The SAC1 domain in synaptojanin is required for autophagosome maturation at presynaptic terminals

The Parkinson's disease mutation R228Q in the synaptojanin 1 SAC1 domain does not affect synaptic vesicle endocytosis at fly excitatory glutamatergic neurons and photoreceptors

SYNJ1 is evolutionarily conserved from yeast to humans (Fig 1A and B). The human Parkinson's disease‐causing R258Q mutation in SYNJ1 blocks the dephosphorylation of specific phosphoinositides including PI(3)P and to assess the functional consequences of this mutation, we used MiMIC technology and knocked in the R228Q mutation (corresponding to the human R258Q) into the endogenous Drosophila synj locus (synj^RQ) (Appendix Fig S1A) (Venken et al, 2011; Vilain et al, 2014). This synj^RQ knock‐in allele expresses Synj at a comparable level to the wild‐type synj locus, indicating that the mutation does not affect protein stability (Appendix Fig S1B, C, and H). We also developed a second system to study Synj function, where we use the UAS/Gal4 system to transgenically overexpress wild‐type or R228Q Synj in neurons of synj null mutant flies that otherwise lack Synj expression (synj^−/−; nSyb > UAS synj⁺ or synj^RQ). We confirm similar expression of these transgenically expressed Synj⁺ and Synj^RQ proteins [~1.2‐fold over the levels produced from the endogenous synj locus (Appendix Fig S1D, E, and I)]. We compared the subcellular localization of endogenous Synj against that produced from the R228Q knock‐in locus, or transgenically expressed wild‐type Synj⁺ and Synj^RQ proteins. We find that all concentrate in the presynaptic terminals of the larval neuromuscular junction (NMJ) (Appendix Fig S1F–G″ and J–M″), indicating the mutant protein localizes correctly and similar to the presynaptic protein EndoA (Verstreken et al, 2002).

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Figure 1. The Parkinson's disease mutation R228Q in the Synj SAC1 domain does not affect synaptic vesicle cycling

• A. Diagram of human Synj1 indicating the two polyphosphoinositide domains (SAC1 and 5‐PPase) and the proline‐rich domain. The SAC1 domain bears at position 258 an arginine to glutamine amino acid change causative for PD.

• B. Arginine at position 258 (green) in Synj1 SAC1 domain and the surrounding region are highly conserved from yeast to human (R258 corresponds to R228 in Drosophila).

• C. Table indicating neuronal expression (nSyb‐Gal4) of Synj^RQ and Synj⁺ rescues the lethality of synj null mutants (synj^1/2 indicated as synj^−/−), “A” and “L3” indicate lethality at the “adult stage” and at the “third‐instar larval stage”, respectively.

• D. Example average traces of ERGs of control animals (left, nSyb‐Gal4/+), flies with synj null mutant eyes (synj^−/−), flies with synj null mutant eyes expressing synj^RQ or synj⁺ (UAS synj^RQ, synj^−/− and UAS synj⁺, synj^−/−), control animals (right, w¹¹¹⁸), and synj^RQ knock‐in animals. Full genotypes appear in the Appendix Supplementary Methods. “ON” and “OFF” peaks are indicated with arrowheads. Note that while synj null mutants display severe defects in their ERG traces, synj^RQ mutants do not. n > 20 animals per genotype.

• E–J Representative images of boutons loaded (1 min, 90 mM KCl) with FM 1‐43 (E–I) and quantification (J) of the following genotypes: control (left, D42‐Gal4/+) (E), synj null mutants (synj^−/−) (F), synj null mutants that express synj^RQ (G), or synj⁺ (H) (UAS synj^RQ, synj^−/− and UAS synj⁺, synj^−/−), control animals (right, w¹¹¹⁸; image not shown, only quantification is included) and synj^RQ knock‐in animals (I). Full genotypes appear in the Appendix Supplementary Methods. Scale bar, 5 μm. Note that synj^RQ internalizes as much FM 1‐43 as controls do. For UAS experiments, statistical analysis with one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, ****P < 0.0001; ns, not significant; n ≥ 7 larvae and n ≥ 14 NMJs per genotype. For synj^RQ experiment: t‐test, P > 0.05, n = 10 larvae and n = 20 NMJs per genotype. Error bars represent SEM.

• K. Quantification of the number of satellite boutons per NMJ of the genotypes in (E–I). Note that while synj null mutants harbor numerous satellite boutons, synj^RQ mutants do not. Statistical analysis for UAS experiments: one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, *P < 0.05, n ≥ 4 larvae and n ≥ 16 NMJs per genotype. For Synj^RQ experiment: t‐test, P > 0.05, n = 6 larvae and n ≥ 14 NMJs. Error bars represent SEM.

• L, M Quantification (L) of the EJC amplitude recorded at third‐instar larval NMJs of the indicated genotypes (see also E–I) and sample EJC traces (M). Statistical analysis, one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, P > 0.05, n ≥ 8 larvae per genotype. Error bars represent SEM.

• N. Cumulative histograms of mEJC amplitudes and sample EJC traces recorded at third‐instar larval NMJs of the indicated genotypes (see also E–I). n ≥ 7 larvae for indicated genotypes.

• O, P Relative EJP amplitudes recorded during a 10‐min 10‐Hz stimulation train of the genotypes indicated in (E–I). Please see Materials and Methods for details. Data for the synj null mutants that express synj^RQ or synj⁺ are not significantly different from control (D42‐Gal4/+), whereas a significant difference is observed from data point 3 till the end of the recording of synj null mutants (synj^−/−) compared to control (ns: data points 1–2, *P < 0.05: data point 3, **P < 0.01: data points 4, 16–21, ***P < 0.001: data points 5, 13–15, ****P < 0.0001: data points 6–12). Statistical analysis for UAS experiments: one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, n ≥ 4 larvae indicated genotypes in (O). Data for Synj^RQ compared to control are not significantly different. Statistical analysis by t‐test, P > 0.05, and n ≥ 8 larvae for indicated genotypes in (P). Error bars represent the SEM. Note that synj null mutants do not maintain neurotransmitter release whereas the synj^RQ mutants maintain release similar to controls.

Synj null mutant animals are not viable (Fig 1C), and deletion of synj specifically in the eye severely affects the efficiency of synaptic transmission between eye photoreceptors and the brain detected by electroretinogram (ERG) recordings (Fig 1D, arrowheads). As expected, these Synj null phenotypes are rescued by neuronal transgenic expression (nSyb‐Gal4) of wild‐type Synj⁺. Surprisingly, however, transgenic expression of the Synj^RQ mutant is as effective as the wild‐type protein (Fig 1C and D). Since even a partially active mutant might result in rescue of the phenotype when overexpressed, we confirmed that synj^RQ knock‐in flies are viable and that they too display normal ERGs (Fig 1D). Hence, it appears that the Synj^RQ mutant protein retains core Synj activities that are essential for viability and synaptic communication in the eye.

