Rab7 Reduces α-Synuclein Toxicity in Rats and Primary Neurons

doi:10.21203/rs.3.rs-277472/v1

Download PDF

Research article

Rab7 Reduces α-Synuclein Toxicity in Rats and Primary Neurons

https://doi.org/10.21203/rs.3.rs-277472/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 01 Jan, 2022

Read the published version in Experimental Neurology →

Version 1

posted

You are reading this latest preprint version

Background

During the pathogenesis of Parkinson’s disease (PD), aggregation of alpha-synuclein (αSyn) induces a vicious cycle of cellular impairments that lead to neurodegeneration. Consequently, removing toxic αSyn aggregates constitutes a plausible strategy against PD. In this work, we tested whether stimulating the autolysosomal degradation of αSyn aggregates through the Ras-related in brain 7 (Rab7) pathway can reverse αSyn-induced cellular impairment and prevent neurodegeneration in vivo.

Methods

The disease-related A53T mutant of αSyn was expressed in primary neurons and in dopaminergic neurons of the rat brain simultaneously with wild type (WT) Rab7 or its dominant-negative T22N mutant as a control. The cellular integrity was quantified by morphological and biochemical analyses.

Results

In primary neurons, WT Rab7 rescued the αSyn -induced loss of neurons and neurites. Furthermore, Rab7 decreased the amount of reactive oxygen species and the amount of Triton X-100 insoluble αSyn. In rat brain, WT Rab7 reduced αSyn -induced loss of dopaminergic axon terminals in the striatum and the loss of dopaminergic dendrites in the substantia nigra pars reticulata. Further, WT Rab7 lowered αSyn pathology as quantified by phosphorylated αSyn staining. Finally, WT Rab7 attenuated αSyn-induced DNA damage in primary neurons and rat brain.

Conclusion

Rab7 reduced αSyn-induced pathology, ameliorated αSyn-induced neuronal degeneration, oxidative stress and DNA damage. These findings indicate that Rab7 is able to disrupt the vicious cycle of cellular impairment, αSyn pathology and neurodegeneration present in PD. Stimulation of Rab7 and the autolysosomal degradation pathway could therefore constitute a beneficial strategy for PD.

Neurology

alpha-synuclein

Rab7

protein aggregation

Parkinson’s disease

DNA damage

oxidative stress

autophagy

neuroprotection

Aggregation and accumulation of alpha-synuclein (αSyn) are key events in Parkinson`s disease (PD) and other synucleinopathies. αSyn monomers are mainly located at presynaptic vesicles whereas misfolded αSyn oligomers and high molecular weight aggregates can be found throughout the entire neuron. αSyn pathology affects diverse cellular functions, including: increasing reactive oxygen species (ROS), impairing mitochondrial function [1,2], reducing proteasomal degradation and autophagy [3,4] as well as inducing DNA damage [5]. However, the pathogenesis of PD does not follow a lineair succession of events and is rather charactersiezed by self-enhancing mechanisms. ROS and mitochondrial dysfunction also increase αSyn pathology [6,7], as does impaired protein quality control and clearance [3,8–11]. Moreover, autophagy can protect against oxidative stress [12–14] while oxidative stress can impair autophagy [15,16]. DNA damage can also result from ROS imbalance [17,18], and it can induce autophagy [19]. Taken together, toxic αSyn species initiate a vicious circle of protein aggregation and cellular dysfunction as summarized in the graphical abstract. Removing toxic αSyn species has the potential to disrupt this toxic feedback loop, improve neuronal health and slow disease progression in synucleinopathies.

Aggregated αSyn is degraded via the autophago-lysosomal pathway. Mutations in genes encoding autophago-lysosomal pathway proteins, such as glucocerebrosidase (GBA), ATPase cation transporting 13A2 (ATP13A2), Ras-related in brain 39B (Rab39B), ADP-ribosylation factor-like protein 8B (Arl8b), lysine acetyltransferase 8 (KAT8), Ras-related in brain guanine nucleotide exchange factor 1 (RabGEF1) or vacuolar protein sorting ortholog 35 (VPS35), cause impairment of degradation, and are risk factors for PD [20–24]. The multi-step process of autophagy is regulated by several critical proteins, including the Ras-related in brain (Rab) family of small GTPases. Ras-related in brain 7 (Rab7) orchestrates trafficking of autophagosomes and late endosomes, catalyzes the fusion of autophagosomes with lysosomes, and regulates late maturation of autophagosomes. Rab7 malfunction hampers autolysosome formation, leads to the accumulation of dysfunctional autophagosomes and consequently impairs protein degradation [23,25–29]. We previously demonstrated that Rab7 is the rate-limiting factor for αSyn aggregate clearance; its overexpression facilitates the removal of αSyn inclusions in human cells and alleviates αSyn-induced motor phenotype in fruit flies [30].

In the present work, we investigated whether mammalian neurons can be protected against αSyn toxicity by Rab7-induced stimulation of aggregate clearance. Expression of αSyn carrying the disease-related A53T mutation was achieved in mouse primary neurons and in dopaminergic neurons of rat substantia nigra (SN) by a recombinant adeno-associated viral vector 2/7(rAAV) – as previously described [31–33]. Wild-type (WT) Rab7 was also expressed using a rAAV2/7 vector. As a negative control, we used the dominant negative (DN) Rab7 mutant T22N, which binds Rab7 effectors but not guanosine-5'-triphosphate (GTP) [34]. First, we assessed the effect of Rab7 on αSyn pathology, αSyn-induced DNA damage, oxidative stress and cell death in mouse primary neurons. Second, we analyzed the integrity of the dopaminergic nigrostriatal neurons in vivo. Finally, we investigated αSyn accumulation and DNA damage in the dopaminergic neurons. We demonstrate that inducing Rab7-mediated clearance of αSyn aggregates in neurons improves cellular integrity and reduces neurodegeneration.

Source of chemicals, antibodies, composition of buffers, equipment and software used in this study are listed in supplemental Table S1.

Recombinant AAV production and purification

Vector production and purification was carried out as previously described [33]. The plasmids include: the constructs for the AAV7 serotype, the AAV transfer plasmid encoding either the human A53T mutant αSyn, the enhanced green fluorescent protein (GFP), human WT Rab7 with a double N-terminal HA-tag or T22N DN Rab7 with a double N-terminal HA-tag, under the control of the CMVie enhanced synapsin1 promoter and the pAdvDeltaF6 adenoviral helper plasmid. Real-time PCR analysis was used for genomic copy (GC) determination.

Primary neuronal culture

Primary cortical cultures were prepared from P1-3 C57BL6/J mouse pups of mixed sex as previously described [1]. Briefly, after dissection and trypsination, dissociated neurons were plated onto poly-L-ornithine coated glass coverslip (100 000 cells per well in 24-well dishes), and maintained in Neurobasal A medium containing 2 % B27, 0.5 mM glutamax and antibiotics. One third of the medium was changed on every third day, and from the second medium change on, no antibiotics were added. On DIV 12, neurons were transduced with one of the following combinations of rAAV2/7 vectors: (1) GFP, (2) GFP with DN Rab7A, (3) GFP with WT Rab7A, (4) A53T αSyn with GFP, (5) A53T αSyn with DN Rab7A or (6) A53T αSyn with WT Rab7A (final concentration for all 1.0e+9 GC/ml). Cells were fixed (4 % paraformaldehyde, 10 min, 20 °C) on DIV 15 for phospho-αSyn and LAMP1 labeling. Cells were fixed on DIV 16 for detecting 53BP1. Cells were fixed or lysed on DIV 18 or DIV 20 for toxicity and comet assays.

