Identification of Glycerol 3-phosphate acyltransferase as a potent modifier of α-Synuclein-induced toxicity

doi:10.21203/rs.3.rs-4199316/v1

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Identification of Glycerol 3-phosphate acyltransferase as a potent modifier of α-Synuclein-induced toxicity

https://doi.org/10.21203/rs.3.rs-4199316/v1

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Although multiple cellular pathways have been implicated in a-Synuclein (a-syn)-associated Parkinson’s disease (PD), the role of lipid metabolism remains elusive. Using the Drosophila system as a genetic screening tool, we identified mino, which encodes the mitochondrial isoform of the lipid synthesis enzyme glycerol 3-phosphate acyltransferase (GPAT), as a potent modifier of a-syn. Silencing the expression of mino significantly suppresses a-syn-induced PD phenotypes in Drosophila, including dopaminergic neuronal loss and locomotion defects as well as circadian rhythm-related activities, whereas mino overexpression yields opposite effects. Mechanistically, we found that mino modulates the levels of mitochondrial reactive oxygen speciesand lipid peroxidation. Importantly, treatment of a-syn-expressing flies with FSG67, a GPAT inhibitor, reproduces the benefits of mino knockdown. FSG67 also inhibited a-syn aggregation and lipid peroxidation in mouse primary neurons transfected with a-syn preformed fibrils. Our study elucidates an important factor contributing to a-syn toxicity and offers a novel therapeutic direction for PD.

Biological sciences/Neuroscience/Diseases of the nervous system/Parkinson's disease

Biological sciences/Genetics/RNAi

Health sciences/Diseases/Neurological disorders/Parkinson's disease

Parkinson's disease

lipid

glycerol 3-phosphate acyltransferase

GPAT

FSG67

Drosophila screen identifies mino (glycerol 3-phosphate acyltransferase) as a novel modifier of a-Synuclein (a-syn)
Silencing of mino expression abrogates the pathological phenotypes in flies induced by elevated levels of a-syn.
Silencing the expression of other GPAT similarly produces beneficial outcomes in a-syn flies, suggesting a common role of GPAT in modulating a-syn neurotoxicity.
Pharmacological inhibition of GPAT via FSG67 treatment inhibits a-syn aggregation and lipid peroxidation in both fly brains and mouse primary neurons.

Parkinson’s disease (PD) is a prevalent neurodegenerative disease that is characterized clinically by a constellation of motoric deficits including bradykinesia, tremors, and rigidity. The principal neuropathology underlying PD is the loss of dopaminergic (DA) neurons in the substantia nigra pars compacta of the midbrain, which results in the disruption of the nigrostriatal pathway leading to movement disorders. Accompanying the DA neurodegeneration is the presence of intraneuronal inclusions known as Lewy bodies (LBs), which are enriched with the presynaptic protein α-syn ^1,2.

Notably, rare mutations and copy number variations in the α-syn-encoding gene SNCA, such as A30P and A53T, are causative of monogenic forms of PD ^3–5. Further, genome-wide association studies have also implicated genetic variations of SNCA increasing the risk for idiopathic PD ⁶. However, the effects of these genetic variations are modest compared to other PD risk genes, like Gba1 or LRRK2 ⁷, potentially attributable to the polygenetic mode of inheritance and genetic modifiers' interaction with SNCA ^8,9. Interestingly, a recent retrospective analysis of studies on α-syn revealed interactions with up to 365 genetic modifiers, underscoring α-syn's intricate involvement in PD pathology ¹⁰. These modifiers are linked to diverse biological pathways, including mitochondrial function, antioxidant/glutathione metabolism, autophagy, protein degradation, vesicle trafficking, neuroinflammation, cytoskeleton dynamics, and lipid metabolism. In particular, the role of lipid dyshomeostasis in the development of PD has gained much attention in recent years. LBs, traditionally seen as comprised mainly of α-syn, is now recognized to be a complex mixture of protein aggregates intertwined with lipid components including from damaged mitochondria, autophagosomes, lysosomes, and fragments of lipid membranes, emphasizing the pivotal role of lipids in the biogenesis of LB associated with PD pathogenesis ^11,12. Furthermore, several genes linked to PD, such as PLA2G6, GBA, LRRK2, Parkin, SMPD1, SCARB2, DGKQ and SYNJ1, play important roles in lipid metabolism ^13–15. Not surprisingly, lipidomic irregularities are observed in the serum and brain tissues of PD patients, including elevated levels of polyunsaturated fatty acids (PUFAs), glycerophosphatidylcholine, diacylglycerol, and triglycerides ^14,16–18.

α-syn is known to interact with lipids and its aberrations disrupts this interaction. Disease-related mutations in α-syn, such as A30P, E46K, and G51D, all of which are clustered in the protein's N-terminus, alter its affinity for lipid membranes and influence the balance between its multimeric and monomeric states ^19–21. α-syn deficiency can lead to changes in palmitate uptake and brain neutral lipid content, affecting the acylation of arachidonic acid and docosahexaenoic acid, and their conversion into phospholipid pools ^22–24. Conversely, excessive α-syn induces the accumulation of lipid droplets in yeast ²⁵, and promotes the synthesis of polyunsaturated ether phospholipids in iPSC-derived human midbrain dopaminergic neurons, leading to lipid peroxidation and increased sensitivity to ferroptosis, a form of cell death marked by iron-dependent accumulation of lipid peroxides ²⁶. α-syn also contributes to ferroptosis by generating lipid peroxidation ²⁷.

Various lipid-metabolizing enzymes are known to affect α-syn aggregation and associated neurotoxicity. For example, stearoyl-CoA desaturase (SCD) regulate the unsaturation level of fatty acids and affect α-syn oligomeric composition in yeast, cells and transgenic mice ^28–32; Fatty acid CoA synthetase long-chain family member 4 (ACSL4) regulates fatty acids partitioning into the fatty acyl-CoA pool and influences lipid peroxidation and ferroptosis that are linked to α-syn-induced neurotoxicity ^26,33,34. Lysophosphatidic acid acyltransferase (LPACT), which converts lysophospholipids to phospholipids and helps to reconstitute phosphatidylcholine, is upregulated in postmortem PD brains and cultured neurons overexpressing SNCA E35K/E46K/E61K mutants ³⁵. Finally, LIPE, the rate-limiting lipase enzyme, regulates α-syn tetramer:monomer equilibrium and α-syn neurotoxicity ³⁶. Accordingly, modulating fatty acid and lipid metabolism genes, including LPCAT, SCD and LIPE, may reverse α-syn aggregation and associated phenotypes, which was observed in several PD models ^30,35,36. Furthermore, YTX-7739, an SCD inhibitor, has progressed to phase 1 clinical trials ²⁹. Taken together, it is important to clarify the role of lipids in the pathogenesis of PD that holds promise to shed light on novel therapeutic directions for PD treatment.

