than mice injected with control virus (Fig. 8), confirming previous studies that lentiviral overexpression of α-synuclein in the substantia nigra produces a dysfunction of motor performance (Lo Bianco et al., 2002; Lauwers et al., 2007; Alerte et al., 2008). A comparison of the effects of various α-synuclein mutants with wild-type α-synuclein revealed that exactly those mutations which increased α-synuclein aggregation (Fig. 7) also increased the motor impairment upon nigrostriatal expression (Fig. 8). The only difference between the aggregation and motor dysfunction effects of α-synuclein mutants was that the deletion of the N-terminal 8 residues did not decrease the increased aggregation of C-terminally truncated α-synuclein (Fig. 7), but did prevent the enhancement of motor dysfunction induced by the C-terminal truncation (Fig. 8).

Figure 8
Motor defects in mice injected with lentiviruses expressing α-synuclein mutants assayed using the beamwalk task

To assess motor function in mice overexpressing mutant α-synuclein in the substantia nigra with a second, independent assay, we monitored these mice on a forceplate actometer (Fowler et al., 2001; Fig. 9A). Quantitation of the movements of the mice on the forceplate revealed that wild-type α-synuclein overexpression significantly decreased the mobility of the mice as measured by the number of low mobility bouts and spatial confinement (Fig. 9B). Strikingly, the overall pattern of the effects of the mutations in α-synuclein on motor impairment was the same as observed with the beamwalk assay, although the forceplate assay was notably noisier (Fig. 9). Specifically, the PD-associated mutations, proline subsitutions in the NAC region, and C-terminal truncations again enhanced the deleterious effect of overexpressed α-synuclein, whereas other mutations had no significant effect on the impairment of motor function by α-synuclein. The only difference between the results with the two assays (beamwalk and forceplate) was that the latter also identified a phenotype associated with one of proline substitutions (A19P), and suggested a trend for several other mutations.

Figure 9
Forceplate analysis of motor function in mice injected with lentiviruses expressing α-synuclein mutants

α-Synuclein induced neuronal cell loss in the substantia nigra

Do motor impairments in mice with lentiviral expression of various mutants of α-synuclein correlate with neuron loss in the injected area? To assess and compare cell loss in the substantia nigra, we sacrificed mice 45 days after injection, and analyzed dopaminergic cell loss by quantitating tyrosine hydroxylase (TH)-positive neurons (Fig. 10). GFP produced by the viruses via an internal ribosomal entry site was used to identify the injection site. Compared to control injections, brains expressing wild-type or mutant α-synucleins exhibited an overall reduction in the number of dopaminergic neurons in the injected area, suggesting that simple overexpression of α-synuclein causes death of dopaminergic neurons (Figs. 10A and 10B). Note that the reduced GFP fluorescence in some brain slices may be caused by loss of infected neurons. We found that α-synuclein with PD-associated mutations caused an even larger loss of dopaminergic neurons than wild-type α-synuclein. In addition, the same proline mutations that produced increased α-synuclein aggregation (Fig. 7) and impaired motor function in the injected mice (Figs. 8 and and9)9) also enhanced the neurotoxic effect of α-synuclein (Fig. 10). The effect of truncated α-synuclein, however, was less strong, although even here significant enhancement of the deleterious effects of α-synuclein was observed.

Figure 10
Neuron loss in mice expressing α-synuclein mutants in substantia nigra

Next, we asked whether α-synuclein over-expression also affects non-dopaminergic neurons in the injected area. Since neurons labeled by NeuN antibodies do not coincide with TH expressing neurons (Cannon and Greenamyre, 2009), we assessed loss of NeuN-positive neurons by measuring the NeuN/DAPI ratio (Fig. 10; Sharma et al., 2011b). We found a reduction of neuron number for all α-synuclein mutants compared to control injections. Compared to WT injections, we observed a significant reduction of neuron number for PD and NAC-domain mutants and the C-terminally truncated α-synuclein1–95. These data suggest, that over-expression of α-synuclein does not specifically induce cell death in dopaminergic neurons, but is toxic to neurons in general.

Overall, all pathological readouts correlate well with each other in that PD-associated mutations and point mutations in the ‘NAC’ region as well as C-terminal truncations of α-synuclein increase α-synuclein aggregation and pathology compared to wild-type α-synuclein. Our data further demonstrate that loss-of-function of α-synuclein does not correlate with increased aggregation propensity/pathology, suggesting that pathology in synucleinopathies may arise from gain-of-toxic function of α-synuclein.