Synj is best defined as the protein that uncoats newly endocytosed synaptic vesicles (Cremona et al, 1999; Harris et al, 2000; Verstreken et al, 2003). To directly examine whether R228Q affects this endocytic function, we performed FM 1‐43 fluorescent dye labeling at the excitatory glutamatergic Drosophila NMJ. FM 1‐43 binds membranes and is incorporated into and labels newly formed synaptic vesicles, thereby labeling synaptic boutons proportional to the amount of vesicle recycling and thus synaptic endocytosis (Ramaswami et al, 1994). As previously described, synj loss significantly reduces FM 1‐43 labeling compared to controls (Verstreken et al, 2003). This labeling is restored by neuronal expression of wild‐type synj, and again also by Synj^RQ (Fig 1E–H and J). We confirmed that Synj^RQ rescues the synj knockout phenotype independent of overexpression by finding normal FM 1‐43 dye uptake in synj^RQ knock‐in animals (Fig 1I and J). Thus, FM 1‐43 labeling suggests that Synj^RQ supports presynaptic vesicle recycling and, under the conditions we tested, does so as efficiently as wild‐type Synj.

The disruption of NMJ endocytosis often correlates with morphological defects. These also occur in the synj null mutants in the form of numerous satellite boutons that are small protrusions emanating from the primary axial branch (Dickman et al, 2006). We therefore examined whether Synj^RQ possesses sufficient Synj activity to suppress this defect. We find a significant rescue of this synj null mutant‐associated satellite bouton phenotype in synj null mutants that transgenically express wild‐type Synj⁺ or mutant Synj^RQ. We also observe no difference in satellite bouton number between synj^RQ knock‐in animals and controls (Fig 1K). Hence, Synj^RQ appears to allow enough endocytosis to suppress this classic readout of NMJ endocytic defects, and is markedly different to the synj null allele.

We next directly examined how well Synj^RQ supports neurotransmission at the larval NMJ. First, we measured basal release characteristics including miniature excitatory junctional currents (mEJC) and EJCs. In controls and synj null mutants expressing Synj⁺ or Synj^RQ, no difference is observed in mEJC and EJC amplitude nor in frequency (Fig 1L–N). We then stimulated the motor nerves for 10 min at 10 Hz to continuously drive synaptic vesicle exo‐ and endocytosis, and measured the post‐synaptic excitatory junctional potentials (EJP). While control NMJs maintain neurotransmitter release under these conditions, synj null mutants fail to do so, and the block in endocytosis causes the EJP amplitude to drop to a low level in the first minutes of stimulation (Fig 1O) (Verstreken et al, 2003). This neurotransmission defect is rescued by transgenic neuronal expression of wild‐type Synj⁺, or Synj^RQ (Fig 1O). Here also, synj^RQ knock‐in animals maintain release during this 10‐Hz stimulation train at a level comparable to controls (Fig 1P). Synj^RQ was previously shown to lack SAC1 domain phosphatase activity (Krebs et al, 2013) and our data now suggest that the Synj^RQ mutant protein can support synaptic vesicle cycling. These results are consistent with data obtained in mouse cortical neurons using SAC1‐dead Synj1 mutants that also supported vesicle recycling during persistent activity (Mani et al, 2007).

Synaptojanin is required for autophagosome formation in presynaptic terminals

Given that Synj is strongly enriched at presynaptic terminals (Verstreken et al, 2003), Synj1^R258Q lacks PI(3)P phosphatase activity (Krebs et al, 2013), and the importance of PI(3)P on autophagosomes (Simonsen & Tooze, 2009; Burman & Ktistakis, 2010; Noda et al, 2010; Dall'Armi et al, 2013; Devereaux et al, 2013), we hypothesized that the Synj SAC1 domain acts in synaptic autophagy. We and others recently showed the presence of autophagic markers at presynaptic terminals (Williamson et al, 2010; Hernandez et al, 2012; Zhu et al, 2013; Maday & Holzbaur, 2014; Wang et al, 2015, 2016; Soukup et al, 2016; Stavoe et al, 2016). To determine whether Synj localizes on autophagic membranes, we examined the interaction between Synj and the transmembrane Atg9 marker of early pre‐autophagosomal membranes (He et al, 2006; Tamura et al, 2010; Yamamoto et al, 2012; Stavoe et al, 2016). The distribution of a genomic Atg9^HA construct where Atg9^HA is expressed under endogenous promotor control is broad (Appendix Fig S2A–A″), consistent with the idea that Atg9 decorates early autophagic vesicles and autophagosomes. Next, we used anti‐HA to immunoprecipitate neuronal autophagosomal membranes from fly heads expressing Atg9^HA at endogenous levels. We detect the co‐immunoprecipitation of Synj with the Atg9‐positive membrane fraction in both wild‐type and synj^RQ knock‐in animals expressing Atg9^HA (Fig 2A). In contrast and indicating specificity, other synaptic proteins (alpha‐SNAP and complexin) do not co‐immunoprecipitate (Fig 2A).

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Figure 2. Synaptojanin is required for synaptic autophagy

• A. Western blot of an anti‐HA immunoprecipitation from control fly heads (w¹¹¹⁸), Atg9^HA‐expressing fly heads (w;;Atg9^HA) and w;synj^RQ;Atg9^HA. Blots probed with anti‐Synj to assess whether Synj (predicted 134 kDa) is present on Atg9‐positive structures; anti‐HA to assess immunoprecipitation specificity, Atg9^HA (predicted 96 kDa). As control alpha‐SNAP (33 kDa) and complexin (cpx; 18 kDa) where probed. Experiments were performed in independent duplicates.

• B–G Live imaging of fed (B–F) and 4‐h‐starved (B′–F′) NMJ boutons of control (D42‐Gal4 > Atg8^mCherry) (B, B′) and synj^−/− null mutants (C, C′) either expressing synj^RQ (D, D′) or synj^CS (E, E′) or synj⁺ (F, F′) and also expressing Atg8^mCherry (D42‐Gal4 > Atg8^mCherry). Full genotypes are included in the Appendix Supplementary Methods section. Quantification of the number of Atg8^mCherry dots (arrowheads) (G). Statistical analysis with one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, ****P < 0.0001, ***P < 0.001, and each genotype individually fed and starved by t‐test, ****P < 0.0001, n ≥ 9 larvae and n ≥ 18 NMJs per genotype. Error bars represent SEM; scale bar, 5 μm. Note that loss of Synj blocks the formation of Atg8^mCherry dots and this is not rescued by expression of synj^RQ nor synj^CS.

• H–J Live Atg8^mCherry imaging following 30 min of 20‐Hz electrical nerve stimulation of indicated genotypes (H, I). Full genotypes are included in the Appendix Supplementary Methods. Quantification of Atg8^mCherry dots (arrowheads) (J). Statistical analysis by t‐test, **P < 0.01, n ≥ 8 larvae and n ≥ 33 NMJs per genotype. Error bars represent SEM; scale bar, 5 μm.