Using transgenic Drosophila overexpressing α-syn as a model for genetic screening, we report here the identification of mino, which encodes the mitochondrial isoform of the lipid synthesis enzyme glycerol 3-phosphate acyltransferase (GPAT), as a potent modifier of α-syn. We find that siRNA-mediated silencing of mino expression significantly mitigates α-syn-induced PD phenotypes in Drosophila, including dopaminergic neuronal loss and associated locomotion defects as well as circadian rhythm-related activities, whereas overexpression of yields opposite effects. The beneficial effects of mino silencing on α-syn-expressing flies are linked to reduced levels of mitochondrial reactive oxygen species and lipid peroxidation. Additionally, we extend our investigation to include the predicted endoplasmic reticulum-localized GPAT (CG15450 and Gpat4) and the related gene Gnpat, that encodes peroxisome-localized glyceronephosphate O-acyltransferase. Knocking down Gpat4, or CG15450 similarly reduces α-syn aggregation and suppresses α-syn neurotoxicity, suggesting a common role of GPAT in modulating α-syn neurotoxicity. Consistent with this, treatment of α-syn-expressing flies with FSG67, a GPAT inhibitor, reproduces the benefits of mino/GPAT knockdown. Notably, FSG67 also inhibits α-syn aggregation and lipid peroxidation in mouse primary neurons transfected with α-syn preformed fibrils. Taken together, our study elucidates an important factor/pathway contributing to α-syn toxicity and offers a novel therapeutic direction for PD.

Drosophila screen identifies mino RNAi as a suppressor of α-syn neurotoxicity

To identify modifiers of α-syn, we performed a genetic screen in a Drosophila model of synucleinopathy, where we overexpressed human wild-type α-syn in the photoreceptor (PR) neurons R1-R6. Overexpression of α-syn in PR neurons is known to induce vacuole formation in the lamina layer, where PR neurons form synapses with downstream lamina monopolar neurons ³⁷, which offers a convenient readout. To identify candidate genes for screening, we utilized a predictive model deltaSVM to analyze a collection of SNPs dataset on the Parkinson's disease genome-wide association study locus browser (v1.9) from 83 human PD risk loci that may confer differential transcription factor binding ^38,39. Based on the SNP locations, we manually selected 1–3 nearest genes for each locus. Subsequently, we identified available RNAi or dCas9/gRNA-VPR overexpression lines from the Bloomington Drosophila Stock Center (BDSC). These lines targeted the Drosophila orthologues of the selected human genes, which were identified using the DIOPT Ortholog Finder ⁴⁰. Finally, we crossed 107 lines targeting 92 genes with the Drosophila synucleinopathy model to examine whether they can influence α-syn-induced vacuole formation in the lamina ⁴¹ (Supplementary Data 1).

From the above pool of 107 RNAi and overexpression fly lines, we found 5 RNAi lines (targeting Ggamma1, mino, nonC, RpII215 or Src64B) and 9 overexpression lines (targeting Acbp1, Bap170, bnl, CG5214, CG7461, Psn, Pu, SdhC, andor wg) that suppress vacuole formation in α-syn flies by more than 2-fold at an adjusted p-value of 0.01 (Fig. 1A). We subsequently selected four lines whose targeted genes are associated with energy and lipid metabolism, i.e. mino, CG5214, CG7461, SdhC, to test if their RNAi or overexpression could rescue the decline in climbing abilities induced by α-syn (Supplementary Fig. 1A). Among these, we found that mino RNAi expression in α-syn flies (nSyb-GAL4/+; UAS-SNCA.J7/+;) is the only one that could alleviate their climbing phenotype for up to 8 weeks post-eclosion (Fig. 1B). This phenomenon is similarly observed when we employed another fly synucleinopathy model expressing α-syn in a pan-neuronal manner (nSyb-QF2 QUAS-SNCA/+) that triggers more severe phenotype (Fig. 1C). Conversely, overexpressing mino using an upstream UAS insertion mutant EY00734 exacerbates α-syn-induced loss of locomotor abilities (Fig. 1C). Collectively, these results underscore mino's role in modulating α-syn neurotoxicity. As due diligence, we also ascertained the expression of mino in both the RNAi and overexpression lines using qPCR of brain-extracted mRNAs (Supplementary Fig. 1B).

Pan-neuronal overexpression of α-syn is known to be toxic to DA neurons ⁴². Among the clusters of DA neurons in the fly brain, the protocerebral anterior medial (PAM) cluster is one of the largest clusters that contain more than 100 tyrosine hydroxylase (TH)-positive DA neurons. PAM cluster is involved in startle-induced negative geotaxis ^43,44. To investigate the role of mino in modulating α-syn-induced DA neurodegeneration, we utilized the Labkit pixel classification method to automatically quantify the PAM dopamine neuron cluster (Fig. 1D and 1E, Supplementary Fig. 1E) ⁴⁵. In the first group of α-syn mutant flies (nSyb-GAL4/+; UAS-SNCA.J7/+;) tested, we observed a significant decrease in PAM TH + cells at day 49, but not day 32, post-eclosion, when compared to nSyb-GAL4/+ control flies. Nonetheless, we found that the observed neuronal loss is rescued in the presence of mino RNAi (Fig. 1D). In the second group of α-syn mutant flies that has more severe phenotype, a decline in TH + cells is already evident at day 32 post-eclosion compared to their nSyb-QF2/+ control counterparts. Again, mino RNAi mitigates the neurodegeneration, although more modestly so (Supplementary Fig. 1C). The beneficial effects of mino RNAi persisted when we elevated the housing temperature to 29°C, which enhances the expression of both α-syn and mino RNAi (Fig. 1F). As expected, mino overexpression triggers the opposite effects, i.e. it exacerbates the loss of TH + cells in α-syn flies (Fig. 1F). Although α-syn expression alone resulted in an overall decrease in TH + cells during the pre-adult stage, we ascertained that mino's impact on α-syn-associated neurodegeneration is manifested at the adult but not pre-adult stage, as evidenced by the absence of significant differences in TH + cells in α-syn flies at day 1 post-eclosion in the presence of mino RNAi or overexpression (Supplementary Fig. 1D). Thus, mino is likely acting on α-syn-related pathways in a post-developmental fashion.

