RESEARCH ARTICLE

The effects of ERmorphology on synaptic structure and function inDrosophila melanogasterJames B. Summerville1,*, Joseph F. Faust1,*, Ethan Fan1, Diana Pendin2, Andrea Daga3, Joseph Formella1,Michael Stern1,‡ and James A. McNew1,‡

ABSTRACTHereditary spastic paraplegia (HSP) is a set of genetic diseasescaused bymutations in one of 72 genes that results in age-dependentcorticospinal axon degeneration accompanied by spasticity andparalysis. Two genes implicated in HSPs encode proteins thatregulate endoplasmic reticulum (ER) morphology. Atlastin 1 (ATL1,also known as SPG3A) encodes an ER membrane fusion GTPaseand reticulon 2 (RTN2, also known as SPG12) helps shape ER tubeformation. Here, we use a new fluorescent ERmarker to show that theERwithin wild-typeDrosophilamotor nerve terminals forms a networkof tubules that is fragmented and made diffuse upon loss of theatlastin 1 ortholog atl. atl or Rtnl1 loss decreases evoked transmitterrelease and increases arborization. Similar to other HSP proteins, Atlinhibits bone morphogenetic protein (BMP) signaling, and loss of atlcauses age-dependent locomotor deficits in adults. These resultsdemonstrate a crucial role for ER in neuronal function, and identifymechanistic links between ER morphology, neuronal function, BMPsignaling and adult behavior.

KEY WORDS: Atlastin, ER, Reticulon, Neuron

INTRODUCTIONThe function of intracellular organelles is tightly coordinated withlocation within the cytoplasm. The endoplasmic reticulum (ER) isan interconnected network of narrow tubes and flattened cisternaeor sheets (Hu et al., 2011; Shibata et al., 2009, 2006; Terasakiet al., 2013; Westrate et al., 2015). In most cells, the ER is themost abundant subcellular organelle and extends elaborateprocesses throughout the cytoplasm. The ER membrane isformed into its tubular architecture by the action of structuralproteins within the reticulon, REEP and DP1 family (English andVoeltz, 2013; Hu et al., 2008; Shibata et al., 2009; Voeltz et al.,2006; Yang and Strittmatter, 2007). The members of this diversefamily of proteins share a common protein motif called thereticulon homology domain (RHD). The hydrophobic ∼200-amino-acid RHD likely forms a helical hairpin structure thatintercalates four hydrophobic helical segments into the outerleaflet of the ER membrane to induce curvature and maintain atubular shape (Voeltz et al., 2006; Zurek et al., 2011). Manymembers of the Reticulon, REEP and DP1 family also contain anextended N-terminal segment ranging from a few hundred to athousand amino acids that likely provides additional functionality

(Di Sano et al., 2012). The nature of most of these secondaryfunctions remains to be revealed.

The large ER network also maintains luminal and membranecontinuity throughout the cytoplasm. This interconnected natureof the ER network is required for ER function and is maintainedby the ER membrane fusion GTPase atlastin (Orso et al., 2009),which is a member of the fusion dynamin-related protein family(fusion DRP) (McNew et al., 2013; Moss et al., 2011; Pendin et al.,2011).

The ER is closely associated with and functionally connected tothe plasma membrane. This connection is often associated withthe management of Ca2+ stores in the ER lumen. The ER proteinSTIM1 diffuses through the ER membrane to find bindingpartners in the plasma membrane including the Orai channel(Jozsef et al., 2014; Soboloff et al., 2012). This set of protein–protein associations works to restore ER Ca2+ through the store-operated Ca2+ channel system. ER–plasma-membrane contact sitesare also generated by the association of ER-integral extendedsynaptotagmins (E-Syt) with phospholipids in the plasmamembrane (Fernández-Busnadiego et al., 2015; Giordano et al.,2013; Schauder et al., 2014) as well as proteins like junctophillinsin certain cell types (Helle et al., 2013; Stefan et al., 2013).