• K–M Images of NMJ boutons of synj^−/− null mutants either expressing synj⁺ (K) or synj^RQ (L) and also expressing Atg8^mCherry (D42‐Gal4 > Atg8^mCherry) fed (data from G) and fed or starved for 4 h with chloroquine and labeled with anti‐mCherry antibodies. Quantification of the number of Atg8^mCherry dots (arrows) of indicated genotypes (in detail in the Appendix Supplementary Methods) (M). Statistical analysis by t‐test, ****P < 0.0001, **P < 0.01, n ≥ 9 larvae and n ≥ 18 NMJs per genotype. Error bars represent SEM; scale bar. 5 μm. Despite that chloroquine blocks autophagosome to lysosome fusion, Atg8^mCherry dots still do not form in animals expressing synj^RQ.

Next, we assessed whether Synj is important for presynaptic autophagy. Such a role had recently been suggested in zebrafish cone photoreceptors (George et al, 2016). We expressed transgenes for the Atg8^mCherry (fly LC‐3) marker of mature autophagosomes (Kabeya et al, 2000) and Lamp1^GFP, an autolysosomal marker (Marzella et al, 1982; Juhász et al, 2008), in motor neurons using D42‐Gal4. In control animals, autophagy induced by starvation (amino acid deprivation) is marked by the presence of punctate dots of Atg8^mCherry signal within presynaptic terminals (Fig 2B, B′, and G) that we previously characterized by correlative light and electron microscopy (CLEM) as synaptic autophagosome‐like structures (Soukup et al, 2016). Moreover, boutons of starved control animals also accumulate the Lamp1^GFP marker (Appendix Fig S2B, B′, and F). In contrast, in synj null mutants and synj null mutants that express Synj^RQ, the Atg8^mCherry and Lamp1^GFP redistribution is blocked (Fig 2C–D′ and G, and Appendix Fig S2C–D′ and F). Unlike our findings with Synj‐driven endocytosis, we observe that only wild‐type Synj⁺, and not Synj^RQ, can recover the Synj function in presynaptic autophagy (Fig 2F and G, and Appendix Fig S2E and F). Our results are also not unique to “starvation‐induced autophagy” because when we induce the process by direct electrical motor neuron stimulation (30 min 20 Hz) (Soukup et al, 2016), the clear increase in the number of Atg8 dots observed in controls is absent in synj^RQ mutants (Fig 2H–J).

To scrutinize our results further and provide evidence that the block in synaptic autophagy is caused by the lack of SAC1 activity, we created flies that express Synj^C396S (Synj^CS), a SAC1 enzymatic dead mutant (Mani et al, 2007). We expressed Synj^CS in synj^−/− null mutants under the same conditions as used to express Synj^RQ using D42‐Gal4 driver. We then assessed Atg8^mCherry localization and found that Atg8^mCherry also does not concentrate in dots upon induction of starvation (Fig 2E, E′, and G). These data are in further support of the notion that Synj SAC1 domain function is required for synaptic autophagy and that the pathogenic PD mutant Synj behaves in this respect similar to a Synj‐SAC1 domain mutant.

Next, to distinguish whether loss of Synj SAC1 function blocks autophagosome formation or promotes autophagosome degradation, we pharmacologically blocked the fusion of autophagosomes with late endosomes/lysosomes using chloroquine (Stauber et al, 1981; Chi et al, 2010). Subsequently, we monitored Atg8^mCherry localization. As expected in wild‐type controls, chloroquine under fed or starved conditions causes an increase in the number of Atg8^mCherry dots. In contrast, in synj^RQ mutants, chloroquine does not result in such an increase in the number of Atg8^mCherry dots (Fig 2K–M), suggesting autophagosome biogenesis is slowed. We provide further evidence for this by using the flux marker Atg8 tagged with GFP and RFP in tandem. Here, Atg8‐positive organelles are green and red fluorescent, but when they fuse with lysosomes, the GFP is quenched and autolysosomes are only red (DeVorkin & Gorski, 2014). We quantified the number of yellow and red Atg8‐positive organelles in controls as well as the rare Atg8‐positive organelles in synj^RQ mutants. We do not observe a significant difference in the ratio of red (mature) over total Atg8‐labeled organelles between control and synj^RQ (Control: 55%; synj^RQ: 57%). The data indicate that once a rare Atg8‐positive autophagosome manages to form in synj^RQ mutants, its further maturation is not impeded (Appendix Fig S2G–I).

Synj mutants accumulate Atg18a within synaptic boutons

Members of the PROPPIN domain‐containing protein family are key PI(3)P‐ and PI(3,5)P₂‐binding proteins with a role in autophagy (Baskaran et al, 2012). Given that Synj1^RQ cannot dephosphorylate PI(3)P (and likely PI(3,5)P₂), we hypothesized that in the synj null mutants that express Synj^RQ, autophagic PROPPIN‐containing proteins would accumulate on synaptic pre‐autophagosomal membranes. Therefore, we generated flies expressing N‐terminal GFP‐tagged Drosophila homologs of the PROPPIN WIPI2 (Atg18a, Atg18b, and CG11975; 70.8, 56.5, and 32.0% identity to human WIPI2 respectively). Both Atg18a^GFP and CG11975^GFP localize in puncta at synaptic boutons when expressed in motor neurons (D42‐Gal4) (Appendix Fig S3A, B, G, and H). In contrast, Atg18b^GFP is retained in neuronal cell bodies (Appendix Fig S3E and F). After starvation, we observe a clear upregulation of the number of Atg18a^GFP puncta, while the number of CG11975^GFP dots does not change (Appendix Fig S3C, D, I, and J and Fig 3A, A′, and E). Additionally, Atg18a^GFP puncta sometimes colocalize with Atg8^mCherry puncta (Appendix Fig S3K and L) and expression of Atg18a^GFP using the ubiquitous Da‐Gal4 driver in atg18a null mutants rescues their lethality. These data indicate that Atg18a^GFP is a functional protein.

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Figure 3. Atg18a‐positive structures accumulate at synaptic boutons of synj mutants

• A–E Live imaging of fed (A–D) and 4‐h‐starved (A′–D′) NMJ boutons in controls (D42‐Gal4 > Atg18a^GFP) (A, A′) and synj^−/− null mutants (B, B′) or synj^−/− null mutants expressing either synj^RQ (C, C′) or synj⁺ (D, D′) and also expressing Atg18a^GFP (D42‐Gal4 > Atg18a^GFP). Quantification of the number of Atg18a^GFP dots (arrowheads) of indicated genotypes under fed and starved conditions (E). Full genotypes appear in the Appendix Supplementary Methods. Statistical analysis by t‐test comparing control to individual genotypes separately in fed conditions and each genotype individually fed and starved, **P < 0.01, *P < 0.05, n ≥ 10 larvae and n ≥ 20 NMJs per genotype. Error bars represent SEM; scale bar, 5 μm. Note that loss of Synj increases the number of Atg18a^GFP dots even under fed conditions. This is not rescued by expression of synj^RQ.