Studies on α-syn PD models have revealed disruptions in circadian rhythm-linked locomotor behavior ^46–48, which may reflect sleep disturbances in human PD. Consistent with these studies, we found that α-syn expression in flies induces an age-dependent decline in their daily activities, i.e. the disappearance of the morning peak during the dark-light transition and a reduced amplitude at the evening peak at the light-dark transition (Fig. 2A and 2B). This disruption in the circadian rhythm-related locomotor activities in α-syn is alleviated in the presence of mjno RNAi (Fig. 2A and 2B). However, although mino RNAi restored the evening peak's average intensity, it did not lead to the recovery of the morning peak (Fig. 2C-E). In contrast, mino overexpression tends to aggravate α-syn-induced reduction of the evening peak (Fig. 2A-E). In the absence of α-syn expression, neither mino RNAi nor its overexpression affects the daily total activities of Drosophila (Fig. 2A-E).

We recognize that the metabolic crosstalk between neurons and glia significantly influences neuronal energy balance. To investigate whether the modifying effects of mino on α-syn operates selectively in neurons or extends to glial cells, we downregulated mino expression either solely in glia (with repo-GAL4) or simultaneously in neurons and glia (with nSyb-GAL4 and repo-GAL4) and assessed their impact on the fly’s daily locomotor activities. We found that expressing mino RNAi in glia alone has no credible α-syn-modifying effects whereas simultaneous mino RNAi expression in neurons and glia mitigates the decline in daily locomotor activities exhibited by α-syn flies (Supplementary Fig. 2A and 2B). Thus, mino seems to exert its effects on α-syn in a cell-autonomous fashion.

Taken together, our results identified mino as a novel modifier of α-syn whose downregulation can alleviate α-syn-induced phenotypes in two distinct Drosophila models of α-syn-associated PD.

mino modulates α-syn-induced lipid peroxidation, 4-HNE accumulation and associated cell death

α-syn neurotoxicity is associated with alterations in lipid profiles, elevated lipid peroxidation and sensitivity towards ferroptosis ^{17,26,27,49,50}. Given that mino facilitates the acylation of acyl-CoA to glycerol-3-phosphate, a pivotal step in synthesizing phospholipids and triglycerides, we were curious to examine whether mino could impact on α-syn-associated lipid peroxidation. To investigate this, we labeled neuronal membranes with a cell membrane-localized infrared fluorescent protein (mIFP) whose pan-neuronal expression is directed by nSyb-GAL4 ⁵¹, and used BODIPY C11 lipid dye to analyze lipid peroxidation levels. Under standard culture conditions (25°C for 3 weeks), we observed that α-syn expression did not appreciably alter neuronal membrane lipid peroxidation levels, although the extent of lipid peroxidation decreased in the presence of mino RNAi and increased when mino is overexpressed (Fig. 3A, 3B and 3C). In the absence of α-syn expression, neither mino RNAi nor its overexpression alone affects the basal lipid peroxidation level (Fig. 3C). When flies were raised at 29°C for 3 weeks, α-syn overexpression alone significantly elevated the level of lipid peroxidation in neuronal membranes (Fig. 3A and 3C). This elevated lipid peroxidation level is significantly blunted by mino RNAi expression but becomes aggravated in the presence of mino overexpression (Fig. 3C). As lipid peroxidation promotes the buildup of products such as 4-hydroxynonenal (4-HNE) and malondialdehyde (MDA), we probed and found an elevated presence of the lipid peroxidation product 4-HNE in α-syn-expressing fly brains (Fig. 3D and 3E). As expected, the observed 4-HNE level is suppressed in the presence of mino RNAi expression and enhanced when mino is overexpressed (Fig. 3D and 3E).

Next, we investigated whether mino's effects on α-syn-induced lipid peroxidation correlates with changes in cell death. By means of TUNEL assay, we found that α-syn expression results in marked cell death compared to its control counterparts, especially at week 2 and 3 but not appreciably at earlier timepoints (day 1 and week 1) post-eclosion (Fig. 3F, 3G, and Supplementary Fig. 3A). Consistent with the results above, α-syn-induced cell death is significantly reduced in the presence of mino RNAi and aggravated when mino is overexpressed (Fig. 3F, 3G, and Supplementary Fig. 3A). Neither mino RNAi or its overexpression alone induces significant cell death in flies (Fig. 3F, 3G, and Supplementary Fig. 3A). To distinguish between apoptotic and non-apoptotic event, we co-stained the brains with cleaved caspase-3 antibody but did not detect positive staining (Fig. 3F). To confirm the antibody's efficacy, we stained hs-hid (heat shock- head involution defective) third instar larva brain (which expresses hid upon heat shock) after a 2-hours heat shock at 37°C and observed the expected strong positive staining (Supplementary Fig. 3B). Despite the possibility that cleaved caspase-3 might be detectable only during specific stages of apoptosis, the absence of its staining at weeks 2 and 3 in α-syn flies suggests that apoptosis may not be primarily occurring in α-syn-expressing fly brains.

A common role of GPAT as modulators of α-syn neurotoxicity

mino encodes an enzyme that belongs to the conserved GPAT gene family that are responsible for catalyzing the initial acylation step in the de novo synthesis of phospholipid ⁵². This prompted us to examine whether other GPAT members may play similar roles to mino in modulating α-syn neurotoxicity. To address this, we extended our RNAi targeting to include two other Drosophila homologs of GPAT, i.e. Gpat4, and CG15450. Whereas mino is predicted to reside on the mitochondria, the product of both Gpat4, and CG15450 are predicted to reside on the ER ⁵³. Similar to mino RNAi, downregulation of the expression of Gpat4, and CG15450 effectively suppresses the decline in startle-induced climbing abilities (Fig. 4A) and also mitigates the reduction in daily locomotor activities in α-syn flies (Fig. 4B and 4C). Aligning with these observations, Gpat4, and CG15450 RNAi expression also attenuates the decline of DA neuronal number in the PAM cluster induced by α-syn (Fig. 4D). Further supporting a common role for GPAT in modulating α-syn-induced toxicity, we found that when α-syn flies were treated with the GPAT inhibitor FSG67 ⁵⁴, they showed improvement in their daily locomotor activities (Fig. 4E and 4F). In contrast, RNAi-mediated silencing of Gnpat expression, a related enzyme predicted to reside on the peroxisome that is responsible for catalyzing the initial acylation step of ether lipid synthesis, did not significantly affect the locomotion activities of α-syn flies (Fig. 4A-C). Surprisingly, Gnpat RNAi reduces the loss of DA neurons in the PAM cluster (Fig. 4D). Despite this unexpected finding, these results collectively highlight mino as a representative of the broader GPAT activity responsible for modulating α-syn neurotoxicity.