Most recently, the ER has been found to be stably associated withendosomal structures (Raiborg et al., 2015a; Rowland et al., 2014).In this circumstance, the specific proteins on each surface thatinteract remain to be precisely defined, but the consequence of theinteraction is functional segregation of certain cargoes within theendosome that permits regulated sorting into membranesubdomains prior to an ER-directed membrane fission event(Raiborg et al., 2015a,b; Rowland et al., 2014).

ER structure appears to be crucially important for cell functiongiven that human disease results when components that control thisstructure are compromised by mutation. The hereditary spasticparaplegias (HSPs) are a group of related genetic disorders causedby mutations in any of more than 70 genes, denoted SPG1 to SPG72(Lo Giudice et al., 2014; Noreau et al., 2014). Lower limb weaknessand spasticity represent two prominent clinical features of thesediseases, which occur as a consequence of dysfunction ordegeneration of the upper motor neurons (Blackstone et al.,2010). The observation that atlastin 1 (ATL1) and reticulon 2(RTN2) are HSP genes responsible for SPG3A and SPG12,respectively, implicates ER morphology in the neuronaldysfunction that causes HSPs.

The properties of three additional HSP genes, spartin (SPG20),spastin (SPAST, also known as SPG4) and NIPA1 (also known asspichthyin or SPG6), implicate receptor trafficking through theendocytic system in HSP neuronal dysfunction. For example, loss ofspartin attenuates both ligand-stimulated EGF receptor uptake(Bakowska et al., 2007) as well as depolarization-stimulated FM1-43 uptake (Nahm et al., 2013), whereas loss of spastin increasesReceived 15 December 2015; Accepted 17 February 2016

1Department of BioSciences, Program in Biochemistry and Cell Biology, RiceUniversity, Houston, TX 77005, USA. 2CNR, Neuroscience Institute, 35121 Padova,Italy. 3E. Medea Scientific Institute, 31015 Conegliano, Italy.*These authors contributed equally to this work

‡Authors for correspondence (stern@rice.edu; mcnew@rice.edu)

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endosome tubule number and alters transferrin receptor sorting(Allison et al., 2013). NIPA1 is also located in endosomes andpromotes the endocytosis of receptors for bone morphogeneticprotein (BMP) (Tsang et al., 2009).Phenotypic analysis of mutations in the HSP orthologs of model

systems has provided additional clues to the cellular function ofthese proteins. In Drosophila, loss of spastin, spartin andspichthyin confers a similar, but not identical, set of phenotypesincluding stabilized microtubules, increased synaptic boutonnumber and decreased evoked transmitter release at the larvalneuromuscular junction (NMJ), age-dependent locomotor deficitsand increased BMP signaling at the larval NMJ (Nahm et al.,2013; Ozdowski et al., 2011; Sherwood et al., 2004; Wang et al.,2007). These shared phenotypes might reflect disruption of acommon pathway in endocytic receptor trafficking in these mutants.Some of these phenotypes, such as stabilized microtubules, age-dependent locomotor deficits and increased synaptic boutonnumber, are also observed in flies lacking atlastin (atl) (Lee et al.,2009, 2008).Here, we extend this phenotypic analysis of altered atl and

reticulon-like 1 (Rtnl1) activities in Drosophila. We show that theER in motor nerve terminals from wild-type larvae forms a networkof tubules that resembles a ‘basket’, but is diffuse in larvae lackingatl. We find that neuronal RNA interference (RNAi)-mediatedknockdown of either atl or Rtnl1 increases arborization at the larvalneuromuscular junction and decreases evoked transmitter releasefrom larval motor nerve terminals, and that elevated bath [Ca2+]fully rescues these transmitter release phenotypes. We also showthat atl is required only in motor neurons to affect transmitterrelease, whereas Rtnl1 is required additionally in the target muscleand peripheral glia. Finally, we show that loss of atl increases BMPsignaling in larval motor neurons and causes age-dependentlocomotor deficits in adults. Thus, loss of atl and Rtnl1 confersphenotypes similar, but not identical, to each other as well as tomutants defective in spartin, spastin and spichthyin. Our resultsdemonstrate specific mechanistic links between ER morphologyand several aspects of neuronal anatomy and function.