• F–P Correlative light and electron microscopy (CLEM) of 4‐h‐starved synj^−/− null mutant NMJ boutons expressing Atg18a^GFP and synj^RQ under the control of D42‐Gal4. NMJ boutons with Atg18^GFP fluorescence were imaged (F), and subsequently surrounded by burn marks using near‐infrared branding (NIRB) (F–H). Branding marks are clearly visible in transmission and fluorescent mode due to autofluorescence of the marks (F, G). Branding marks allow subsequent detection of the same region of interest in electron micrographs (H). The marks then allow for the correlation of fluorescent and electron micrographs of Atg18a^GFP‐positive boutons (I–K). Arrows indicate Atg18^GFP‐positive labeling (intensities were adjusted as to identify structures) in EM micrographs and boxes are enlarged in subsequent panels. Magnification of Atg18a^GFP‐positive structures identified in (I–K) is shown in (L) and further examples of Atg18^GFP‐labeled structures are taken from independent CLEM experiments (M–P). Scale bars: 50 μm (F–H), 5 μm (I), 50 nm (J, K), and 200 nm (L–P).

• Q–U Representative transmission electron micrographs of 4‐h‐starved synj^−/− null mutants expressing either synj⁺ (Q) or synj^RQ (R) using D42‐Gal4, as well as of 4‐h‐starved synj^−/− null mutants (S), controls (w¹¹¹⁸ (T)), and synj^RQ knock‐in animals (U). Full genotypes appear in the Appendix Supplementary Methods. Scale bars, 1 μm.

• V. Quantification of the number of synaptic vesicles (diameter < 80 nm) and cisternae that resemble Atg18a^GFP‐positive structures identified using CLEM. Note that at boutons of synj^−/− null mutants, synj^−/− null mutants expressing synj^RQ and synj^RQ, there are cisternae that resemble the Atg18a^GFP‐positive structures identified in CLEM; in synj^−/− null mutants expressing synj^RQ and in synj^RQ knock‐in, there are numerous synaptic vesicles detectable. For statistical analysis of both SV and cisternae, t‐test comparing genotypes individually, *P < 0.05, ***P < 0.001, ****P < 0.0001. n ≥ 3 larvae and n ≥ 8 NMJs per genotype. Error bars represent SEM.

We then assessed Atg18a^GFP localization in synj null mutants that express synj^RQ. Consistent with Synj^RQ not being able to dephosphorylate PI(3)P (Krebs et al, 2013), we find a significant increase in the number of Atg18a^GFP dots in fed synj mutants in comparison with fed controls (Fig 3A–C′ and E). In addition, the number of Atg18a^GFP dots is also increased in synj null mutant boutons that express synj^RQ compared to synj null mutant boutons that express synj⁺ (Fig 3C–E). Atg18a can bind PI(3)P and PI(3,5)P₂ that are both dephosphorylated by the SAC1 domain of Synj1. To delineate which phosphoinositide is involved, we took a pharmacological approach. We used wortmannin (WM) that inhibits Vps34, a kinase that produces PI(3)P, or YM201636 (YM) that inhibits PIKfyve, a kinase that produces PI(3,5)P₂. We applied these drugs during starvation to control and synj^RQ mutant animals that express Atg18a^GFP. While Atg18a^GFP accumulates in dots in starved controls, application of either drug blocks this effect and Atg18a^GFP does not accumulate anymore, indicating that Atg18a^GFP likely binds both PI(3)P and PI(3,5)P₂ during autophagy at synapses (Appendix Fig S3M). In synj^RQ mutants, the number of Atg18a^GFP dots is always upregulated. However, when we starve the larvae in the presence of either WM or YM, the number of Atg18a^GFP is lower than in untreated starved synj^RQ mutants (Appendix Fig S3M). These data suggest that consistent with the previously established function of the Synj SAC1 domain, both PI(3)P and PI(3,5)P₂ accumulate and bind Atg18a in synj^RQ mutants.

We tried to find further evidence for upregulation of PI(3)P in synj^RQ using expression of 2xFYVE‐GFP, a probe that binds PI(3)P. We were however unsuccessful and did not detect obvious differences (not shown). Nonetheless, we do believe our results are consistent with the specific loss of SAC1 domain function in synj^RQ. We assessed 5‐phosphatase domain activity of Synj using the PLCδ‐PH‐eGFP probe that detects PI(4,5)P₂. We find that the synaptic labeling intensity of PLCδ‐PH‐eGFP is much less affected in synj null mutants that express Synj^RQ (where 5‐phosphatase activity is not affected) compared to synj null mutants (where 5‐phosphatase activity is affected) (Appendix Fig S3N–R), suggesting synj^RQ mutants support PI(4,5)P₂ dephosphorylation. Taken together, the data are consistent with normal PI(4,5)P₂ dephosphorylation activity of Synj1^RQ measured in vitro (Krebs et al, 2013), but altered PI(3)P dephosphorylation, thereby leading to the accumulation of Atg18a at synaptic boutons in synj^RQ mutants.

Visualization of Atg18a‐labeled structures at synaptic boutons

Synapses in synj mutants expressing Synj^RQ harbor increased synaptic Atg18a^GFP accumulations and largely lack Atg8^mCherry accumulations. To further visualize the Atg18a^GFP‐labeled structures, we resorted to CLEM, a methodology that allows us to directly determine the ultrastructural appearance of Atg18a‐positive structures within synaptic boutons. Synj mutants expressing Synj^RQ and Atg18a^GFP were dissected, fixed, and imaged. Subsequently, a near‐infrared laser was used to produce burn marks in the vicinity of the imaged region. These marks are easily discernable at the light and EM level and help us when producing serial sections of the imaged boutons (Fig 3F–H). After alignment of the confocal images with the serial TEM images, we are able to identify the Atg18a^GFP‐positive structures (Fig 3I–K). As indicated in Fig 3L–P, Atg18a^GFP marks profiles that have the appearance of cisternae that sometimes contain a few smaller vesicular structures in their lumen. Given that the flux of synaptic autophagy is blocked in synj mutants expressing Synj^RQ and that these mutants do not harbor Atg8^mCherry dots, we propose that these cisternae are early presynaptic autophagic intermediates.