Microfluidic chambers for axonal regeneration assay

The effect of WT and DN Rab7 on αSyn-induced inhibition of axonal regeneration was measured as previously described [35]. Primary cortical neurons were seeded in silicone microfluidic devices with 450 µm groove length according to the manufacturer`s protocol. After bonding the microfluidic devices to a glass coverslip pre-coated with poly-L-ornithine, 100 000 cells (5.0e+6/ml) were plated in the cell compartment. The axonal side of the chamber was filled with 200 µl complete Neurobasal medium. After attachment of the cells, 200 µl medium was added in the cell compartment as well. One third of the medium was changed every third day on both sides. On DIV 12, neurons were infected with rAAV (final concentration for all 5e+8 GC/ml). On DIV 15, axons were mechanically cut by an air bubble. Cells were fixed on DIV 20 and stained for tubulin (PRB-435P, followed by Alexa 555-labeled anti-mouse secondary antibody). Axonal length in the distal compartment was measured using ImageJ by drawing a segmented line along each individual axon projecting out of the microgroove.

Animals and surgery

All animal experiments were carried out in accordance with the European Communities Council Directive of November 24, 1986 (86/609/EEC) and approved by the Bioethical Committee of the KU Leuven (ECD project P067-2013 and P085-2014, Belgium). Eight-week-old female Wistar rats (Janvier, France, 200-250 g) were housed under a 12-hour light and/or dark cycle with free access to pelleted food and tap water. All surgical procedures were performed using aseptic techniques. Rats were anaesthetized with ketamine (60 mg/kg, intraperitoneal (i.p.), Ketalar, Pfizer, Belgium) and medetomidine (0.4 mg/kg, i.p., Dormitor, Pfizer). 3 µl of vector mix containing the rAAV2/7 A53T αSyn vector (9.0E11 GC/ml, injected genome copies 1,35E9) and rAAV2/7 HA-Rab7A T22N vector (9,3E11 GC/ml, injected genome copies 1,39E9) or the rAAV2/7 HA-Rab7A WT vector (9,3E11 GC/ml, injected genome copies 1,39E9) was injected in the substantia nigra (AP: -5.3; L: -2.0; DV: -7.2 from dura, using Bregma as reference) with a Hamilton syringe (Hamilton, Bonaduz, GR, Switzerland) as previously described [32,36].

Forelimb use was quantified by the cylinder test four weeks after the unilateral rAAV-injection. Contacts made by each forepaw with the wall of a 20-cm-wide clear glass cylinder were scored from the videotapes by an observer blinded to the animal’s identity. A total of 25 contacts were recorded for each animal, and the impaired (left) forelimb usage was expressed as a percentage of total forelimb contacts. Non-lesioned control rats score around 50 % in this test.

Rats were sacrificed four weeks after the rAAV-injection with an overdose of sodium pentobarbital (200 mg/kg, i.p.) followed by intracardial perfusion with 4 % paraformaldehyde in PBS. After post-fixation (4 % paraformaldehyde, overnight), 50 µm-thick coronal brain sections were cut with a vibrating microtome.

Immunohistochemical and immunofluorescent stainings

To quantify the number of dopaminergic neurons in the substantia nigra (SN) and the density of dopaminergic axon terminals (fibers) in the striatum, free-floating sections were stained for tyrosine hydroxylase (TH, Ab152). αSyn pathology was quantified after staining for αSyn phosphorylated at Serine 129 (phospho-αSyn, 11A5). Sections were pretreated with 3 % hydrogen peroxide (10 min) and incubated with primary antibody in 10 % normal rabbit serum overnight. Incubation with biotinylated anti-rabbit IgG (TH) and anti-mouse IgG (phospho-αSyn) secondary antibodies was followed by streptavidin - horseradish peroxidase complex. TH immunoreactivity was visualized using Vector SG as a chromogen and phospho-αSyn immunoreactivity was visualized using 3,3-diaminobenzidine (DAB).

To determine neuroinflammation, every sixth striatal section was stained for the astroglia marker Glial fibrillary acidic protein (GFAP, ab4674) and for the microglia marker Iba1 (019-19741) by incubation with primary antibodies at 4 °C overnight and fluorescently labelled secondary antibodies (Alexa 488 conjugated goat anti-chicken and Alexa 555 conjugated donkey anti-rabbit, 120 min at 20 °C). Sections were counterstained with Hoechst and mounted with Fluoromount-G.

To detect DNA damage, brain slices from the SN (minimum 3 sections per animal) were stained for 53BP1 (PA5-54565), human αSyn (ALX-804-258-L001) and the HA-tag of Rab7 (MMS-101P). After incubation with secondary antibodies (Alexa 488 conjugated anti-rat, Alexa 555 conjugated anti-rabbit, and Alexa 405 conjugated anti-mouse), sections were also mounted with Fluoromount-G.

Fixed primary neurons were permeabilized using 0.2 % Triton X-100 in Tris-buffered saline (TBS, pH 7.4, 10 min). Non-specific sites were blocked by 2 % bovine serum albumin in TBS (RT, 1 hr). The following primary antibodies were applied in blocking solution (4 °C, overnight): Microtubule-associated protein 2 (MAP2, ab5392), Lysosomal-associated membrane protein 1 (LAMP1, ab208943), 53BP1, human αSyn and HA-tag (as above). Secondary antibodies were: Alexa 647 conjugated anti-chicken, Alexa 555 conjugated anti-rabbit, Alexa 488 conjugated anti-rat and Alexa 405 conjugate anti-mouse. Coverslips were mounted with Fluoromount G. Co-occurance of phospho-αSyn and LAMP1 signal was determined by using the Manders` coefficient (ImageJ), as fraction of phospho-αSyn signal within LAMP1 signal as previously [1,37].

Analysis of dopaminergic neurons, αSyn pathology and gliosis

The number of cells in the SN positive for TH and phospho-αSyn was determined by stereological measurements using the Optical fractionator method in a computerized system as described before [38] (StereoInvestigator). Every fifth section throughout the entire SN was analyzed, 7 sections for each animal. The coefficient of error calculated according to the procedure of Schmitz and Hof [39], varied between 0.05 and 0.10. The volume of striatal lesion was determined by quantifying the TH-positive terminals using the Cavalieri method (Stereologer). Every fifth section covering the entire extent of the striatum was included in the counting procedure, 7 sections for each animal. An investigator blinded to the different groups performed all the analyses.

For quantification of gliosis, fluorescent images were acquired from slices stained for GFAP and Iba1 using a 20x objective (NA 0.8) with an Axio Imager 2 microscope (Zeiss). After thresholding individual images (separately for GFAP and Iba1 channels), the area fraction was determined by ImageJ from 3-5 sections per animal and from 10 images per section as described previously [40].

For quantification of striatal phospho-αSyn pathology, images from 3-diaminobenzidine stained sections (minimum 4 sections per animal, and minimum 4 images per section) were acquired using a 20x objective (NA 0.8) with an Axio Imager 2 microscope (Zeiss). 4 independent blinded investigators rated the pattern of phospho-αSyn signal in 40-60 randomly selected images per evaluator. Blinding was achieved by 5 digit codes. Images were rated with 0 when mostly longer segments were present, and with 1 when a punctate pattern or dotted lines dominated the image. The results were analyzed in a hierarchical nested design as previously [41]. The evaluator was set as a random factor in the generalized linear mixed model, and it had no effect on the result. From the same images, the amount of phospho-αSyn staining was quantified as area fraction of the DAB signal as described for GFAP and Iba1.

Triton X-100 solubility

For αSyn protein quantiﬁcation and detection of Triton X-100 insoluble proteins, neurons were lysed in buffer containing 1 % Triton X-100, 25 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA and protease inhibitors as previously described [1]. After centrifugation (14000 g, 30 min, 4 °C), supernatant was used as Triton X-100 soluble fraction. The pellet (Triton X-100 insoluble fraction) was washed in ice-cold PBS, centrifuged again and re-dissolved with sonication (10 s) in 50 µl buffer containing 2 % SDS, 75 mM Tris, 15 % glycerin, 3.75 mM EDTA pH 7.4 and protease inhibitors. 10 µg of Triton X-100 soluble lysate or 10 µl from the re-dissolved pellet were loaded onto a 4-20 % Tris/glycine SDS gel for western blot analysis. After blocking, membranes were incubated first in the presence of antibodies against human αSyn (ALX-804-258-L001), HA tag (MMS-101P), PSD-95 (3450) and alpha-tubulin (ab6046), then with horse radish peroxidase-conjugated secondary antibodies (donkey anti-rat, donkey anti-mouse and donkey anti-rabbit). Signal was visualized with chemiluminescent substrate and detected with Luminescent Image Analyzer.