GPAT RNAi suppresses α-syn higher-order oligomers formation and mitochondrial ROS accumulation

We have previously demonstrated that α-syn expression in the Drosophila brain leads to a metabolic shift from oxidative phosphorylation to glycolysis, a phenomenon that is regulated by hypoxia-inducible factor HIF1A and the kinase module component of the mediator complex Med13 ⁴¹. We were curious whether mino would influence this metabolic reprogramming induced by α-syn. Unlike HIF1A and Med13, we did not observe significant alterations in the α-syn-induced increase of glycolysis enzymes Pgi and Ldh in the presence of mino RNAi or other GPAT RNAi tested (Fig. 5A). Moreover, we found no credible evidence of changes in the total α-syn level upon mino/GPAT RNAi co-expression, suggesting that mino is likely exerting its effects downstream of α-syn, potentially impacting on α-syn aggregation and associated mitochondria-related oxidative stress level.

Several studies have highlighted the involvement of phospholipid membranes and cardiolipin, a mitochondrial lipid, in initiating the early stages of α-syn misfolding and aggregation ^55,56. Given that GPAT catalyzes crucial early steps in glycerolipid synthesis, we investigated whether GPAT influences the formation of α-syn oligomers/aggregates. In contrast to many α-syn-based PD models employing mammalian cells, α-syn expressed in the Drosophila brains is predominantly detected as monomers in both soluble and insoluble fractions ^42,57. However, oligomeric species of α-syn can be revealed through removing Triton X-100 and DTT from buffers while using cross-linkers like Dimethyl dithiobispropionimidate (DTBP) ⁵⁸. Supporting this, we observed that α-syn oligomers can indeed be stabilized in DTT- and Triton X-100-free buffers; the levels of which increases with increasing DTBP concentrations of up to 5 mmol, thereafter a sharp decline follows (Supplementary Fig. 4A and 4B). Using this method of detection for α-syn oligomers, we found that mino/GPAT RNAi, but not Gnpat RNAi, suppress the formation of higher-order α-syn oligomers in the presence of 4 mmol DTBP (Fig. 5B and 5C). Interestingly, GPAT RNAi specifically reduces α-syn tetramers and higher-order oligomers, without impacting α-syn dimers or trimers, either in comparison to their respective monomers or in total α-syn levels (Fig. 5B and 5C).

The formation of α-syn oligomers and aggregates is known to correlate with the generation of mitochondrial reactive oxygen species (ROS) including H₂O₂ ⁵⁵. To assess mitochondrial H₂O₂ levels, we utilized the fluorescent H₂O₂ sensor mito-roGFP2-Orp1 ⁵⁹. Although mino RNAi or overexpression did not significantly affect the mitochondrial H₂O₂ index in the absence of α-syn, their effects are evident in α-syn-expressing neurons, where mino RNAi results in a significant reduction of the H₂O₂ index, whereas mino overexpression exacerbates it (Fig. 5D and 5E). Alongside this, we employed MitoTimer, a sensor that reflects the cumulative redox history of mitochondria, to examine mitochondria maturation in these flies ⁶⁰. We observed a dramatically accelerated mitochondria maturation process in wild-type brains expressing α-syn, a phenomenon that is mitigated by mino RNAi (Fig. 5F and 5G). We also noted a minor suppression of mitochondria maturation by mino RNAi in the absence of α-syn (Fig. 5F and 5G).

Taken together, our results above revealed a connection between GPAT and the formation of α-syn oligomers and associated mitochondrial maturation/H₂O₂ generation, an important link that was not previously reported that may have pathophysiological relevance to α-syn-related PD pathogenesis.

Pharmacological inhibition of GPAT suppresses α-syn-induced lipid peroxidation in mouse primary neurons treated with α-syn preformed fibrils (PFF)

To examine the relevance of our findings above in the mammalian context, we investigated whether GPAT inhibition could similarly retard the formation of pathological α-syn species and the events they trigger in primary mouse neurons. To address this, we treated primary mouse neurons with α-syn preformed fibrils (PFF). In the presence of PFF treatment, we observed the appearance of elongated α-syn fibers that stain prominently with antibody against α-syn phosphorylation at serine 129 (pS129) (Fig. 6A and 6B). These phosphorylated α-syn fibers form dense clusters in both neurites and cell bodies of the PFF-treated neurons (Fig. 6A and 6B). Neither such morphology nor pS129 α-syn staining was observed when the primary neurons were treated with α-syn monomers (Fig. 6A and 6B). Importantly, co-treatment of neurons with PFF and the GPAT inhibitor FSG67 significantly attenuated the formation of p129 α-syn fibers and clusters (Fig. 6A and 6B).

Interestingly, PFF-induced pS129 fiber/aggregation colocalizes with the lipid peroxidation product 4-HNE and PFF-treated neurons exhibit elevated levels of both 4-HNE and MDA (Fig. 6A and 6B). Moreover, these neurons show excessive BODIPY 488 background staining, which is absent in in mock or monomers-treated neurons, suggesting the presence of lipid-related abnormalities in PFF-treated neurons. These molecular phenotypes induced in neurons by α-syn are all ameliorated in the presence of FSG67 co-treatment (Fig. 6A and 6B). Finally, we also stain PFF-treated neurons with BODIPY C11 to detect for lipid peroxidation. As expected, neurons treated with PFF, but not α-syn monomers, exhibit enhanced lipid peroxidation levels that are significantly reduced when they are co-treated with FSG67 (Fig. 6C).

Collectively, our results suggest that GPAT are also key players in α-syn-induced cytotoxicity in the mammalian context and that their modulation would similarly offer benefits.