Using the morphological features of Atg18a^GFP‐positive structures as a guide, we assessed the ultrastructural morphology of synapses in starved animals. We used TEM images to quantify the number of synaptic vesicles and the number of structures that have an appearance similar to the Atg18a‐positive cisternae in our CLEM studies (Fig 3V). Compared to starved controls and synj null mutants expressing Synj⁺, starved synj null mutants show a strong reduction in the number of synaptic vesicles per bouton area, consistent with a defect in synaptic endocytosis. These mutants also show an increased number of cisternal structures (Fig 3Q, S, T, and V), correlating with an increased number of Atg18a‐positive dots in synj null mutants. Starved synj null mutants that express synj^RQ and synj^RQ knock‐in animals also show an increased number of cisternal structures at boutons (Fig 3R, U, and V). However, there is a largely normal number of synaptic vesicles compared to controls (Fig 3Q–V), consistent with the normal synaptic vesicle endocytosis that we measured in these mutants (see Fig 1).

Synaptojanin regulates Atg18a mobility at autophagosomal membranes

In synaptic vesicle endocytosis, the dephosphorylation of PI(4,5)P₂ by Synj on nascent vesicles regulates the detachment of adaptors with affinity for this lipid. We reasoned that a similar mechanism may be at play during autophagosome formation and tested whether Synj regulates Atg18a attachment and detachment. We expressed Atg18a^GFP and assessed the fluorescence recovery after bleaching one specific Atg18a^GFP dot. Synj null mutants and synj null mutants expressing Synj^RQ show significantly slower recovery of Atg18a fluorescence in the bleached area compared to synj null mutants expressing Synj⁺ or compared to wild‐type controls (Fig 4A–E).

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Figure 4. Synaptojanin regulates Atg18a uncoating at autophagosomal membranes

• A–E Representative time‐lapse images of a FRAP experiment where the fluorescence recovery after bleaching of an individual Atg18a^GFP dot (D42‐Gal4 > Atg18a^GFP) was monitored at a synaptic bouton of a control (A) and of a synj^−/− null mutant (B) and of a synj^−/− null mutant expressing either synj^RQ (C) or synj⁺ (D) (expressed using the D42‐Gal4). (E) Quantification of fluorescence intensities of a bleached Atg18a^GFP dot (arrowheads) in the indicated genotypes (in % of initial fluorescence of the Atg18a^GFP dot before bleaching), error bars represent SEM. Full genotypes appear in the Appendix Supplementary Methods. Note the reduced fluorescence recovery of Atg18a in synj null mutants and synj null mutants expressing synj^RQ compared to wild‐type controls. Even in wild‐type Atg18a^GFP, fluorescence never recovers more than 60%, suggesting an immobile pool of protein on the nascent autophagosome.

• F–J Transgenic larvae expressing Atg18a^mEos3.2 were imaged at 33 Hz. (F–F‴) Representative low‐resolution image of NMJ bouton (F), sptPALM trajectory map (F′), average intensity (F″), and diffusion coefficient (F‴) of Atg18a^mEos3.2. Scale bar, 5 μm. (G) Comparison of MSD of Atg18a^mEos3.2 in fed and starved conditions (n ≥ 6 larvae and n ≥ 18 NMJ per condition). (H) Analysis of the area under the MSD curve (μm²s) showed significant decrease in starved compared to fed controls. (I) Diffusion coefficient distribution of Atg18a^mEos3.2 in fed and starved controls. (J) Mobile‐to‐immobile ratio for fed (13.4 ± 1.8) significantly reduced in starved conditions (7.1 ± 0.5). Statistical analysis by one‐way ANOVA with Tukey's multiple comparison test, **P < 0.01. Error bars represent SEM.

• K–M Representative time‐lapse images of mEOS3.2 photoconversion experiment were the RFP fluorescence recovery on an individual Atg18a^mEOS3.2 after GFP conversion of the neighboring bouton, an individual Atg18a^mEOS3.2 dot (D42‐Gal4 >  Atg18a^mEOS3.2) was monitored at a synaptic bouton of a control (K) and of a synj^RQ (L). (M) Quantification of the ratio RFP/GFP fluorescence intensities of an Atg18a^mEOS3.2 dot (arrowheads) in the indicated genotypes (in % of final time point (10 min after photoconversion) RFP/GFP fluorescence ratio of the Atg18a^mEOS3.2 dot), error bars represent SEM. Full genotypes appear in the Appendix Supplementary Methods. Statistical analysis comparing curves fitted by linear regression. n ≥ 3 larvae and n ≥ 13 NMJs per genotype. Note the reduced RFP/GFP fluorescence recovery of Atg18a^mEOS3.2 in synj^RQ compared to wild‐type controls.

To find further evidence for our model, we also generated Atg18a^mEOS3.2‐expressing flies. mEos is a photoconvertible fluorescent protein that shifts from green to red fluorescence when activated (Zhang et al, 2012). We reasoned that Atg18a mobility would decrease after induction of autophagy, as more Atg18a would bind to newly forming autophagosomes. Therefore, we used single‐particle tracking photoactivation localization (sptPALM) microscopy to track single molecules within synaptic boutons expressing Atg18a^mEOS3.2 (Bademosi et al, 2017). Tracking each fluorescent molecule (Video EV1) allows us to generate trajectory maps of Atg18a^mEOS3.2, as well as intensity and instantaneous diffusion coefficient maps (Fig 4F–F‴). Moreover, we quantified Atg18a^mEOS3.2 mobility by analyzing the mean square displacement (MSD) of sptPALM trajectories of single fluorescent localizations (Fig 4G). Compared to fed larvae, Atg18a mobility significantly decreases in starved larvae (Fig 4H), in good agreement with recruitment of cytosolic molecules to presynaptic autophagosomes. Analysis of the diffusion coefficient distribution reveals a decrease in the mobile population of Atg18a^mEOS3.2 in starved conditions (Fig 4I). This is most apparent in the ratio of the mobile‐to‐immobile fractions (Fig 4J). This change in mobility of Atg18a after induction of autophagy is consistent with the recruitment of Atg18a to presynaptic autophagosomes.

We subsequently followed the accumulation of Atg18a on synaptic autophagosomes by converting a boutonic pool of Atg18a^mEOS3.2. We converted Atg18a^mEOS3.2 in one bouton and followed accumulation of red Atg18a^mEOS3.2 on (green Atg18a^mEOS3.2‐positive) autophagosomes in the neighboring boutons of control and synj^RQ knock‐in flies. While the amount of red photoconverted Atg18a in the cytoplasm of controls and synj^RQ is similar, the red Atg18a accumulates significantly less quickly on nascent autophagosomes in synj^RQ compared to controls (Fig 4K–M). This slowed acquisition of red fluorescence indicates reduced exchange between membrane bound and soluble pools of Atg18a, and thus indicates that the SAC1 domain of Synj normally promotes Atg18a mobility at synaptic autophagosomes.

Synj^RQ reduces lifespan upon starvation and causes neurodegeneration

To more broadly assess the physiological consequences of the Synj^RQ mutation, we examined whether this affects animal survival under starvation conditions, a readout that is strongly modulated by autophagy (Juhász et al, 2007). While wild‐type flies reared in these conditions can survive for ~4 days, we detect significantly reduced survival times for synj null mutants that express Synj^RQ in neurons or synj^RQ knock‐in animals which display 100% lethality within 3 days (Fig 5A and A′). The results correlate with impaired autophagosome maturation in animals lacking Synj SAC1 activity, suggesting that presynaptic autophagy connects to an important survival mechanism during starvation in the fruit fly.