RESULTSChemical fixation disrupts ER morphology in Drosophilamotor neurons, muscles and S2 cellsER structure has been difficult to examine in Drosophila. Toaddress this issue, we developed improved reagents for in vivoimaging of the ER. First, we utilized a mutant GFP, termedSuperfolder GFP (sfGFP) (Aronson et al., 2011; Costantini et al.,2015; Pédelacq et al., 2006), that has been found to fold properly inoxidizing environments such as the ER lumen. This protein trafficsto the ER because of addition of theDrosophilaBiP signal sequenceand is retained by addition of the HDEL ER retention signal fromcalreticulin (BiP–sfGFP–HDEL). Next, we introduced an ERmembrane marker containing Sec61β linked to the fluorescentmarker tdTomato. In Fig. 1, we show the ER labeled by BiP–sfGFP–HDEL expressed in the motor neuron cell body (Fig. 1A),the neuromuscular junction (NMJ) (Fig. 1B), larval body wallmuscle (Fig. 1C) and cultured S2 cells (Fig. 1D).Previous results have suggested that organelle structure in vivo is

labile to fixation (Johnson et al., 2015), and therefore an accuratedepiction of ER structure might require imaging in live cells. Todetermine whether ER structures in Drosophila larvae or S2 cellswere similarly labile to fixation, we compared ER structure in livecells with the fixed tissue. We found that very mild fixation (4%formaldehyde for 5 min with no permeabilization) significantly

altered ER structure in all tissues that were examined (Fig. 1). Uponfixation, ER tubules appeared to fragment into discrete punctae andthe uniformly labeled tubules seen in live imaging converted into a‘beads on a string’ morphology consistent with ruptured tubules. Itis possible that the ER tubules are maintained under stress byassociation with the underlying cytoskeletal network and thatchemical fixation disrupts this mechanical tension. Regardless ofthe mechanism, these observations reveal the disruptive effect oftissue fixation on the ER in vivo. For this reason, all subsequent ERimaging exhibited here was performed on live cells.

BiP–sfGFP–HDEL marks the ERWe investigated ER anatomy in larval motor nerve terminals inmore detail. To orient the BiP–sfGFP–HDEL signal with the plasmamembrane, we co-expressed BiP–sfGFP–HDEL with the plasmamembrane marker myr::tdTomato. We found that BiP–sfGFP–HDEL was located immediately adjacent to the myr::tdTomatosignal (Fig. 2A–C), indicating that the ER baskets in motor nerveterminals directly underlie the plasma membrane. We also co-expressed BiP–sfGFP–HDEL along with the ER membrane markertdTomato–Sec61β. As expected, these two markers showedextensive overlap (Fig. 2D–F), thus validating both transgenes asER markers.

The atl2 null mutation alters ER structure inmotor axons andpresynaptic boutonsatl encodes an ER fusion GTPase (Orso et al., 2009), and thus weanticipated that, by preventing ER fusion, the null mutation atl2

would fragment the ER. To test this prediction, we compared ERmorphology in motor neuron cell bodies, motor axon initialsegments, and motor nerve terminals in control and atl2 animals(Fig. 3A). We found little effect on ER morphology in cell bodies(Fig. 3Bi versus Fig. 3Bii), but in axon initial segments, we foundthat the ER crossbridges found in wild-type were almost completelyeliminated in atl2 cells, and that the long straight ER tubulesobserved in wild-type became wavy (Fig. 3Biii,iv). In motor nerveterminals, we found that atl2 eliminated the basket structures andcaused a diffuse ER signal, which we attribute to ER fragmentation(Fig. 3Ci,ii).