Biochemical assays

To determine the toxic effect of αSyn in cultured neurons, the concentration of LDH was measured in the medium 24 h after the last medium change using the Cytotoxicity Detection Kit (Roche) according to the manufacturer’s protocol. Briefly, for each independent preparations, 3-4 technical replicates were used and averaged. Absorbances were measured at 492 nm, reference wavelength was 620 nm. Background (medium) absorbance was substracted from all values, and values were normalized to control (non-treated) and to maximal lysed (treated with 2 % Triton X-100, according to the protocol) cells. Caspase activity was measured with EnzChek Caspase-3 Assay Kit (Thermo Fisher). Apoptosis was induced with 100 µM NMDA for 24 h. Cells were lysed and protein concentration determined (BCA assay). 100 µM Z-DEVD–AMC substrate was added to a sample of 100 µg protein. After 45 min incubation at 20 °C, fluorescence of cleaved Z-DEVD–AMC was acquired (Excitation/Emission: 342/441 nm; Inﬁnite 200 Pro). MAP2 stained primary neurons were quantified as area fraction from 20x images as described for GFAP and Iba1.

Mitochondrial membrane potential, cytoplasmic reactive oxygen species (ROS) production and mitochondrial ROS production were measured in separate assays as described previously [1]. Tetraethylbenzimidazolylcarbocyanine iodide (JC-1, 50 nM, 37 °C, 30 min) was used to measure the mitochondrial membrane potential. Fluorescence at 485/505 nm (excitation/emission) and 535/570 nm was detected by a plate reader (Inﬁnite 200 Pro). After measuring the basal mitochondrial membrane potential, the mitochondrial uncoupler FCCP (1 mM) was used to induce a mitochondrial stress.

Cytoplasmic ROS was measured using dichlorodihydroﬂuorescein diacetate (DCFH-DA; 10 mM, 37 C, 30 min), mitochondrial ROS was measured using MitoSOX (200 nM, 20 min, 37C). Fluorescence at 485/530 nm (DCFH-DA) or 510/580 nm (MitoSox) was measured with plate reader. To detect the response to stress-induced ROS production, preparations were treated with 1 mM FCCP (MitoSox) or with 1 % H₂O₂ (DCFH-DA).

DNA damage

To determine DNA double strand breaks, the number of 53BP1-positive nuclear loci was counted in brain sections containing nigral neurons transduced with human αSyn (minimum 3 sections per animal, and minimum 3 images per section, 15-30 neurons per animal) or in primary neuronal cultures (3 repetitions, minimum 15 neurons per repetition /condition) as previously described [18].

DNA single- and double-strand breaks were detected via single cell gel electrophoresis (comet) assay, according to the guidelines provided in [42]. Adherent microscopy slides were coated with 1 % low melting agarose (PeqLab). Primary neurons were harvested in PBS, mixed with 1 % agarose in PBS and placed on the pre-coated slide. After agarose gelling, slides were submerged in lysis buffer (2.5 M NaCl, 100 mM Na₂EDTA, 0.1 % Triton X-100, 10 mM TRIS, pH 10, 4 °C, 1 h) and denatured in alkaline electrophoresis buffer (0.3 M NaOH, 1 mM Na₂EDTA, pH>13, 30 min). Electrophoresis was carried out for 25 min at 20 V in the same buffer. Slides were removed from the electrophoresis chamber, fixed in 70 % ethanol and stained with 2.5 μg/ml of propidium iodide for 20 min at 20 °C. Excess of stain was removed by rinsing the slides with distilled water. For each slide, at least 10 images were acquired using an epifluorescence microscope (Zeiss Axio Observer). For each comet, the “tail moment” was determined as the product of the percentage of DNA in the tail and the tail length using the Open Comet plugin for Image J as described previously [17].

Statistical analysis

For in vitro experiments, “n” was set to the number of individual preparations. Within one experiment, mean of the technical replicates was determined for each treatment group. In animal experiments, “n” was set to the number of animals. No animals were excluded from the analysis. In graphs, markers represent individual preparations or individual animals, lines represent mean and standard deviation (SD) of all individual preparations or all animals. Data normality was tested by the Shapiro-Wilk test and graphically by QQ plot (R). t-test, one-way ANOVA or two-way ANOVA were performed using GraphPad Prism. Linear regression was performed using R (v 2.8.0). Binomial data from pattern rating was analyzed using generalized linear mixed-effects model fitted with binomial distribution. Evaluator, animal, section and image were used as random effects nested within each other (R package: lme4). P values are indicated in the graphs by symbols with * or # representing p<0.05, ** or ## representing p<0.01, *** or ### representing p<0.001. Exact p values are given in the Figure legend.

Rab7 decreases αSyn toxicity in primary neuronal cultures

αSyn toxicity was induced in primary neurons by expressing human A53T αSyn via rAAV2/7 vector transduction on day 12 in vitro (DIV 12). On day 20, cultures expressing αSyn presented fewer neurons and a less dense network of neurites than cultures expressing the GFP control (Figure 1A), as confirmed by quantification of the MAP2-positive area fraction (Figure 1B). Cultures expressing αSyn showed an increased concentration of the cytosolic enzyme LDH in the medium (Figure 1C), indicating that the difference resulted from cell death and not only from reduced neurite outgrowth. Accordingly, neurons with αSyn expression were more vulnerable and showed a much more pronounced activation of caspase-3 when exposed to excitotoxic NMDA (supplemental Figure S1A).

Next, we assessed whether Rab7 is able to reduce αSyn toxicity in primary neurons by co-transducing cultures with rAAVs encoding either WT Rab7, DN Rab7 or GFP. WT Rab7 – but not DN Rab7 or GFP – alleviated the αSyn-induced reduction in MAP2-positive area by 20 % (Figure 1A, B). Similarly, the release of LDH into the culture medium was significantly decreased by WT Rab7 but not by DN Rab7 (Figure 1C). Furthermore, the NMDA-induced caspase-3 activation was lowered by WT Rab7 (supplemental Figure S1A).

To characterize αSyn aggregation in our neuronal culture model, we assayed the Triton X-100 solubility of αSyn in neuronal cell lysates (Figure 1D-G). αSyn was detected both in the Triton X-100 soluble fraction (Figure 1D) and in the Triton X-100 insoluble fraction (Figure 1F). Co-expression of WT but not DN Rab7 (evidenced by the HA-tag in Figure 1D) significantly reduced the amount of Triton X-100 insoluble αSyn (Figure 1F,G). The amount of Triton X-100 soluble αSyn was not decreased by Rab7 (Figure 1D) but rather showed a slight, but not significant increase (quantified in supplemental Figure S1B).

In order to corroborate the functional significance of αSyn aggregation on the neuronal network in culture, we quantified the amount of the postsynaptic protein PSD-95 using immunoblotting (Figure 1D,E). αSyn reduced the amount of PSD-95 in neuronal cultures, and this effect was mitigated by WT Rab7.