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Figure 5. Defects in synaptic autophagy result in reduced lifespan upon starvation and neurodegeneration

• A, A′ Complete starvation (water‐only diet) assay in synj null mutants expressing synj^RQ (light green) or synj⁺ (blue) (expressed using nSyb‐Gal4) (A) and synj^RQ knock‐in animals (dark green) as well as controls (w¹¹¹⁸, black) (A′). Statistical analysis of survival curves by log‐rank (Mantel–Cox) test, ****P < 0.0001, n ≥ 134 flies per genotype for (A) and n ≥ 292 flies per genotype for (A′), dotted lines surrounding the curves indicate SEM. Note, synj null mutants expressing synj^RQ as well as synj^RQ knock‐in flies have a shorter life span under these starvation conditions.

• B–E Representative electron micrographs of the retina after 7 days of continuous light exposure of control (nSyb‐Gal4/+) (B) and synj null mutants expressing synj^RQ (C) or synj⁺ (D) (expressed using nSyb‐Gal4). Green highlighting in (C) shows disintegrating rhabdomeres as a sign of degeneration. Scale bar, 2 μm. (E) Quantification of number of intact ommatidia per indicated genotype when flies were kept in complete darkness or when exposed to light. Full genotypes are in the Appendix Supplementary Methods. Statistical analysis by t‐test, ****P < 0.0001, n = 3–5 flies per genotype. Error bars represent SEM.

• F–F″ Dopaminergic neuron degeneration assay in 30‐day‐old control (w¹¹¹⁸) and synj^RQ knock‐in flies. Schematic representation (F) and image (z‐stack, sum of slices) of posterior side of an anti‐TH‐labeled 30‐day‐old adult fly brain (F′) to show the location of the different dopaminergic neuron cell bodies in this part of the brain. Scale bar, 40 μm. (F″) Bar graph showing the number of dopaminergic neurons (TH‐positive) in each cluster indicated in (F) in 30‐day‐old flies of control (black bars) and synj^RQ knock‐in (dark green bars) flies. Statistical analysis per dopaminergic cluster by t‐test, **P < 0.01, *P < 0.05, n ≥ 15 brains per genotype. Error bars represent SEM. Note at PPM3 and PPL1 dopaminergic neuron clusters the reduction of anti‐TH‐positive neuronal cell bodies in the of synj^RQ mutant animals compared to controls.

Next, we tested neuronal survival, another parameter sensitive to autophagy (Komatsu et al, 2006; Juhász et al, 2007). To induce neurodegeneration, flies were kept 7 days in constant light challenging photoreceptor neurons of the fly eye. In synj null mutant flies that express Synj^RQ, a significant reduction in the number of intact ommatidia and an accumulation of vacuoles was observed. In contrast to control animals, when synj null mutant animals expressing Synj⁺ or animals kept in dark for 7 days, these photoreceptors appear unaffected (Fig 5B–E). Finally, given the vulnerability of dopaminergic neurons in PD, we assessed the integrity of such neurons in fly brains of 30‐day‐old synj^RQ knock‐in animals using anti‐TH labeling. In these mutants, anti‐TH‐positive neuronal cell bodies in the PPM3 and PPL1 dopaminergic neuron clusters were significantly reduced compared to controls (Fig 5F–F″). Hence, Synj SAC1 domain function is required for dopaminergic neuron survival in the fly brain.

WIPI2/Atg18a accumulates in neurites of SYNJ1 R258Q patient‐derived human induced neurons

To confirm our key findings in a disease‐relevant condition, we resorted to human iPSC obtained from two patients with autosomal recessive, early‐onset PD carrying the R258Q mutation in SYNJ1 as well as two age‐matched controls (Appendix Table S1). Following validation, including expression analysis of pluripotency and proliferation markers (Fig 6A–B′, Appendix Fig S4 and Appendix Table S1), we differentiated these iPSC into neurons. After 28 days of differentiation, we labeled the neurons with anti‐WIPI2 (mammalian Atg18a) and imaged the neurites (Fig 6C–F). Similar to our observations at fly synapses, we find a significant increase in the number of WIPI2 dots in starved control neurons compared to fed control neurons (Fig 6C, D, and G). Moreover, in line with our observations made with fly synj^RQ mutants, we find a significant increase in the number of WIPI2 dots in fed SYNJ1^RQ neurons compared to fed controls and which is even further increased in starved SYNJ1^RQ neurons (Fig 6C and E–G). These data indicate that also in human neurons, Synj1 is required for autophagy, and further supports our model that implicates SYNJ1‐SAC1 function in removing Atg18a/WIPI2 from nascent autophagosomal membranes.

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Figure 6. WIPI2/Atg18a accumulates in neurites of SYNJ1 R258Q human patient iPSC‐derived neurons

• A–B′ Immunocytochemistry analysis of human iPSC‐derived neurons (after 28 days of differentiation) showing expression of neuronal markers, anti‐MAP2 (magenta) and anti‐synapsin 1 (green) in control (A, A′) and SYNJ1^RQ (B, B′). Scale bars, 15 μm.

• C–G Images of neurons stained with anti‐WIPI2 in fed and starved conditions of controls (C, D) and SYNJ1^R258Q (E, F). (G) Quantification of the number of WIPI2 dots of indicated genotypes (two control lines and two patients lines (NAPO16 and 17), circles and triangles indicate separate lines for each genotype) under fed and starved conditions. Statistical analysis with one‐way ANOVA Kruskal–Wallis followed by Dunn's multiple comparison post hoc test, ****P < 0.0001, **P < 0.01, ns, not significant. n ≥ 21 images per condition per genotype. Error bars represent SEM; scale bar, 15 μm. Note that SYNJ1^RQ has increased number of WIPI2 dots under fed conditions.

Fly stocks and genetics

Flies were grown on standard cornmeal and molasses medium supplemented with yeast paste at 25°C. The following Drosophila stocks were obtained from the Bloomington Stock Center: w; P{hs‐I‐CreI.R}1A Sb1/TM6, y[1] w[*]; Mi{y[+mDint2]=MIC}Synj[MI11871]/SM6a (Venken et al, 2011). The following fly stocks were obtained from the laboratory where the stock was first described: w;UAS‐mCherry‐Atg8a (Chang & Neufeld, 2009), UAS GFP::Lamp1/CyO;TM6B/Sb bos1 (Pulipparacharuvil et al, 2005), w;;UAS‐PLCd‐PH‐EGFP (Verstreken et al, 2009), yw,eyFLP GMR‐LacZ;FRT42D synj¹/CyO, and yw,eyFLP GMR‐LacZ;FRT42D synj²/CyO (Verstreken et al, 2003), w;;atg18a(W3)/TM6b,Hu,Tb and w;;atg18a(W4)/TM6b,Hu,Tb (Shen & Ganetzky, 2009), genomic w;;Atg9^HA (G. Juhász, unpublished).