In this study, we identified mino, alongside members of the GPAT family, as novel genetic modifiers of α-syn-induced neurotoxicity. Using both Drosophila and primary mouse neuronal model of synucleinopathy, we demonstrated that pharmacological or genetic reduction of mino/GPAT activity effectively mitigates α-syn-associated pathological phenotypes that includes a reduction in the neuronal lipid peroxidation level. These new insights that we have provided in this report support a key role for lipid-related pathways in the pathogenesis of PD and therein offers a therapeutic angle focused on lipid aberrations in PD.

α-syn comprises of an amphipathic N-terminal region, a hydrophobic non-amyloid-component (NAC) domain, and an acidic C-terminal domain. The N-terminal and NAC regions house multiple lipid membrane binding motifs critical for the protein's interaction with lipid membranes ^19,61. α-syn aggregation is enhanced in the presence of phospholipid surfaces ^62–64 and its toxicity can also be altered by the saturation level of fatty acids ²⁸. In turn, α-syn can trigger various lipid abnormalities including variations in neutral and polar lipid levels, ester and ether lipids, unsaturation levels of fatty acids and lipids, division of specific fatty acid species in the fatty acid pool and lipid pools, and lipid peroxidation level. Using two Drosophila PD models exhibiting mild or strong disease phenotypes, we demonstrated that α-syn neurotoxicity can be modulated by the lipid synthesis enzyme mino. RNAi against mino ameliorates α-syn-associated pathological phenotypes including the decline in locomotor activities and the loss of DA neurons, while elevated mino expression exacerbates these phenotypes. The beneficial effects of mino downregulation are similarly observed in α-syn flies harbouring RNAi species targeting the predicted differentially subcellular-localized Gpat4, and CG15450, suggesting a common role of GPAT in modulating α-syn neurotoxicity. Notably, previous report indicated that fluorescence-tagged transgenic mino, when expressed in the female reproductive system, colocalized with both mitochondria and ER ⁶⁵. Although we detected mitochondrial localization of ectopically expressed α-syn in Drosophila photoreceptor neurons (data not shown), it is plausible that dynamic lipid exchange between mitochondria and ER contributes to the subcellular-independency of the modifier role of GPAT ⁶⁶.

Our study also indicates that the modulation of higher-order oligomers of α-syn can potentially be achieved by employing RNAi targeting GPAT. The detection of α-syn oligomers in Drosophila brains through western blot has historically posed greater challenges compared to mammalian cells, despite successful identification of ectopically expressed α-syn in both TritonX-100 soluble and insoluble fractions ^42,57,67. Through an in vivo crosslinking with DTBP originally described by Cole and colleagues ⁵⁸, we were able to enhance the sensitivity of detecting α-syn oligomeric species. While RNAi against GPAT did not result in a significant change in α-syn protein levels, we observed a relative redistribution of α-syn from higher-order oligomers to lower-order oligomers and monomers. This shift was associated with a decrease in α-syn neurotoxicity.

Interestingly, we observed a significantly accelerated mitochondria cumulative redox level in pan-neuronal α-syn-expressing fly brains during the first week, that preceded the onset of cell death from the second week onward. Although elevated α-syn is known to trigger caspase activation and apoptosis in various PD models ^68,69, it remains uncertain if α-syn-induced neuronal death can occur through alternative pathways. Recent observations of enhanced lipid peroxidation in neurons containing SNCA gene triplication suggest their potential sensitivity to ferroptosis ²⁶. In the enzymatic ferroptotic pathway, lipoxygenase (LOX) catalyzes iron-dependent specific oxidation of poly-unsaturated PE-AA and PE-AdA ⁷⁰. Mitochondria, the primary ROS generating organelle, contributes to the non-enzymatic pathway of ferroptosis via Fenton reaction ⁷¹. Drosophila, with a distinct lipidomic composition lacking polyunsaturated fatty acids including AA and AdA, may utilize shorter chain polyunsaturated fatty acids, such as linoleic acid (18:2n-6) and linolenic acid (18:3n-3), as substrates for ROS-induced non-enzymatic lipid peroxidation on ER and mitochondria ^72,73. Linoleic acid in choline glycerophospholipids is prone to form hydroperoxide during peroxidation in human erythrocytes membrane ⁷⁴.The lipid peroxidation product 4-HNE has been observed in fat body, neurons, and muscles of aging flies and was further enhanced by paraquat feeding ⁷⁵. Although our results favor a model by which α-syn elicits neuronal death through ferroptosis, we cannot rule out apoptosis as a co-driver of neurodegeneration in this case. We appreciate that the TUNEL assay, which is commonly used for cell death detection, lacks specificity in distinguishing apoptosis, necrosis, and autolytic cell death ⁷⁶, notwithstanding that TUNEL-positive nuclei have also been identified in cells undergoing ferroptosis ⁷⁷. Moreover, despite the absence of cleaved caspase 3 in our study, previous study using an hPARP-based apoptosis reporter has detected apoptosis in pan-neuronal α-syn-expressing brains in Drosophila ⁴². It is therefore plausible that both apoptosis and ferroptosis coexist to drive neurodegeneration in α-syn fly brains ⁷⁸.

Finally, we demonstrated in both the α-syn fly as well as primary mouse neuronal models that pharmacological inhibition of mino/GPAT activity through FSG67 treatment promotes beneficial outcomes that illustrates the transferability of our findings from non-mammalian to mammalian context. Notably, current disease-modifying experimental therapeutics for PD aimed at α-syn as a target are mostly focused on reducing its aggregation or enhancing its degradation. Our approach targeting a key pathway that α-syn acts on, i.e. lipid synthesis and modifications, offers a unique and at the same time complementary therapeutic angle that could augment the anti-aggregation, and/or pro-degradation approaches in mitigating α-syn-induced neurotoxicity. Interestingly, FSG67 was originally developed as a drug to treat obesity and diabetes ⁷⁹. Given the optimism surrounding the use of GLP agonists to treat PD, which was similarly first developed to treat diabetes (and subsequently to manage weight loss), it is interesting to postulate a converged pathway between FSG67 and GLP agonists that emphasizes the need to better understand the role of lipid aberrations in the pathogenesis of PD.

In conclusion, our research not only reveals a pivotal role of glycerol 3-phosphate acyltransferase in modulating α-syn aggregation and its neurotoxic effects, but also highlights its potential as a novel target for mitigating the progression of PD.