Rab7 is known for orchestrating trafficking of autophagosomal and endosomal vesicles. Accordingly, we previously demonstrated in HEK293T cells that Rab7 promotes the processing of αSyn into acidic vesicles such as autolysosomes [30]. To revisit this mode of Rab7 action, we determined in our primary neuronal cultures the fraction of phospho-αSyn present in LAMP1-positive vesicles. In αSyn expressing neurons, we observed fibrillary structures positive for phospho-αSyn (Figure 1H). These structures are reminiscent of αSyn structures observed after PFF-induced seeding of αSyn pathology in primary neurons [43]. In addition, we obsereved phosph-αSyn-positive puncta that colocalized with LAMP2. Upon quantification, the fraction of phospho-αSyn signal that colocalized with Lamp1 signal was larger with WT Rab7 than with DN Rab7 (Figure 1I). This finding is consistent with the hypothesis that Rab7 increases maturation of αSyn-containing autophagosomes.

Taken together, these findings confirmed the protective effect of WT Rab7 against αSyn aggregates and associated toxicity in mammalian neurons. The protective effects were not observed with DN Rab7, indicating that GTP binding is necessary for this Rab7 effect. Without αSyn expression, WT Rab7 had no obvious effects, confirming that its protective effect resulted from modulation of αSyn toxicity and not from direct, αSyn-independent effects.

Rab7 protects dopaminergic axon terminals and reduces αSyn accumulation in vivo

In order to further validate our findings, we tested the effect of WT Rab7 on αSyn-induced toxicity in dopaminergic neurons of the rat SN. To do this, animals were injected with the rAAV2/7 A53T αSyn vector combined with the rAAV2/7 WT Rab7 or rAAV2/7 DN Rab7 vector (Figure 2A). We chose DN Rab7 as the negative control for WT Rab7 because DNA and protein are identical except for the T22N point mutation. Additionally, we did not observe toxic effects of DN Rab7 in primary neurons – even when co-expressing αSyn (Figure 1). As expected, we observed in the SN of the injected hemisphere neurons positive for both human αSyn (Figure 2B, green) and the HA-tag of Rab7 (Figure 2B, red). Most of them were positive for both epitopes. The number of neurons in the SN positive for the HA-tag was not different between animals (Figure 2C). Neurons with human αSyn or HA-tag expression were not observed in the contralateral hemisphere (supplemental Figure S2A).

Dopaminergic neurons were identified by staining for tyrosine hydroxylase (TH, Figure 2D). Stereological quantification revealed a 65 % loss of TH-positive nigral neurons upon overexpression of αSyn (Figure 2D, E). In contrast to the protective effect of WT Rab7 in cultured neurons (Figure 1), the numbers of dopaminergic neurons were not different between animals injected with either αSyn + DN Rab7 or αSyn + WT Rab7 (Figure 2E).

Next, we analyzed the effect of Rab7 on the number of pathological phosphorylated αSyn-positive neurons in the SN. Four weeks after rAAV vector injection, the number of phospho-αSyn positive cells in the SN was not different between animals injected with αSyn + DN Rab7 and animals injected with αSyn + WT Rab7 (Figure 2F, G). In contrast, WT Rab7 decreased the amount of phospho-αSyn -positive structures in the area between the neurons (“neuropil”) in both the SN pars compacta (SNc, Figure 2F, H), and the SN pars reticulata (SNr, Figure 2F, I). We assume that a significant part of the phospho-αSyn-positive structures in the SN correspond to dendrites of dopaminergic neurons (basal dendrites in the SNc and apical dendrites in the SNr), but axonal structures and non-dopaminergic neurons could contribute to the signal as well.

Next, we characterized the effect of Rab7 on αSyn-induced loss of dopaminergic axon terminals (“fibers”) in the striatum (Figure 3A, B). In both groups, the density of striatal dopaminergic fibers was reduced to <50 % of the contralateral, non-injected hemisphere. Yet, the extent of the reduction was less pronounced in animals injected with αSyn + WT Rab7 compared to animals injected with αSyn + DN Rab7 (Figure 3B, C) – consistent with the protective effect of WT Rab7 on the density of the neuronal network in cultured neurons (Figure 1B). The extent of fiber loss in the striatum did not correlate with loss of somata in the SN (supplemental Figure S2B), but a differing response to toxic insults by somata of dopaminergic neurons in the SN and their axonal compartments in the striatum has been observed in numerous previous studies [44–46].

Retraction of dopaminergic axon terminals in the striatum could result from different stimuli, like the impaired health of the neuronal somata in the SNc, or from the propagation of αSyn pathology into the striatum. αSyn pathology was observed only in the injected hemisphere, and mostly detected along neurite-like structures (Figure 3D, insets green arrows). We observed both fibrillar and punctuated phospho-αSyn staining patterns in the striatum (Figure 3D, insets, green and red arrowheads) and assumed that the punctuating pattern represents a more advanced stage of neurite degeneration. Double staining for phospho-αSyn and TH revealed an overlapping distribution (supplemental Figure S2C), confirming that the majority of phospho-αSyn in the striatum is localized in dopaminergic terminals. The density of phospho-αSyn positive structures was significantly lower in animals expressing WT Rab7 compared to animals expressing DN Rab7 (Figure 3E). Additionally, blinded rating of images revealed that the more advanced punctuating pattern was more prevalent in animals expressing DN Rab7, while animals expressing WT Rab7 showed more of the fibrillary pattern (Figure 3F). These findings are in line with the observed protective effect of WT Rab7 on the striatal dopaminergic axonal terminals (Figure 3C), the reduced amount of Triton X-100 insoluble αSyn in cultured neurons (Figure 1G), the differences in phospho-αSyn positive dendrites in SNc and SNr (Figures 2H, I), and with our previous finding that WT Rab7 can induce the degradation of αSyn aggregates (Dinter et al., 2016).

The significant negative correlation between the amount of striatal phospho-αSyn and the density of dopaminergic fibers (Figure 3G) suggests that αSyn pathology is closely linked to the degeneration of axon terminals. The strong correlation between the density of striatal dopaminergic fibers in each animal and the amount of phospho- αSyn in the SNc (Figure 3H) and SNr (Figure 3L) indicates that αSyn pathology and degeneration are not local processes, but rather affect the entire neuron.

Next, we assessed whether Rab7 affected motor performance. Forelimb use is regulated by the extracellular concentration of dopamine in the contralateral striatum. αSyn-induced motor deficits were therefore quantified by the cylinder test, carried out four weeks after rAAV injection. Unilateral injection of rAAV αSyn combined with WT Rab7 or DN Rab7 into the right hemisphere reduced left forepaw use in almost all animals (Figure 3J), consistent with the fact that dopamine is secreted from nigrostriatal axon terminals and TH fibers were reduced in the right striatum (Figure 3C). Surprisingly, the forelimb asymmetry was larger with co-injection of WT Rab7 than with DN Rab7 (Figure 3J). This finding was unexpected given the blunted TH fiber loss by WT Rab7. However, it could be explained by an effect of Rab7 on the dopamine transporter (DAT), which is responsible for dopamine reuptake and termination of dopamine signaling. To test this hypothesis, we co-stained striatal sections for DAT and vesicular monoamine transporter 2 (VMAT2) (Figure 3K). Although the total area of striatal DAT signal was not different between groups (supplemental Figure S2D), the area of the DAT signal within VMAT2-positive fibers was significantly higher with WT Rab7 expression than with DN Rab7 (Figure 3L). Such a difference in DAT availability can explain the observed asymmetry in forelimb use [47].

αSyn pathology induces the activation of glia cells, which can induce toxic as well as protective responses [48,49]. αSyn expression induced microglia activation in the injected striatum (supplemental Figure S3A, C), but there was no difference between WT Rab7 and DN Rab7 injected animals. For astroglia, the αSyn-induced activation was moderate in animals with DN Rab7 and significantly higher in animals with WT Rab7 (supplemental Figure S3A, B). The negative correlation between astroglia reaction and striatal phospho-αSyn pathology (supplemental Figure S3D) could suggest a beneficial role of astroglia. Yet, the lacking correlation between the astroglia reaction and dopaminergic fiber density (supplemental Figure S3E) argues against a direct role of astroglia.