DISCUSSION

Although extensive evidence established a central role for α-synuclein in the pathogenesis of neurodegenerative disorders, especially PD (Spillantini and Goedert, 2000; Masliah et al., 2001; Dawson and Dawson, 2003), questions remain regarding the structure/function relations of α-synuclein in neurodegeneration. Moreover, previous studies examining how α-synuclein acts in neurodegeneration have rarely correlated in vitro and in vivo activities of α-synuclein mutants, and no systematic mutagenesis of the entire molecule was previously performed. Perhaps the most important issue, however, is whether the neuropathological effects of α-synuclein are related to its physiological functions. This issue is particularly pertinent because the physiological role of α-synuclein as a SNARE chaperone can at least in the context of the neurodegeneration caused by deletion of CSPα act to prevent neurodegeneration (Chandra et al., 2005), thus raising the question whether the neuropathological effects of α-synuclein are possibly caused by a loss of α-synuclein function. The present study attempts to address these issues in a systematic analysis of 26 α-synuclein mutants using 7 assays for its physiological and pathological actions. The relative effects of the α-synuclein mutations on the physiological and pathological actions of α-synuclein are summarized in Fig. 11. Overall, our data suggest the following conclusions:

Figure 11
Summary of function and pathology for α-synuclein mutants
  1. There is no correlation between the sequences required for physiological and pathological actions of α-synuclein. Functionally inactive α-synuclein does not necessarily produce pathology, whereas many mutations that enhance the neuropathogenic effect of α-synuclein do not detectably alter its physiological function as a SNARE chaperone.
  2. Full phospholipid binding is not required for the physiological function of α-synuclein as a SNARE chaperone at the synapse. Only when >75% of lipid binding is reduced (e.g. helix 1 is missing), is the physiological function of α-synuclein impaired.
  3. Synaptic targeting of α-synuclein is mediated by both lipid and synaptobrevin-2 binding.
  4. The two α-helices and the C-terminal region of α-synuclein are both essential for its function as a SNARE-chaperone, the former probably because they bind to phospholipids and the latter probably because it binds to synaptobrevin-2.
  5. The propensity of α-synuclein to aggregate and the pathogenic effects of α-synuclein are independent of its N-terminal α-helix, but enhanced by point mutations in its central α-helix and by deletion of its C-terminal region. Thus, the aggregation and neurotoxicity of α-synuclein correlate precisely, suggesting that the two processes are related.

Overall, our data support the notion that aggregation of α-synuclein is a central element of its neurotoxicity, extending previous studies showing that PD mutants of α-synuclein increase the propensity of protofibril or fibril formation of α-synuclein (Narhi et al., 1999; Conway et al., 2000; Greenbaum et al., 2005; Ono et al., 2011) and thus pathology in familial PD cases (Polymeropoulos et al., 1997; Kruger et al., 1998; Zarranz et al., 2004). Mechanistically, α-synuclein aggregation could cause neurotoxicity at least via two pathways: Aggregation of α-synuclein could be in itself neurotoxic. The resulting aggregates could damage neurons either as oligomers or as inclusion bodies, thereby impairing neuronal viability (Spillantini and Goedert, 2000; Masliah et al., 2001; Kayed et al., 2003; Volles and Lansbury, 2003; Lindersson et al., 2004; Lansbury and Lashuel, 2006; Tsika et al., 2010; Colla et al., 2012). Alternatively, α-synuclein aggregates could serve to nucleate aggregation of all cellular synucleins, thereby depleting the neuron of synucleins and causing an indirect loss-of-function effect. Although we cannot at present completely rule out the second hypothesis, several lines of evidence argue against it. Possibly the most compelling argument against the indirect loss-of-function hypothesis is the observation that a complete loss of all synucleins in triple α/β/γ-synuclein KO mice does not lead to the same major neuropathology as the overexpression of mutant α-synuclein. Specifically, synuclein-deficient mice display motor impairments only after 200 days and have no neuronal loss (Burré et al., 2010), suggesting that the loss-of-function phenotype of α-synuclein is a process different from the observed neuropathology.

In agreement with the PD pathology observed in humans (Singleton et al., 2003; Ibanez et al., 2004), we found that increased expression of wild-type α-synuclein is sufficient to cause pathology. All three PD mutants aggravated the α-synuclein pathology in our assays. We found that the pathology induced by lentiviral expression of α-synuclein in the substantia nigra manifests not only as the loss of dopaminergic neurons but also as a loss of other neurons in the injected area. While loss of dopaminergic neurons in the substantia nigra is responsible for motor symptoms in Parkinson’s disease (Fahn and Sulzer, 2004), a variety of other neurons in the central and peripheral nervous system also exhibit signs of pathology in postmortem analysis, suggesting that pathology induced by α-synuclein is not limited to dopaminergic neurons (Braak et al., 2007; Baba et al., 2012; Sulzer and Surmeier, 2012).