Transgenic flies

A list of fly genotypes used in this study can be found in the Appendix Supplementary Methods section. All constructs were generated using a seamless cloning method (Gibson Assembly, NEB). UAS synj^RQ, UAS synj^CS, and UAS synj⁺ were created using gBlocks (Integrated DNA Technologies, Inc.) and PCR products generated using primers listed in the Appendix Supplementary Methods section from CH322‐135G1 P[acman] plasmid [BACPAC Resources Center (BPRC)] (Venken et al, 2006, 2009), cloned into the EcoRI site of pUAST‐attB, sequence‐verified, and inserted into attP site VK37 (22A3) using PhiC31‐mediated integration at BestGene Inc.

synj^RQ knock‐in flies were created as described in Vilain et al (2014). Targeting vector MiMIC 1322 pBS‐KS‐attB1‐2 I‐SceI Synj^RQ I‐CreI was generated using primers listed in the Appendix Supplementary Methods section from CH322‐135G21 P[acman] plasmid (BPRC) and cloned first into XhoI and PstI sites of pSV004 (containing I‐SceI and I‐CreI sites, see Appendix Supplementary Methods section) followed by restriction with SmaI, gel purification, and cloning into BbsI sites of MiMIC 1322 pBS‐KS‐attB1‐2 vector with an additional linker sequence containing BbsI sites (see Appendix Supplementary Methods section) into XbaI and EcoRI sites. Next, the construct was inserted into flies with Mi{MIC}Synj^MI11871 using PhiC31‐mediated integration by in‐house injection. Efficiencies were 30% for integration into the Mi{MIC}Synj^MI11871, 15% for desired orientation, and 37% of I‐CreI double‐strand break/repair. After genomic manipulation, the whole locus was sequence‐verified.

UAS GFP::Atg18a, UAS GFP::Atg18b, UAS GFP::CG11975, UAS mEOS3.2::Atg18a, UAS eGFP‐mCherry::Atg8a were created using gBlocks (Integrated DNA Technologies, Inc.) and PCR products generated using primers listed in the Appendix Supplementary Methods section, all Atg18 homologues cDNAs as template were obtained from the Proteomic ORF Collection (Yu et al, 2011) (respectively, LD38705, GH07816, and LD32381), mEOS3.2 was PCR from Addgene plasmid #54550 and cloned into the XhoI site of pUAST‐attB, and Atg8 was cloned in between the EcoRI and XbaI sites of pUAST‐attB. All constructs were sequence‐verified and inserted into attP site VK20 (99F8) using PhiC31‐mediated integration at BestGene Inc or by in‐house injection.

Induction of autophagy by starvation, electrical stimulation, and chloroquine assay

Autophagy induction and flux assay were performed by placing early third‐instar larvae on petri dishes containing 20% sucrose and 1% agarose for 3–4 h prior to experimentation (Scott et al, 2004). Electrical stimulation, 20 Hz for 30 min, to induce autophagy was conducted as described before (Soukup et al, 2016). For the chloroquine assay, we fed third‐instar larvae 4 h on standard and starvation food containing 10 mg/ml chloroquine diphosphate salt (Sigma).

Correlative light and electron microscopic studies (CLEM and TEM)

Correlative light and electron microscopy studies were performed as described in Soukup et al (2016). Before dissection in HL3, larvae were starved for 3–4 h. The samples were directly fixed for 2 h at room temperature in the dark (0.5% glutaraldehyde, 2% paraformaldehyde in 0.1 M phosphate buffer, pH = 7.2), followed by washing steps in 0.1 M phosphate buffer, pH = 7.2. near‐infrared branding (NIRB) was performed on a LSM 710 upright confocal microscope using the bleaching function as described (Urwyler et al, 2015). Before and after NIRB, z‐stacks of the region of interest were acquired. Next, larvae were post‐fixed in 1% glutaraldehyde, 2% paraformaldehyde in 0.1 M phosphate buffer, pH = 7.2 at 4°C overnight or until further processing as described in Fernandes et al (2014). NIRB introduced branding marks were used as a guide to identify Drosophila NMJ boutons previously imaged with the confocal microscope. Electron and fluorescent micrographs overlay, and identification of Atg18a^GFP‐positive ultrastructural features and processing of micrographs using brightness and contrast were performed with NIH ImageJ and Adobe Photoshop Elements 14.

TEM studies of larval boutons were performed as described in Kasprowicz et al (2008). Larvae were starved for 3–4 h prior to dissection and fixation. TEM of the adult fly heads was performed as described previously in Slabbaert et al (2016).

Imaging and quantification

For live imaging of Atg8^mCherry, Atg8^{GFP‐mCherry}, Atg18a^GFP, Atg18b^GFP, CG11975^GFP, or Lamp1^GFP, larvae expressing these markers were dissected in HL3 on sylgard plates and imaged on a Nikon A1R confocal microscope with a 60× NA 1.0 water‐dipping lens. All images were collected with a pinhole of 1 airy unit and a resolution of 1,024 × 1,024. Imaging settings between experiments with the same marker were identical. Images are shown either in grayscale or using a 16‐color lookup table (the LUT is shown in the figure).

Quantification of number of puncta or intensity was performed in NIH ImageJ. Atg8^mCherry, Atg8^{GFP‐mCherry}, Atg18a^GFP, or CG11975^GFP dots were counted using particle analysis of a binary image combined with applying a threshold mask on the cytoplasmic signal to calculate the bouton area. Lamp1^GFP fluorescence intensities were measured by first applying a threshold to mark the boutonic areas followed by measuring the average pixel intensities in boutonic areas. The different conditions were blinded before imaging and quantifying the data.

Fluorescence recovery after photobleaching (FRAP) was performed on third‐instar larval fillets dissected in HL3 on sylgard‐coated plates using a protocol adapted from Seabrooke et al (2010). Experiments were conducted on a Nikon A1R confocal microscope with a 60× NA 1.0 water‐dipping lens. Before bleaching, 10 images were acquired to set the pre‐bleach intensity. Next, bleaching (minimum 50% of the pre‐bleach intensity, by two rapidly iterated laser bursts), using a 488‐nm laser, was focused on an Atg18a^GFP dot, directly followed by image acquisition for 120 s (60 scans with a 2,12‐s interval). The fluorescence intensity of the selected Atg18a^GFP dot was analyzed in NIH ImageJ.