It is well established that dopaminergic axon terminals regenerate after toxic insults [50], and that αSyn can inhibit this regeneration [41]. In order to rule out that the differences in the density of striatal dopaminergic terminals in Figure 3C results from differences in terminal regeneration, we investigated whether WT Rab7 affects axon regrowth. Primary neurons were seeded in microfluidic chambers that separate axons from somata. After rAAV transduction of neurons with αSyn and Rab7, axons were dissected and the length of regenerated axons was measured five days later. Neither DN Rab7 nor WT Rab7 alone had an effect on regeneration (supplemental Figure S4A, B). In contrast, αSyn significantly reduced axon regeneration. The length of regenerated axons was shorter in cultures co-expressing WT Rab7 and αSyn (supplemental Figure S4A, B). Differences in axon sprouting therefore do not account for the differences between the densities of dopaminergic terminals in rat striatum.

Rab7 increases cellular viability

Accumulation of human αSyn impairs neuronal viability and neuronal functioning, which, in turn, causes further accumulation of toxic αSyn species. Therefore, mitigation of αSyn pathology by WT Rab7 overexpression should improve cellular functioning and increase viability (see graphical abstract). To test this hypothesis, we measured the amount of reactive oxygen species (ROS) and the mitochondrial membrane potential, two classical indicators of cellular functioning in primary neurons [1,51]. ROS levels and the mitochondrial membrane potential were normalized to control, non-αSyn -transduced cultures (represented as dashed lines in Figures 4A-C). Cultures transduced with rAAV-αSyn showed about 20 % more basal cytosolic ROS than cultures without αSyn (Figure 4A), consistent with our earlier findings [1]. αSyn increased mitochondrial ROS to a similar extent (Figure 4B), and reduced the mitochondrial membrane potential (Figure 4C). In addition to these steady-state measurements, we tested neurons’ response to an acute induction of oxidative and mitochondrial stress: Treatment with H₂O₂ increased cytoplasmic ROS (Figure 4A) while treatment with the mitochondrial uncoupler FCCP increased mitochondrial ROS (Figure 4B) and decreased the mitochondrial membrane potential (Figure 4C).

The αSyn-induced increase in steady-state cytoplasmic ROS was not affected by Rab7 (Figure 4A), but handling stress-induced ROS was more efficient in cultures with WT Rab7 and αSyn expression than in cultures expressing GFP or DN Rab7 with αSyn (Figure 4A). For mitochondrial ROS, both steady-state and stress-induced levels of mitochondrial ROS were lower in cultures expressing WT Rab7 with αSyn than in cultures expressing GFP or DN Rab7 with αSyn (Figure 4B). Finally, WT Rab7 stabilized mitochondrial membrane potential in the presence of αSyn and mitigated the stress-induced reduction in the mitochondrial membrane potential (Figure 4C). These findings suggest that the protective effects of WT Rab7 observed in cultured neurons (Figure 1A) and in the striatum (Figure 3B) could involve improved handling of αSyn-induced oxidative damage and reduction of mitochondrial impairment. In none of the measurements did DN Rab7 differ from GFP (Figure 4A-C), confirming that DN Rab7 does not induce additional toxicity in this paradigm.

Aggregated αSyn and ROS induce DNA damage [17,18,52,53]. We therefore measured αSyn-induced DNA damage by the comet assay. Nuclei of GFP infected neurons remained intact during electrophoresis (Figure 4D), whereas a significant amount of fragmented DNA was mobilized into comet tails from nuclei of αSyn transduced neurons (Figure 4D). The αSyn-induced tail moment was significantly decreased by co-transduction with WT Rab7 as compared to DN Rab7 or GFP (Figure 4D, E).

DNA breaks recruit specific repair proteins to the sites of the damage. Tumor suppressor p53-binding protein 1 (53BP1) is an effector of the canonical DNA damage response [54], and 53BP1 foci can be used to quantify the extent of DNA damage. The number of 53BP1 foci was significantly higher in αSyn expressing neurons than in the GFP control (Figure 4F, G). WT Rab7, but not DN Rab7, reduced the number of 53BP1 foci (Figure 4G), confirming the reduction of DNA fragmentation as reported by the comet assay (Figure 4E). Both methods also confirmed that expression of DN Rab7 per se did not induce DNA damage in neurons.

To confirm these Rab7 effects which were obtained in neuronal cultures, we determined the effect of αSyn and Rab7 on DNA damage in rat brain by quantifying the number of 53BP1 foci in neurons of rat SN (Figure 5). αSyn induced 53BP1 foci in SN neurons, and fewer 53BP1 foci were observed in animals transduced with WT Rab7 than with DN Rab7 (Figure 5A, B). The extent of DNA damage as reported by the number of 53BP foci correlated well with the density of dopaminergic axon terminals in the striatum of the same animal (Figure 5C), and similarly with the amount of phospho-αSyn in striatum (Figure 5D), SNr (Figure 5E) and SNc (Figure 5F). These correlations confirm the close link between degeneration of axon terminals, aggregate load and DNA damage. In conclusion they suggest that WT Rab7 reduces DNA damage by reducing αSyn aggregates.

Here we report that the expression of WT Rab7 reduced αSyn pathology in primary neurons and in the rodent brain. Transduction with WT Rab7 abated αSyn-induced pathology as shown by the increased density of MAP2-positive neurons, the decreased LDH release and caspase-3 activation in primary cultures and by the decreased lesion of dopaminergic axon terminals in the rat striatum. The fraction of αSyn in lysosomal vesicles was increased by Rab7, indicating that toxic αSyn species are reduced via autolysosomal clearance. WT Rab7 reduced αSyn-induced ROS production, mitochondrial impairment and DNA damage, confirming the functional relevance of reducing αSyn pathology.

We induced αSyn pathology and toxicity by overexpressing A53T αSyn in the SN of rats and in primary mouse neurons using rAAV. Overexpression of αSyn in the rat brain induced a robust phenotype with a 60 % reduction of both the number of dopaminergic somata in the SN (Figure 2E) and dopaminergic axon terminals in the striatum (Figure 3C) – in line with previous reports [32]. To promote the autolysosomal degradation of toxic αSyn species and to reduce αSyn dependent phenotypes, we expressed WT Rab7. The DN mutant of Rab7 was used as a negative control. With WT Rab7, the densities of phospho-αSyn positive structures in SNc and SNr were reduced (Figure 2), as was the density of phospho-αSyn positive axon terminals in the striatum (Figure 3 and supplemental Figure S2C). In primary neurons, we observed a reduced amount of αSyn in the Triton X-100 insoluble fraction (Figure 1F-G) and a higher colocalization between phospho-αSyn and Lamp1 with WT Rab7 compared to DN Rab7 or GFP (Figure 1H), indicating that Rab7 promotes the fusion of αSyn containing autophagosomes with lyososomes. This is consistent with our previous finding that Rab7 increases the fraction of αSyn in autolysosomes as reported by RFP-GFP tandem fluorescence [30], and with the known promotion of autophagosome maturation by Rab7 [29,55–58].

DN Rab7 did not increase the accumulation of insoluble αSyn (Figure 1F, G) or induce toxicity (Figure 1B, C, Figure 4). Accordingly, there is no direct evidence that PD can be caused by a deficit in Rab7 activity. Yet, mutations in the RAB7 gene are associated with melanoma [59], which is more common in PD patients. In addition, several other proteins involved in trafficking of endosomes and autophagosomes are linked to PD, including RabGEF1 and Arl8b [24]. RabGEF1 activates Rab5 [60] which characterizes early endosomes and is subsequently exchanged for Rab7. Both Rab7 and Arl8 are localized on late endosomes [60–62], and Rab7 is exchanged for Arl8 during endosome maturation [62]. Thus, the autophagolysosomal pathway clearly is important during PD pathogenesis, and stimulating autophagosome maturation can be a viable therapeutic strategy even without a causative role of Rab7 during PD pathogenesis.