To quantify the effects of atl2 on ER morphology in nerveterminals, we reasoned that the BiP–sfGFP–HDEL signal in atl2

larvae would be uniformly distributed across the bouton and thus afrequency histogram for the pixel intensities within each terminalwould exhibit a Gaussian distribution. In contrast, we anticipatedthat pixel intensities in wild-type nerve terminals would comprise asmall number of bright pixels indicating the tubules, and a largernumber of dark pixels indicating voids between tubules. Thus, wepredicted that a frequency histogram for the pixel intensities fromwild-type would be positively skewed. We compared frequencyhistograms of pixel intensities, normalized to mean intensity,between wild-type and atl2 and found, as predicted, a strong positiveskew in wild-type (Fig. 3Ciii) but a more Gaussian distribution inatl2 (Fig. 3Civ). Furthermore, the mode and median pixel intensitieswere close to the mean in atl2, consistent with a Gaussiandistribution, but were less than the mean in wild-type, consistentwith a positive skew (Fig. 3D). In addition, we compared line scansacross nerve terminals in atl2 and wild-type. Whereas wild-type linescans showed multiple peaks, corresponding to tubules within thebaskets, atl2 line scans were uniform, consistent with a diffuse ERsignal (Fig. 3Cv,vi).

Next, we determined whether the atl2 ER fragmentation could berecapitulated by targeted atl knockdown in motor neurons. Thus, we

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expressed both an atl RNAi transgene and the dominant-negativeatlK51A allele (Orso et al., 2009) in neurons and found a similardecrease in ER tubule crossbridges in axon initial segments(Fig. 4D–F) and a similar diffuse ER signal at motor nerveterminals (Fig. 4G–I). These results demonstrate that maintenanceof the correct ER structure in motor neurons requires atlastinactivity.In contrast to atl2, the null Rtnl11 allele conferred no detectable

abnormalities in ER structure in motor neuron cell bodies, axons ornerve terminals (data not shown).

Impaired evoked transmitter release in atl2 and Rtnl11 nullmutantsAltered levels of several HSP proteins, including Spartin andSpastin, alter transmitter release from Drosophila motor nerveterminals (Nahm et al., 2013; Ozdowski et al., 2011; Sherwoodet al., 2004). We tested the possibility that atl2 and Rtnl11 mightsimilarly affect transmitter release. We used the larvalneuromuscular preparation (Jan and Jan, 1976; Stewart et al.,1994) to measure synaptic transmission and found that eachmutation indeed decreased evoked transmitter release, using the

Fig. 1. Chemical fixation disrupts the ER network in motor neurons, muscles, and S2 cells. Representative confocal slices through the center (A) andperiphery (A′) of live and fixed wild-type motor neuron (MN) cell bodies in the ventral nerve cord. (B) ER in wild-type larval boutons under live and fixed conditions.(B′) Magnification of boxed regions in B. (C) ER in wild-type muscle 6 from segment A3 under live and fixed conditions. (C′) Magnification of boxed regions inC. (D) ER in S2 cells under live and fixed conditions. (D′) Magnification of boxed regions in D. Chemical fixation was achieved by a 5 min exposure to 4%paraformaldehyde. ER was imaged using BiP–sfGFP–HDEL. Scale bars: 5 µm, except B′ (2 µm).

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consequent muscle depolarization termed excitatory junctionalpotential (EJP) as a readout. Fig. 5A and B show averaged EJPs inwild-type versus atl2, respectively, at the four indicated bath [Ca2+].At a bath [Ca2+] of 0.6 mM, both atl2 and Rtnl11 decreasetransmitter release almost two-fold (Fig. 5C).To confirm that these transmitter release phenotypes were caused

by loss of atl and Rtnl1, we determined whether expression of UAS-atl+ and UAS-Rtnl1+ transgenes would be sufficient to restore wild-type transmitter release to atl2 and Rtnl11. We found that the atl2

transmitter release phenotype was fully rescued by expression ofUAS-atl+ driven by the weak ubiquitous Gal4 driver arm-Gal4(Fig. 5D). In contrast, the Rtnl11 transmitter release phenotype wasrescued only partially by arm-Gal4-driven UAS-Rtnl1+; this partialrescue most likely reflected weak expression of the Rtnl1+ transgenebecause, as shown below, driving UAS-Rtnl1+ expression with thestronger Da-Gal4 driver elicited complete rescue (see Fig. 7B).Previous reports have indicated that nerve terminal ER is capable

of controlling transmitter release by influencing cytoplasmic [Ca2+](Emptage et al., 2001; Liang et al., 2002; Llano et al., 2000). Wewondered whether the decreased evoked transmitter release in atl2