Drosophila stocks

Flies were grown at 25°C with a standard yeast-corn meal-dextrose diet. Aging experiments were conducted at 25°C, unless otherwise stated. Adult male flies were gathered in groups of 20–30 per vial for aging. Throughout the aging process, the food was refreshed every two days for the initial two weeks and subsequently replaced daily from the third week onward. The Drosophila stocks used are listed in Supplementary Table 1.

Drosophila brain immunostaining

Larvae or adult fly brains were dissected in PBS, fixed in 3.7% paraformaldehyde in PBS with 0.1% Triton X-100 on ice for 0.5 hour, and subsequently fixed at room temperature for an additional 0.5 hour. Following fixation, brains were rinsed in PBS with 0.4% Triton X-100 (PBST) and subjected to primary antibodies staining for 1 day at 4°C, then washed in PBST and incubated in secondary antibodies for 1 day at 4°C, followed by washing with PBST. In the TUNEL assay, after removal of the buffer, the TUNEL reaction mixture was added, incubated in a 37°C water bath for 1h, and briefly washed. Subsequently, brains were stained in PBST with 4',6-diamidino-2-phenylindole (DAPI) for 3 hours, briefly washed, and mounted with Prolong Gold (Thermo Fisher Scientific) on bridged slides, positioned as a line with the anterior side facing upward and both sides padded with double layers of Scotch transparent tape (except for a single layer for TH staining in the PAM cluster). For cleaved caspase 3 staining, third instar larvae of w¹¹¹⁸ or hs-hid were heat-shocked in a 37°C water bath for 2 hours and allowed to recover for 5 hours before dissection. Imaging was performed using an FV3000 confocal microscope with consistent image settings for samples within each experiment. The primary antibodies used include rabbit anti-Tyrosine Hydroxylase TH (Pel Freeze, P40101-0, 1:500), rabbit anti-Cleaved Caspase-3 (Asp175) (Cell Signaling Technology, 9661, 1:500), rabbit anti-Cleaved Caspase-3 (Asp175) (5A1E) (Cell Signaling Technology, 9664, 1:500), mouse anti-Repo (DSHB, 8D12, 1:500), mouse anti-alpha-Tubulin (DSHB, 12G10, 1:5000), mouse anti-Malondialdehyde (Abcam, 11E3, ab243066, 1:500), mouse anti-4 Hydroxynonenal (Abcam, HNEJ-2, ab48506, 1:500), rabbit anti-α-syn (phospho S129) (Abcam, ab51253, 1:500).

Labkit-assisted Drosophila PAM cluster TH + neuron quantification

The TH + neurons within a single PAM cluster were imaged using FV3000 with 512x512 resolution and 0.34 µm slice interval for both TH and DAPI channels, generating z-stacks. Labkit training and TH + region prediction were performed on a computer featuring an NVIDIA Quadro P6000. Twenty representative z-stacks, encompassing a spectrum of TH signals and backgrounds, were vertically merged into a unified stack using Fiji/ImageJ. In Labkit (Fiji), the stack underwent more than 100 cycles of "scribbling-predicting-correcting" to train the TH + pixel classifier against backgrounds. The resulting classifier file with a size of 379 megabytes was applied consistently for subsequent TH + neuron analysis. For TH + region identification, the classifier file was loaded into Labkit to batch process a folder of z-stack TH images. The defects including noise and vacuoles on the Labkit-predicted mask images were further minimized using customized Matlab (Mathwork) code “Correct_Labkit_TH.m” (version 1.0, see the source code, instructions for installation and use, description of code dependencies and the test data in https://github.com/rmd13/FlyBrain_PAM_TH_DAPI_Labkit/ ). The code sequentially processes each slice of a Labkit-generated 0/1 mask image. It first smooths the mask using a Gaussian smoothing filter with a sigma of 1.5 to enhance the image quality. Following smoothing, a threshold of 0.25 is applied to the resulting image, thus only the pixels above this threshold are labeled as a new mask. This new mask slice is then dilated using a square-shaped structuring element with a width of 5 pixels. This step is crucial for filling in any small gaps or holes present within the TH + regions, ensuring that the mask is continuous. Additionally, the code identifies and addresses any larger holes that may still exist within the mask, provided these holes have an area of less than 200 pixels. These holes are filled to maintain the integrity of the mask. Finally, the mask is eroded using the same square-shaped structuring element with a width of 5 pixels. The above process helps to reduce the errors, resulting in a high-quality, refined mask suitable for subsequent image analysis steps. The processed mask images were imported into Imaris 8.4 (Bitplane) as a new channel alongside the raw images. TH + surfaces and DAPI spots were created using Imaris in the Labkit channel and DAPI channel, respectively. The number of TH + neurons in the PAM cluster was counted as DAPI spots located within the TH + surface by running the surface distance transformation extension (Imaris) outside the TH + surfaces.

Drosophila brain BODIPY C11 lipid peroxidation assay

Fly brains expressing pan-neuronal infrared fluorescent protein (mIFP) were dissected in PBS and transferred into in Eppendorf tube 1 containing 100 ul PBS. In Eppendorf tube 2, 0.5 ul of 5 mmol BODIPY C11 DMSO solution (Thermo Fisher Scientific, D3861) and 0.5 ul of 1% saponin (Sigma, S7900) were added into 350 ul PBS (B), mixed and transferred to the tube 1, which was then rolled at room temperature for 30 min. Following the rolling process, brains were briefly washed in PBS and mounted with PBS on a bridged slide using two layers of Scotch transparent tape, creating a 5 mm gap. Thick nail polish was used to secure the four corners of the coverslip, and additional PBS was added to fill any gaps if necessary. BODIPY C11 was immediately imaged in FV3000 using a 100x objective lens. The brain optic lobe was located and live-imaged using a 640 nm laser to determine the position of labeled neuronal membranes. Subsequently, images were acquired in the green channel (488 nm laser, 0.04% laser power, 515–545 nm emission) and red channel (561 nm laser, 0.02% laser power, 575–605 nm emission), respectively. Five images were acquired for each brain. For each group the brains were imaged within 1 hour. The lipid peroxidation index was analyzed as the ratio of the mean intensity of membrane area in the green channel to the mean intensity in the red channel using ImageJ/Fiji. For each brain, the lipid peroxidation index from five images was averaged. A pseudo-colored image was generated by creating a ratio channel and masking out the unstained area in Imaris.