Because Rab7 is a major signaling hub [63–65], directly increasing its activity might induce several off-target effects. This is evidenced by our behavioral findings (Figure 3J). Neurons transduced with WT Rab7 are healthier as evidenced by reduced DNA damage in the SNc (Figure 5B) and by the preserved mitochondrial function in our neuronal cultures (Figure 4B-C). Synaptic dopamine is cleared through the DAT, a secondary active transporter. Improved cellular health in neurons expressing WT Rab7 can therefore explain increased dopamine clearance, which in turn can underlie reduced forelimb use [47]. In addition, we observed a higher abundance of DAT in axon terminals expressing WT Rab7 as compared to DN Rab7 (Figure 3L). This is not unexpected given that Rab7 regulates endosomal trafficking. A higher abundance of DAT in striatal axon terminals can also explain increased dopamine clearance and reduced forelimb use. Along similar lines, altered striatal dopamine clearance underlies the paradoxically improved motor performance in a mouse model of αSyn pathology [66]. These observations highlight the Janus-faced consequences of dopaminergic terminal loss in the striatum: On the one hand, fewer terminals secrete less dopamine. On the other hand, reduced dopamine clearance and receptor sensitisation potentiate dopamine effects. Accordingly, dyskinesias are induced more easily in dopamine-denervated striatum and supporting striatal DAT activity can reduce dyskinesias [67].

We observed a tight correlation between (1) the degeneration of dopaminergic terminals in the striatum, (2) αSyn pathology and (3) DNA damage (Figure 3G-I and Figure 5C-F). The fact that corresponding values for animals treated with either WT Rab7 or DN Rab7 aligned on the same regression curve suggests that the protection of dopaminergic axon terminals by WT Rab7 was achieved by the reduction of αSyn pathology and the subsequent reduction of DNA damage. Cellular functioning and αSyn pathology are intricately linked and the individual pathogenic steps potentiate each other (see graphical abstract). Our findings suggest that stimulating the rate limiting step of autolysosomal αSyn aggregate clearance by rAAV-mediated WT Rab7 expression breaks this vicious circle, and the improved cellular functioning limits further αSyn aggregation.

WT Rab7 reduced αSyn-induced degeneration of primary neurons and striatal dopaminergic axon terminals. WT Rab7 also lowered αSyn pathology and associated toxicity, including oxidative stress, mitochondrial impairment and DNA damage. These findings confirm that improving autophagosome maturation is a possible therapeutic strategy for PD.

53BP1 - tumor suppressor p53-binding protein 1

DAB - 3,3-diaminobenzidine

DAT – dopamine transporter

DCFH-DA - dichlorodihydroﬂuorescein diacetate

DIV – days in vitro

DLB – dementia with Lewy bodies

DN – dominant negative

GC – genome copy

GFP – green fluorescent protein

JC-1 - tetraethylbenzimidazolylcarbocyanine iodide

LDH – lactate dehydrogenase

LN – Lewy neurites

MSA – multiple system atrophy

PD – Parkinson’s disease

phospho-αSyn – phosphorylated alpha-synuclein

rAAV – recombinant adeno-associated viral vector

Rab7 – Ras-related in brain 7

ROS – reactive oxygen species

SN – substantia nigra

SNc – substantia nigra pars compacta

SNr – substantia nigra pars reticulata

TBS – Tris-buffered saline

TH – tyrosine hydroxylase

VMAT2 – vesicular monoamine transporter 2

WT – wild type

αSyn – alpha-synuclein

Ethical approval and consent to participate

Animals were approved by local authorities (Bioethical Committee of the KU Leuven, Belgium, ECD project P067-2013 and P085-2014) and conducted in accordance with guidelines of the Federation for European Laboratory Animal Science Associations (FELASA).

Consent for publication

All authors approved the manuscript.

Availability of data and materials

All data are included in the text. Materials are available upon reasonable request.

Competing interests

The authors declare that they have no competing interests.

Funding

This work was supported by the German Research Foundation (DFG to BF, FA 658/3-1) and by the FWO (postdoctoral fellowship to AVDP and project G080517N), the KU Leuven (C14/18/102), the Medical Foundation Queen Elisabeth and the ERA-NET JPco-fuND 2015 SYNACTION

Authors' contributions

AVDP, VB and BF conceived research. EMSz, CVDH, LH, AVDP performed research. EMSz, AVDP and BF wrote the manuscript, all authors contributed and approved the manuscript.

Acknowledgements

We thank Sabine Hamm, Anett Böhm and Annelies Aertgeerts for excellent technical assistance.