Photoconversion of mEOS3.2 was performed on third‐instar larval fillets dissected in HL3 on sylgard‐coated plates; next, nerves were cut in HL3 with NAS (100 μM) followed by mounting the larval prep on a glass slide in HL3 with NAS. Experiments were conducted on a Nikon A1R confocal microscope with a 60× NA1.4 oil lens; all images were collected with an open pinhole and a resolution of 1,024 × 1,024. Images of the Atg18a^mEOS3.2 dot of interest were taken before, directly after, and 5 and 10 min after photoconversion (conducted using a 405‐nm laser) of the neighboring bouton. The fluorescence intensity of the selected Atg18a^mEOS3.2 dot was analyzed in NIH ImageJ.

Single‐particle tracking photoactivation localization microscopy (sptPALM)

SptPALM was carried out on transgenic Atg18a^mEOS3.2 third‐instar Drosophila larvae dissected on sylgard base as previously described (Bademosi et al, 2017). Briefly, Atg18a^mEOS3.2 was imaged at the larva NMJ on muscles 6 and 7 of the second abdominal segments, using total internal reflection (TIRF) microscopy under oblique illumination. Dissected larvae on the sylgard base were inverted onto glass‐bottomed imaging dishes (In Vitro Scientific). Localization and tracking of single Atg18a^mEOS3.2 molecules were carried out using a 63× water‐immersion objective (1.2 numerical aperture, C‐Apochromat) on the ELYRA PS.1 microscope (Zeiss). NMJs were located using 488‐nm laser (18.72 mW) illumination. A 405‐nm laser (0.6 mW) was used for photoconversion, while imaging of photoconverted Atg18a^mEOS3.2 molecules was done using 561‐nm laser (20.6 mW) to spatially visualize and temporally distinguish individual molecules (405‐nm laser was used at 0.009–0.3% power, while 75% power was used for 561 nm). Single‐molecule fluorescence was collected by a sensitive electron‐multiplying charge‐coupled device (EMCCD) camera (Evolve, Photometric). Zen Black acquisition software (2012 version, Carl Zeiss) was used for movie acquisition. Images were captured at 33 Hz and 15,000 frames were obtained per NMJ. sptPALM analysis is described in Appendix Supplementary Methods section.

Complete starvation

Complete starvation assay was performed as described in Juhász et al (2007), briefly 1‐day‐old fed flies were transferred to vials containing a gel of water (1% agarose in water) and transferred to a new vial every day. Dead flies were counted manually multiple times a day.

Light‐induced degeneration

Light‐induced retinal degeneration studies were performed by rearing flies for 7 days in constant illumination at 1,300 lux and 25°C (Soukup et al, 2013).

Visualization of TH‐positive neuronal clusters after aging adult flies

Aging and visualization of dopaminergic clusters was performed by aging 1‐ to 3‐day‐old adult flies for an additional 30 days at 25°C (flies were placed on fresh medium every 3 days). At day 30, fly brains were dissected and stained (see immunohistochemistry). Images of dopaminergic clusters (labeled by the anti‐TH antibody) were captured with a Nikon A1R confocal microscope using a 20× objective 0.75 NA (Nikon). The total number of TH‐positive neurons throughout the brain and in both hemispheres for the PPM1, PPM2, PPM3, and PPL1 clusters was counted manually.

Neural progenitor differentiation

The iPSCs (generation and karyotyping described in Appendix Supplementary Methods and Appendix Table S1) were grown on irradiated MEFs and cultured in normal iPSC medium (DMEM/F12 Advanced (Gibco Life Technologies) supplemented with 20% KOSR, 2 mM l‐glutamine, 0.1 mM MEM‐NEAA, 0.1 mM 2‐mercaptoethanol, 100 units/ml penicillin/streptomycin (all from Life Technologies)) and 10 ng/ml bFGF (Perprotech) at 37°C/5% CO₂. Medium was changed daily and cells were passaged every 4–6 days either mechanically or enzymatically with collagenase type IV (100 U/ml) (Life Technologies), and addition of 10 μM ROCK inhibitor (Sigma) to the medium after passaging.

Neural progenitor cells (NPCs) were created according to published protocols (de Esch et al, 2014; Gunhanlar et al, 2017) with minor modifications. Briefly, for embryoid body (EB) generation, iPSCs were dissociated and transferred to non‐adherent plates in normal iPSC medium without bFGF and cultured on a shaker at 37°C/5% CO₂. After 2 days in vitro (DIV), medium was replaced by neural induction medium (DMEM/F12 Advanced, 1:100 N2 supplement, 100 units/ml penicillin/streptomycin (all from Life Technologies) and 2 μg/ml heparin (Sigma)). After 7 DIV, the EBs were partially dissociated and plated on laminin‐coated culture dishes in neural induction medium.

Neural progenitor cells growing out of the attached EB fragments were cultured for 8 DIV at 37°C/5% CO₂, and medium was changed every other day. Subsequently, the NPCs were split between 1:2 and 1:6 with collagenase every 5–7 days and cultured on laminin (Sigma)‐coated plates. The NPCs were maintained in NPC medium [DMEM/F12 Advanced, 1:100 N2 supplement, 1:50 B27‐RA supplement, 100 units/ml penicillin/streptomycin (all from Life Technologies), 1 μg/ml laminin (Sigma), and 20 ng/ml bFGF (Preprotech)].

Neural differentiation

Neural progenitor cells were passaged 1:1 to poly‐D‐lysine (PDL) (Sigma)‐ and laminin (Sigma)‐coated coverslips and cultured in neural differentiation medium [Neurobasal medium supplemented with 1:100 N2 supplement, 1:50 B27 supplement, 0.1 mM MEM‐NEAA, 100 units/ml penicillin/streptomycin (all from Life Technologies), 20 ng/ml BDNF (ProspecBio), 20 ng/ml GDNF (ProspecBio), 1 μM db‐cAMP (Sigma), 200 μM ascorbic acid (Sigma), and 2 μg/ml laminin (Sigma)]. The differentiating cells were cultured for 28 DIV at 37°C/5% CO₂, and medium was changed every other day.

Neuron starvation

At 28 DIV, neurons were either incubated for 3 h in neural differentiation medium supplemented with 100 μM leupeptin (Roche) for the control condition, or in preconditioned EBSS (Life Technologies) supplemented with 100 μM leupeptin (Roche) for the starved condition.

Statistics

GraphPad Prism 6 (San Diego, USA.) was used to determine statistical significance. Prior to this, datasets were tested for normal distribution using a D'Agostino–Pearson Omnibus test. Next, normally distributed data were tested with parametric tests: Student's t‐test was used when two datasets were compared and we used a one‐way analysis of variance test (ANOVA) followed by a post hoc Tukey test when more than two datasets were compared. Non‐normally distributed data were tested using non‐parametric tests: when comparing two datasets, we used a Mann–Whitney test, and when comparing more than two datasets, we used an ANOVA Kruskal–Wallis test followed by a Dunn's post hoc test. Significance levels are defined as ****P < 0.0001, ***P < 0.001, **P < 0.01, *P < 0.05, ns (not significant) = P > 0.05. “n” in the legends indicates either the number of animals used or the number of NMJs analyzed.