Drosophila daily locomotor activity assay

Fly food, extracted from a vial, was trimmed into two semi-circles with a height of 5 mm, and then positioned on 3 layers of 5 cm x 5 cm tissue paper with smooth side up for a 0.5 hour drying period. To administer FSG67 (HY-112489, MedChemExpress LLC), 10 µl of 0 mmol,1 mmol, 2.5 mmol, or 5 mmol FSG67 solution in PBS was introduced to the smooth side of fly food and evenly distributed until complete absorption. One end of clean glass tube for locomotor activity assay was inserted down vertically into the food and capped. Anesthetized enclosed flies were individually transferred into each tube, and then the tube was blocked with a sponge on the opposite side. These loaded glass tubes were then affixed to the DAM2 Drosophila activity monitors (TriKinetics). The locomotor activities were recorded at a frequency of 10 s over a period of one to three weeks, maintaining a 12-hour light/dark cycles of LED illumination controlled via an Arduino-controlled relay by the same computer. The food was replaced every three days. Actograms captured during this period were analyzed using the ActogramJ plugin ⁸⁰ in ImageJ/Fiji. Daily locomotor activities were further analyzed and plotted using Excel 2010 and GraphPad Prism 6.01.

Western blot

Fly brains were dissected in PBS and homogenized with pestles in brain lysis buffer (Tris-HCl, pH 8.0 10 mmol, NaCl 200 mmol, Triton X-100 1%, EDTA 5 mmol, Glycerol 5%, PMSF 0.5 mmol, DTT 1 mmol, protease inhibitor cocktail (Roche, 11873580001)). Subsequently, an equal volume of 2X laemmli loading buffer (Tris-HCl, pH 6.8, 125 mmol, bromophenol blue 0.02%, SDS 4%, glycerol 20%, β-mercaptoethanol 10%) was added and heated at 50^oC for 0.5 hour. For detection of α-syn oligomers, fly brains were dissected in PBS and cross-linked in same-day prepared buffer A (HEPES 20 mmol pH 8.5, KCl 120 mmol, EDTA 2 mmol, sucrose 0.25 M, glycerol 1%, Dimethyl dithiobispropionimidate (DTBP) 4 mmol, protease inhibitor cocktail (Roche)) on ice for 1h. Then reaction was halted by adding 10 mmol Tris-HCl pH 8.8 for 10 min on ice. Brains were homogenized with a pestle, and an equal volume of 2X Laemmli loading buffer was added without β-mercaptoethanol or DTT, followed by heating at 50^oC for 0.5 hour. The samples were loaded into Mini Protean TGX (stain-free) 4–20% gel (Bio-Rad, 4568095) to run for 100V 1.5 hour. The gel was UV-activated and transferred onto PVDF membrane using transfer buffer (Tris 25 mmol, Glycine 192 mmol, 20% methanol) at 100V for 1.5 hour. The membrane was fixed in PBS with 3.7% PFA and 0.025% glutaraldehyde for 0.5 hour, stain-free imaged, and blocked in 5% milk in Tris-buffered saline with 0.1% Tween-20 (TBST). Primary antibodies used include mouse anti-α-syn (BD 610787, 1:1000), rabbit anti-Gpi (Merck, HPA024305, 1:1000), rabbit anti-LDH (Thermo Fisher,PA5-26531, 1:1000), and secondary antibodies include anti-mouse HRP (CST, 7076S, 1:3000) and anti-rabbit HRP (Amersham, NA934-1ML, 1:3000). The HRP was detected using WesternBright Sirius in ChemiDoc (Bio-rad). The membrane was subsequently re-blotted with mouse anti-alpha tubulin (DSHB, 12G10, 1:5000). Image was analyzed using Image Lab (Bio-Rad) and ImageJ/Fiji, and the results were plotted in GraphPad prism.

Mito-roGFP2-Orp1 imaging for H₂O₂

Retina expressing mito-roGFP2-Orp1 were dissected in PBS and fixed in PBS with 3.7% PFA, 0.1% Triton X-100 and 20 mmol N-Ethylmaleimide (Thermo Fisher Scientific, 23030) for 0.5 hour on ice, followed by another 0.5 hour at room temperature. The fixed retina was washed in PBST with 5% goat serum overnight at 4^oC and then was mounted with Prolong Gold anti-fade mountant (Thermo Fisher Scientific, P10144). Mito-roGFP2-Orp1 was imaged by exciting with a 405 nm or 488 nm laser, respectively, and collecting 500–530 nm emission light. The H₂O₂ index was determined by the ratio of mean intensity of the mitochondria in the two channels using ImageJ/Fiji.

MitoTimer imaging for cumulative redox history

Brains were dissected in PBS and fixed in PBS with 3.7% PFA and 0.1% Triton X-100 for 0.5 hour on ice and an additional 0.5 hour at room temperature. Following fixation, the brains were washed in PBST with 5% goat serum and DAPI overnight at 4^oC and then mounted with Prolong Gold anti-fade mountant (Thermo Fisher Scientific, P10144). The green channel (488 nm laser, 500–540 nm emission) and the red channel (561 nm laser, 570–620 nm emission) were imaged on the cortex layer of the optic lobe. The ratio of the mean intensity of MitoTimer between the red and the green channel was analyzed using ImageJ/Fiji.

Drosophila climbing assay and quantification

The climbing assay was conducted on a customized workstation composed of a background white foam, side LED light, bottom light, 50 ml glass cylinders, an elastic cushion, and a Nikon digital camera securely affixed to a tripod. A sponge pad was positioned in the inner bottom of the cylinder to minimize potential harm to the flies during the climbing assay. 10 ~ 20 flies were flipped into a capped cylinder, tapped to the bottom and allowed to climb for 5 s, following which an image was promptly captured using the camera. The average climbing height for all the images was analyzed using a customized FlySpotter program (Srinivas Gorur-Shandilya, https://github.com/sg-s/fly-spotter) and graphically represented using GraphPad prism.