Szegő ÉM, Dominguez-Meijide A, Gerhardt E, König A, Koss DJ, Li W, et al. Cytosolic Trapping of a Mitochondrial Heat Shock Protein Is an Early Pathological Event in Synucleinopathies. Cell Rep. 2019;28.
Junn E, Mouradian MM. Human α-Synuclein over-expression increases intracellular reactive oxygen species levels and susceptibility to dopamine. Neurosci Lett; 2002;320:146–50.
Winslow AR, Rubinsztein DC. The Parkinson disease protein α-synuclein inhibits autophagy. Autophagy; 2011;429–31.
Suzuki G, Imura S, Hosokawa M, Katsumata R, Nonaka T, Hisanaga SI, et al. α-synuclein strains that cause distinct pathologies differentially inhibit proteasome. Elife; 2020;9:1–21.
Wang D, Yu T, Liu Y, Yan J, Guo Y, Jing Y, et al. DNA damage preceding dopamine neuron degeneration in A53T human α-synuclein transgenic mice. Biochem Biophys Res Commun; 2016;481:104–10.
Goodwin J, Nath S, Engelborghs Y, Pountney DL. Raised calcium and oxidative stress cooperatively promote alpha-synuclein aggregate formation. Neurochem. Int; 2013; 703–11.
Scudamore O, Ciossek T. Increased oxidative stress exacerbates α-synuclein aggregation in vivo. J Neuropathol Exp Neurol; 2018;77:443–53.
Bence NF, Sampat RM, Kopito RR. Impairment of the ubiquitin-proteasome system by protein aggregation. Science (80- ). 2001;292:1552–5.
Klucken J, Poehler AM, Ebrahimi-Fakhari D, Schneider J, Nuber S, Rockenstein E, et al. Alpha-synuclein aggregation involves a bafilomycin A1-sensitive autophagy pathway. Autophagy; 2012;8:754–66.
Minakaki G, Menges S, Kittel A, Emmanouilidou E, Schaeffner I, Barkovits K, et al. Autophagy inhibition promotes SNCA/alpha-synuclein release and transfer via extracellular vesicles with a hybrid autophagosome-exosome-like phenotype. Autophagy; 2018;14:98–119.
Poehler AM, Xiang W, Spitzer P, May VEL, Meixner H, Rockenstein E, et al. Autophagy modulates SNCA/α-synuclein release, thereby generating a hostile microenvironment. Autophagy; 2014;10:2171–92.
Kiffin R, Bandyopadhyay U, Cuervo AM. Oxidative stress and autophagy. Antioxidants Redox Signal; 2006;152–62.
Guo JY, Karsli-Uzunbas G, Mathew R, Aisner SC, Kamphorst JJ, Strohecker AM, et al. Autophagy suppresses progression of K-ras-induced lung tumors to oncocytomas and maintains lipid homeostasis. Genes Dev; 2013;27:1447–61.
Galati S, Boni C, Gerra MC, Lazzaretti M, Buschini A. Autophagy: A Player in response to Oxidative Stress and DNA Damage. Oxid Med Cell Longev; 2019
Burgoyne JR. Oxidative stress impairs autophagy through oxidation of ATG3 and ATG7. Autophagy; 2018. p. 1092–3.
Frudd K, Burgoyne T, Burgoyne JR. Oxidation of Atg3 and Atg7 mediates inhibition of autophagy. Nat Commun; 2018;9.
Paiva I, Pinho R, Pavlou MA, Hennion M, Wales P, Schütz AL, et al. Sodium butyrate rescues dopaminergic cells from alpha-synuclein-induced transcriptional deregulation and DNA damage. Hum Mol Genet. 2017;26:2231–46.
Milanese C, Cerri S, Ulusoy A, Gornati S V., Plat A, Gabriels S, et al. Activation of the DNA damage response in vivo in synucleinopathy models of Parkinson’s disease. Cell Death Dis; 2018;9.
Eapen V V., Waterman DP, Bernard A, Schiffmann N, Sayas E, Kamber R, et al. A pathway of targeted autophagy is induced by DNA damage in budding yeast. Proc Natl Acad Sci U S A; 2017;114:E1158–67.
Sidransky E, Nalls MA, Aasly JO, Aharon-Peretz J, Annesi G, Barbosa ER, et al. Multicenter analysis of glucocerebrosidase mutations in Parkinson’s disease. N Engl J Med; 2009;361:1651–61.
Chang D, Nalls MA, Hallgrímsdóttir IB, Hunkapiller J, Brug M van der, Cai F, et al. A meta-analysis of genome-wide association studies identifies 17 new Parkinson’s disease risk loci. Nat Genet; 2017;49:1511–6.
Verstraeten A, Theuns J, Van Broeckhoven C. Progress in unraveling the genetic etiology of Parkinson disease in a genomic era. Trends Genet; 2015. p. 140–9.
Gao Y, Wilson GR, Stephenson SEM, Bozaoglu K, Farrer MJ, Lockhart PJ. The emerging role of Rab GTPases in the pathogenesis of Parkinson’s disease. Mov. Disord; 2018. p. 196–207.
Bandres-Ciga S, Saez-Atienzar S, Bonet-Ponce L, Billingsley K, Vitale D, Blauwendraat C, et al. The endocytic membrane trafficking pathway plays a major role in the risk of Parkinson’s disease. Mov Disord; 2019;34:460–8.
Baba T, Toth DJ, Sengupta N, Kim YJ, Balla T. Phosphatidylinositol 4,5‐bisphosphate controls Rab7 and PLEKHM 1 membrane cycling during autophagosome–lysosome fusion. EMBO J; 2019;38.
Jimenez-Orgaz A, Kvainickas A, Nägele H, Denner J, Eimer S, Dengjel J, et al. Control of RAB7 activity and localization through the retromer-TBC1D5 complex enables RAB7-dependent mitophagy. EMBO J; 2018;37:235–54.
Kjos I, Borg Distefano M, Sætre F, Repnik U, Holland P, Jones AT, et al. Rab7b modulates autophagic flux by interacting with Atg4B. EMBO Rep.; 2017;18:1727–39.
Kuchitsu Y, Fukuda M. Revisiting Rab7 Functions in Mammalian Autophagy: Rab7 Knockout Studies. Cells; 2018;7:215.
Zhan L, Chen S, Li K, Liang D, Zhu X, Liu L, et al. Autophagosome maturation mediated by Rab7 contributes to neuroprotection of hypoxic preconditioning against global cerebral ischemia in rats. Cell Death Dis; 2017;8:e2949.
Dinter E, Saridaki T, Nippold M, Plum S, Diederichs L, Komnig D, et al. Rab7 induces clearance of α-synuclein aggregates. J Neurochem; 2016;138:758–74.
Karpinar DP, Balija MBG, Kügler S, Opazo F, Rezaei-Ghaleh N, Wender N, et al. Pre-fibrillar alpha-synuclein variants with impaired beta-structure increase neurotoxicity in Parkinson’s disease models. EMBO J; 2009;28:3256–68.
Van der Perren A, Toelen J, Casteels C, Macchi F, Van Rompuy AS, Sarre S, et al. Longitudinal follow-up and characterization of a robust rat model for Parkinson’s disease based on overexpression of alpha-synuclein with adeno-associated viral vectors. Neurobiol Aging; 2015;36:1543–58.
Van Der Perren A, Toelen J, Carlon M, Van Den Haute C, Coun F, Heeman B, et al. Efficient and stable transduction of dopaminergic neurons in rat substantia nigra by rAAV 2/1, 2/2, 2/5, 2/6.2, 2/7, 2/8 and 2/9. Gene Ther; 2011;18:517–27.
Spinosa MR, Progida C, De Luca A, Colucci AMR, Alifano P, Bucci C. Functional characterization of Rab7 mutant proteins associated with Charcot-Marie-Tooth type 2B disease. J Neurosci; 2008;28:1640–8.
Park JW, Vahidi B, Taylor AM, Rhee SW, Jeon NL. Microfluidic culture platform for neuroscience research. Nat Protoc; 2006;1:2128–36.
Van der Perren A, Casteels C, Van Laere K, Gijsbers R, Van den Haute C, Baekelandt V. Development of an Alpha-synuclein Based Rat Model for Parkinson’s Disease via Stereotactic Injection of a Recombinant Adeno-associated Viral Vector. J Vis Exp; 2016;2016:53670.
Dunn KW, Kamocka MM, McDonald JH. A practical guide to evaluating colocalization in biological microscopy. Am. J. Physiol; 2011;p. C723.
Baekelandt V, Claeys A, Eggermont K, Lauwers E, De Strooper B, Nuttin B, et al. Characterization of lentiviral vector-mediated gene transfer in adult mouse brain. Hum Gene Ther; 2002;13:841–53.
Schmitz C, Hof PR. Design-based stereology in neuroscience. Neuroscience; 2005;p. 813–31.
Szegő ÉM, Gerhardt E, Outeiro TF. Sirtuin 2 enhances dopaminergic differentiation via the AKT/GSK-3β/β-catenin pathway. Neurobiol Aging; 2017;56:7–16.
Szegő ÉM, Gerhardt E, Kermer P, Schulz JB. A30P α-synuclein impairs dopaminergic fiber regeneration and interacts with L-DOPA replacement in MPTP-treated mice. Neurobiol Dis; 2012;45:591–600.
Oliveira JMA, Chen S, Almeida S, Riley R, Gonçalves J, Oliveira CR, et al. Mitochondrial-dependent Ca2+ handling in Huntington’s disease striatal cells: Effect of histone deacetylase inhibitors. J Neurosci; 2006;26:11174–86.
Volpicelli-Daley LA, Luk KC, Lee VM-Y. Addition of exogenous α-synuclein preformed fibrils to primary neuronal cultures to seed recruitment of endogenous α-synuclein to Lewy body and Lewy neurite-like aggregates. Nat. Protoc; 2014 ;p. 2135–46.
Masliah E, Rockenstein E, Veinbergs I, Mallory M, Hashimoto M, Takeda A, et al. Dopaminergic loss and inclusion body formation in α-synuclein mice: Implications for neurodegenerative disorders. Science; 2000;287:1265–9.
Chung CY, Koprich JB, Siddiqi H, Isacson O. Dynamic changes in presynaptic and axonal transport proteins combined with striatal neuroinflammation precede dopaminergic neuronal loss in a rat model of AAV α-synucleinopathy. J Neurosci; 2009;29:3365–73.
Koch JC, Bitow F, Haack J, d’Hedouville Z, Zhang J-NN, Tönges L, et al. Alpha-Synuclein affects neurite morphology, autophagy, vesicle transport and axonal degeneration in CNS neurons. Cell Death Dis; 2015;6:e1811.
Lee J, Zhu WM, Stanic D, Finkelstein DI, Horne MH, Henderson J, et al. Sprouting of dopamine terminals and altered dopamine release and uptake in Parkinsonian dyskinaesia. Brain; 2008;131:1574–87.
Pekny M, Wilhelmsson U, Pekna M. The dual role of astrocyte activation and reactive gliosis. Neurosci. Lett; 2014. p. 30–8.
Nichols MR, St-Pierre MK, Wendeln AC, Makoni NJ, Gouwens LK, Garrad EC, et al. Inflammatory mechanisms in neurodegeneration. J. Neurochem; 2019. p. 562–81.
Kowsky S, Pöppelmeyer C, Kramer ER, Falkenburger BH, Kruse A, Klein R, et al. RET signaling does not modulate MPTP toxicity but is required for regeneration of dopaminergic axon terminals. Proc Natl Acad Sci U S A; 2007;
Musgrove RE, Helwig M, Bae EJ, Aboutalebi H, Lee SJ, Ulusoy A, et al. Oxidative stress in vagal neurons promotes parkinsonian pathology and intercellular α-synuclein transfer. J Clin Invest; 2019;129:3738–53.
Schaser AJ, Osterberg VR, Dent SE, Stackhouse TL, Wakeham CM, Boutros SW, et al. Alpha-synuclein is a DNA binding protein that modulates DNA repair with implications for Lewy body disorders. Sci Rep; 2019;9:1–19.
Martin LJ, Pan Y, Price AC, Sterling W, Copeland NG, Jenkins NA, et al. Parkinson’s disease α-synuclein transgenic mice develop neuronal mitochondrial degeneration and cell death. J Neurosci; 2006;26:41–50.
Panier S, Boulton SJ. Double-strand break repair: 53BP1 comes into focus. Nat. Rev. Mol. Cell Biol; 2014. p. 7–18.
Gutierrez MG, Munafó DB, Berón W, Colombo MI. Rab7 is required for the normal progression of the autophagic pathway in mammalian cells. J Cell Sci; 2004;117:2687–97.
Jäger S, Bucci C, Tanida I, Ueno T, Kominami E, Saftig P, et al. Role for Rab7 in maturation of late autophagic vacuoles. J Cell Sci; 2004;117:4837–48.
Zhao YG, Zhang H. Autophagosome maturation: An epic journey from the ER to lysosomes. J. Cell Biol; 2019;757–70.
Wang Z, Miao G, Xue X, Guo X, Yuan C, Wang Z, et al. The Vici Syndrome Protein EPG5 Is a Rab7 Effector that Determines the Fusion Specificity of Autophagosomes with Late Endosomes/Lysosomes. Mol Cell; 2016;63:781–95.
Alonso-Curbelo D, Riveiro-Falkenbach E, Pérez-Guijarro E, Cifdaloz M, Karras P, Osterloh L, et al. RAB7 Controls Melanoma Progression by Exploiting a Lineage-Specific Wiring of the Endolysosomal Pathway. Cancer Cell; 2014;26:61–76.
Horiuchi H, Lippé R, McBride HM, Rubino M, Woodman P, Stenmark H, et al. A novel Rab5 GDP/GTP exchange factor complexed to Rabaptin-5 links nucleotide exchange to effector recruitment and function. Cell; 1997;90:1149–59.
Marwaha R, Arya SB, Jagga D, Kaur H, Tuli A, Sharma M. The Rab7 effector PLE KHM1 binds Arl8b to promote cargo traffic to lysosomes. J Cell Biol; 2017;216:1051–70.
Jongsma ML, Bakker J, Cabukusta B, Liv N, Elsland D, Fermie J, et al. SKIP HOPS recruits TBC1D15 for a Rab7‐to‐Arl8b identity switch to control late endosome transport. EMBO J; 2020;39.
Wen H, Zhan L, Chen S, Long L, Xu E. Rab7 may be a novel therapeutic target for neurologic diseases as a key regulator in autophagy. J Neurosci Res; 2017;95:1993–2004.
Langemeyer L, Fröhlich F, Ungermann C. Rab GTPase Function in Endosome and Lysosome Biogenesis. Trends Cell Biol; 2018;957–70.
Zhang M, Chen L, Wang S, Wang T. Rab7: Roles in membrane trafficking and disease. Biosci. Rep; 2009;193–209.
Hunn BHM, Vingill S, Threlfell S, Alegre-Abarrategui J, Magdelyns M, Deltheil T, et al. Impairment of Macroautophagy in Dopamine Neurons Has Opposing Effects on Parkinsonian Pathology and Behavior. Cell Rep; 2019;29:920-931.e7.
Tomas D, Stanic D, Chua HK, White K, Boon WC, Horne M. Restoration of the Dopamine Transporter through Cell Therapy Improves Dyskinesia in a Rat Model of Parkinson’s Disease. PLoS One; 2016;11:e0153424.