Surprisingly, we found that mutations in the NAC region cause pathology similar to familial PD mutations. ‘NAC’ was originally isolated as a component of β-amyloid from brain tissue of Alzheimer’s patients (Ueda et al., 1993), and was later identified as a fragment of α-synuclein (Nakajo et al., 1993; Jakes et al., 1994). The NAC region comprises residues 61–95 of α-synuclein, a region which is thought to drive fibril formation (Giasson et al., 2001; Uversky and Fink, 2002; Du et al., 2003; Waxman et al., 2009). Indeed, all of our mutants falling into this region (G67P, V70P, A78P, T81P, A89P) consistently increased the aggregation and pathogenicity of α-synuclein. Introduction of proline residues into the already aggregation-prone NAC region is predicted to reduce its helical content, and may thus shift the conformational equilibrium of α-synuclein further towards β-sheet content and subsequently towards more fibril formation, resulting in increased pathology. It is interesting that the N-terminal α-helix of α-synuclein (Chandra et al., 2005) binds more tightly to lipids than the C-terminal α-helix (which coincides with the NAC region) (Drescher et al., 2008), which may account for the greater aggregation propensity of the NAC region. Overall, our data suggest that if familial mutations within the NAC domain were to be found in PD patients, they would be similarly or more severe as those reported in known familial mutations.

C-terminally truncated α-synuclein is enriched in Lewy body extracts (Baba et al., 1998; Tofaris et al., 2003). Abnormal neurites containing C-terminally truncated α-synuclein are present in Alzheimer’s disease without conventional Lewy body pathology (Lewis et al., 2010), and transgenic mice expressing C-terminally truncated forms of human α-synuclein develop PD-like symptoms and exhibit PD-like neuronal pathology (Tofaris et al., 2006; Wakamatsu et al., 2008). Furthermore, co-expression of C-terminal truncated α-synuclein enhances full length α-synuclein induced pathology (Liu et al., 2005; Ulusoy et al., 2010). The charge in the unstructured C-terminus exerts a profound effect on the aggregation rate of α-synuclein (Hoyer et al., 2004; Levitan et al., 2011), and it has been suggested that interaction between the C-terminal and N-terminal or central NAC region are important in maintaining the natively unfolded structure of α-synuclein and prevent α-synuclein from changing conformation (Hong et al., 2011). Thus, lack of the C-terminus may increase aggregation rate and account for the observed pathology in our assays.

In our assays, all proline mutants we analyzed bind to synaptobrevin-2 and reveal similar impairment in lipid binding. However, only the A30P mutation demonstrated decreased synaptic targeting and thus less SNARE-complex stabilization. This result is puzzling, since mutation of the corresponding residues within other 11mer repeats did not result in a similar loss-of-function or aggregation of α-synuclein. What makes A30P so special? We cannot exclude the possibility that under physiological conditions, lipid binding may be affected by other factors. Alanine30 may be crucial for proper helix formation of α-synuclein. For A30P, loss of helicity up- and downstream of the substitution has been suggested (Bussell and Eliezer, 2004; Ulmer and Bax, 2005), and disruption of helicity in A30P may be more severe than for other proline mutants. Whatever mechanism turns out to be responsible for the increased cytosolic localization of A30P, this increased cytosolic localization compared to other proline mutations within the first α-helix of α-synuclein may account for the increased pathology: shifting of α-synuclein structure towards a more unfolded state may allow increased beta-sheet formation, aggregation and thus increased pathology.

Truncation of 8 N-terminal residues of α-synuclein resulted in a 60% decrease in lipid binding. Presumably, deletion of a helical stretch affects helical structure more severely than introduction of single proline residues within the helix, which disturb protein folding only locally, as shown for A30P (Bussell and Eliezer, 2004; Ulmer and Bax, 2005). Similarly, most proline mutants resulted in comparable impairments in lipid binding except for G41P, which did not affect binding of α-synuclein to liposomes. Likely, introduction of a helix-breaking proline residue at the very end of helix 1 has less impact on helicity than proline residues in the middle of a helical domain.

In summary, our data suggest that α-synuclein pathology is not caused by loss-of-function of α-synuclein, but by gain-of-toxic function. Since α-synuclein knockout mice do not show significant neuropathology (Abeliovich et al., 2000; Schluter et al., 2003), and removal of all three synuclein genes is required for severe pathology (Burré et al., 2010; Greten-Harrison et al., 2010; Anwar et al., 2011), suppression of α-synuclein expression alone may not be pathogenic in humans. Therapies to reduce or abolish α-synuclein expression may thus be a worthwhile strategy to decrease/prevent α-synuclein induced pathology.

Acknowledgments

We thank Dr. Xiaofei Yang for breeding of synaptobrevin-2 knockout mice, Dr. Stephan Lammel for assistance with lentiviral injections, and Christian Burré (CircumFlex Computer Systems) for help with data analysis. This work was supported by a grant from NINDS (NS077906 to T.C.S.) and postdoctoral fellowships from the German Academy of Sciences Leopoldina (BMBF-LPD 9901/8-161 to J.B), and from the Human Frontiers Program (LT00527/2006-L to M.S.).

Footnotes

The authors declare no competing financial interests.

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