Real-time PCR for quantitative of fly brain transcripts

Fly brains of 3-weeks old were dissected in PBS, with trachea sacs and fat bodies detached, were subsequently transferred into RNAlater (Thermo Fisher Scientific, R0901). Following removal of excess fluid, RNA from the brain was extracted using TRIzol™ Reagent (Thermo Fisher Scientific, 15596026). The extracted RNA was treated with dsDNase (Thermo Fisher Scientific, EN0771). Subsequently, cDNA was prepared using Invitrogen SuperScripII First-Strand Synthesis System for RT-PCR (11904-018). qPCR was performed using iQ™ SYBR Green Supermix (Bio-rad, 1708882) and LightCycler 96 Instrument (Roche). The data was analyzed using LightCycler 96 software (Roche). RPL32 served as an internal control for mRNA quantification. The qPCR primers used include mino-F1: CAAGCCATCTGCCAGGGTCC, mino-R1: GACATCACAGAGGTGGCTGCTG, RpL32-qF1: ATGCTAAGCTGTCGCACAAATG, RpL32-qR1: GCTTGTTCGATCCGTAACCG.

Synuclein Preformed fibrils preparation

15µl of 2mg/ml synuclein PFF (StressMarq SPR-324) aliquoted into screw-capped tubes were sonicated in an ultrasonic bath sonicator for 2.5h at the following conditions: Sweep, 37Hz frequency, 60% power. Water was changed at 30mins interval to maintain the water temperature at 15°C.

Mouse primary neuron culture and staining

Mice-related studies were approved by and conformed to the guidelines of the Institutional Animal Care Committee (IACUC) of the Nanyang Technological University Lee Kong Chian School of Medicine. Embryonic day 17.5 mouse fetuses from C57BL/6 were obtained. The cortices were isolated and dissociated with papain (LS003119, Worthington) for 30 min at 37°C. Papain was removed and cortices were washed with DMEM/FBS and spin down at 1,000rpm, 5 mins. Cells were titrated, spin down and resuspended in Neurobasal medium supplemented with B27 and GlutaMAX (Gibco). 80,000 cells/ well were plated. At day 7, mice primary cortical neurons were treated with 1µg/ml α-syn monomers (SPR-323, StressMarq) or 1µg/ml synuclein PFF (SPR-322, StressMarq) with or without 75 mmol FSG (HY-112489, MedChemExpress LLC) for 2 weeks. Following treatment, cells were fixed in 4% PFA/4% sucrose/PBS for 30 minutes at room temperature, then washed three times with PBS. Subsequently, cells were blocked in 0.2 M glycine, 0.1 mg/ml saponin, and 30 mg/ml BSA in PBS for 45 minutes. Primary antibodies staining was carried out in antibody diluent (0.1 mg/ml saponin and 1 mg/ml BSA in PBS) for 1 hour at room temperature using anti-Malondialdehyde antibody (Abcam, 11E3, ab243066, 1:500), anti-4-Hydroxynonenal antibody (Abcam, HNEJ-2, ab48506, 1:500) and anti-α-syn phospho S129 (Abcam, ab51253, 1:500). After washing three times with PBS, cells were incubated with secondary antibodies in antibody diluent for 1 hour, followed by four washes with PBS. Finally, cells were stained with BODIPY 493/503 (Thermo Fisher Scientific, D-3922, 1:1000) and 1 µg/ml DAPI, mounted, and imaged using an FV3000 confocal microscope (Olympus). For lipid peroxidation assay, 5 mmol BODIPY C11 was added to the culture medium of glass-bottom culture dish in 1:1000 dilutions and incubated for 0.5 hour, then directly imaged in the green and red channel respectively. The ratio of mean intensity in the cell was analyzed using ImageJ/Fiji.

Statistics

Statistical analyses were conducted using Prism 8 (GraphPad) and Matlab (MathWorks). For the genetic screening, the Anderson-Darling test was applied to assess the non-normal distribution of all the data, followed by the Mann-Whitney U-test. The p-values were adjusted by the Benjamini-Hochberg method using the Matlab function padj = PVAL ADJUST (p-values, ‘BH’), where p is the input vector of all the p-values (fakenmc/pval_adjust, version 1.2.0.2, by Nuno Fachada. https://github.com/nunofachada/pval_adjust). One-way or two-way ANOVA, supplemented by Tukey’s and Dunnett’s post hoc test, was used for multi-group comparisons. All tests were two-tailed. Sample sizes are specified in the figure legends. Data are presented as mean ± S.D. Significance levels are *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Data Availability

All the data is included in this manuscript and supplementary data.

Code Availability

The Labkit classifier file for PAM cluster neuron prediction and the code for the correction of Labkit-predicted TH+ regions could be downloaded from Github (https://github.com/rmd13/FlyBrain_PAM_TH_DAPI_Labkit). The code was under MIT license.

ACKNOWLEDGEMENTS

This work was supported by Singapore Ministry of Health Open Fund-Large Collaborative Grant (MOH-OFLCG000207) to K.L.L.

AUTHOR CONTRIBUTIONS

M.R. and K.L.L. participated in the design of experiments, data analysis, and interpretation. W.T and G.G.Y.L. designed and performed the experiments using primary mouse neurons. M.R. and K.L. wrote the manuscript with inputs from all the authors.

CONFLICT OF INTERESTS

The authors declare no competing interests.

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Identification of Glycerol 3-phosphate acyltransferase as a potent modifier of α-Synuclein-induced toxicity

Status:

Version 1

Abstract

Figures

HIGHLIGHTS

INTRODUCTION

RESULTS

A common role of GPAT as modulators of α-syn neurotoxicity

GPAT RNAi suppresses α-syn higher-order oligomers formation and mitochondrial ROS accumulation

DISCUSSION

MATERIALS AND METHODS

Western blot

Mito-roGFP2-Orp1 imaging for H₂O₂

MitoTimer imaging for cumulative redox history

Real-time PCR for quantitative of fly brain transcripts

Synuclein Preformed fibrils preparation

Mouse primary neuron culture and staining

Statistics

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

Identification of Glycerol 3-phosphate acyltransferase as a potent modifier of α-Synuclein-induced toxicity

Status:

Version 1

Abstract

Figures

HIGHLIGHTS

INTRODUCTION

RESULTS

A common role of GPAT as modulators of α-syn neurotoxicity

GPAT RNAi suppresses α-syn higher-order oligomers formation and mitochondrial ROS accumulation

DISCUSSION

MATERIALS AND METHODS

Western blot

Mito-roGFP2-Orp1 imaging for H2O2

MitoTimer imaging for cumulative redox history

Real-time PCR for quantitative of fly brain transcripts

Synuclein Preformed fibrils preparation

Mouse primary neuron culture and staining

Statistics

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1

Mito-roGFP2-Orp1 imaging for H₂O₂