SzegoRab7aSynFigureSI1.jpg
A) αSyn-induced activation of caspase-3 in the presence of 100 µM NMDA (n=3, mean ± SD, ANOVA followed by Bonferroni posthoc test; **p=0,00421). (B) Quantification of the amount of human αSyn (normalized to β-tubulin signal) in the Triton X-100 soluble fraction of lysates from three independent preparations (n=3, mean ± SD, ANOVA followed by Bonferroni posthoc test). Representative western blot is shown on Figure 1D. (C) Uncropped membranes from Figure 1D showing human aSyn, rodent aSyn, PSD95, HA-tag and β-tubulin signal.
SzegoRab7aSynFigureSI2.jpg
(A) Representative images showing HA-tag staining in the substantia nigra, contralateral to the rAAV injection. Images showing HA-tag staining in the ipsilateral hemisphere are presented on Figure 2C. Scale bar: 200 µm (B) Linear regression of the numbers of dopaminergic neurons in the substantia nigra (from Figure 2D, E) and densities of dopaminergic axon terminals in the striatum (from Figure 3B, C; p=0,8546; r2=0,001917). (C) Representative images showing colocalization of phospho-αSyn (red) and TH (green) in the striatum of αSyn + DN Rab7 (left panel) or WT Rab7 (right panel) rAAV-injected animals. Scale bar: 20 µm (D) Quantification of total area fraction of the striatal DAT signal (n=10, mean ± SD, t-test; p=0,0534). Representative images showing DAT (green) and VMAT2 (red) staining are on Figure 3K, thresholded images used for total DAT signal measurement are on Figure 3K2 and K2.
SzegoRab7aSynFigureSI3.jpg
(A) Representative images showing the astroglia marker GFAP (green) and the microglia marker Iba1 (red) in the contralateral (left panel) and ipsilateral (right panel) striatum of αSyn + DN Rab7 (upper panel) and αSyn + WT Rab7 (lower panel) rAAV-injected animals. Scale bar: 100 µm (B) Quantification of the area fraction of GFAP signal (normalized to the contralateral hemisphere; n=10, mean ± SD, t-test; *p=0,020). (C) Quantification of the area fraction of Iba1 signal (normalized to the contralateral hemisphere, n=10, mean ± SD, t-test). (D) Linear regression of the GFAP ratio (from supplemental Figure 3B) and the density of phospho-αSyn-positive structures in the striatum (from Figure 3E, p=0,0119, r2=0,3028). (E) Linear regression of the GFAP ratio (from supplemental Figure 3B) and the density of dopaminergic terminals in the striatum (from Figure 3C; p=0,0663, r2=0,1752)..
SzegoRab7aSynFigureSI4.jpg
(A) Representative, stiched images showing tubulin-stained axons grown in microfluidic chambers (axonal compartment), 5 days after mechanical lesion. Primary cultures were infected with (GFP (left) or αSyn (right)) and with (GFP (upper), DN Rab7 (middle) or WT Rab7 (lower) rAAV. Scale bar: 100 µm (B) Quantification of the length of tubulin-stained axons in the axonal compartment of microfluidic chambers (n=3, mean ± SD, ANOVA followed by Bonferroni posthoc test; p=0,0468).
SzegoRab7aSynSupplementalTableS1.docx

Download PDF

Journal Publication

published 01 Jan, 2022

Read the published version in Experimental Neurology →

Version 1

posted

You are reading this latest preprint version

Rab7 Reduces α-Synuclein Toxicity in Rats and Primary Neurons

Status:

Journal Publication

Version 1

Abstract

Figures

Background

Methods

Results

Discussion

Conclusion

Abbreviations

Declarations

References

Supplementary Files

Status:

Journal Publication

Version 1