and Rtnl11 might reflect attenuation of the increased cytoplasmic[Ca2+] caused by nerve stimulation. If so, then we predicted that thisdecreased transmitter release would be rescued by elevated bath[Ca2+], at which Ca2+ is less limiting for transmitter release (Wonget al., 2014). For both atl2 and Rtnl11, we found that as we increasedbath [Ca2+], transmitter release became progressively similar to thatin wild type (Fig. 5E, which shows quantal content, corrected fornonlinear summation and normalized to wild type). Elevated bath[Ca2+] completely rescued the Rtnl11 transmitter release phenotype,but rescued the atl2 phenotype only partially. In addition, we foundthat RNAi-mediated knockdown of either atl or Rtnl1 in neuronssimilarly failed to decrease transmitter release at elevated bath[Ca2+] (Fig. 5F; efficacy of the atl and Rtnl1 RNAis is demonstratedin experiments described below). It is not known why elevated bath[Ca2+] rescues the atl RNAi knockdown phenotype completely, butatl2 only partially. However, it is possible that atl acts in non-neuronal tissues to affect transmitter release at elevated bath [Ca2+].In contrast to the strong effects of altered atl and Rtnl1 on EJP

amplitude, the amplitudes of ‘mini’ EJPs, generated by spontaneous

release of individual vesicles of transmitter, were not substantiallyaffected in atl2 or Rtnl11 larvae (Fig. 5G). The modest, albeitsignificant, increase in mini EJP amplitude observed in atl2 is likely agenetic background effect unrelated to the atl genotype, as mini EJPamplitude is not significantly affected byarm-Gal4-driven expressionof the atl+ transgene (data not shown), and elav-Gal4-drivenexpression of atl RNAi does not affect mini EJP amplitude (Fig. 5H).

It has been previously observed that atlastin and reticulon conferopposite effects on ER morphology. In particular, whereas atl2

fragments the ER (Orso et al., 2009), Rtnl1 knockdown by RNAi(O’Sullivan et al., 2012) converts ER tubes into sheets. We found amutual suppression of the evoked transmitter release phenotype in theRtnl1; atl2 double mutant (Fig. 5C,E), which supports the possibilitythat atlastin and reticulon have antagonistic effects onERmorphology.

We examined synapse architecture in atl2 and Rtnl11 mutants byperforming immunocytochemistry with antibodies directed at thepresynaptic active zone protein Bruchpilot and the postsynapticneurotransmitter receptor GluRII. We found no obvious structuralor organizational defects in the number or size of active zones or inthe pattern of GluRII immunoreactivity (data not shown).

atl acts in the motor neuron to control transmitter releaseNext, we wished to identify the tissues in which Atl and Rtnl1 arerequired to control neurotransmitter release. Because atl2 and Rtnl11

exhibited the greatest deficit in neurotransmitter release at the lowestbath [Ca2+] (Fig. 5), we performed the next set of measurements atthe low bath [Ca2+] of 0.1 mM, in a solution (HL3.1) conducive forlow [Ca2+] recordings.

Inhibiting atl by driving atl RNAi expression with the motorneuron driver D42-Gal4 (Fig. 6A, left, green diamonds) or the pan-neuronal driver elav-Gal4 (Fig. 6A, left, green squares) significantlydecreased EJP amplitude. Similarly, inhibiting atl by expressing thedominant-negative atlK51A with either D42-Gal4 (Fig. 6A, left,green inverted triangles) or the pan-neuronal driver nSyb-Gal4(Fig. 6A, left, green triangles) also significantly decreased EJPamplitude. These results indicate that atl activity is required in themotor neuron for wild-type evoked transmitter release.

We confirmed that the decrease in EJP amplitude representeddecreased transmitter release rather than decreased muscle

Fig. 2. The ER lumen marker BiP–sfGFP–HDEL colocalizes with the ER membranemarker tdTomato–Sec61β. Third-instar larvalmotor nerve terminals from segment A2 muscle4. (A–C) Representative confocal z-projectionsshowing the plasma membrane marker myr::tdTomato (A), the ERmarker BiP–sfGFP–HDEL(B) and the merged signals (C).(D–F) z-projections showing the ER membranemarker tdTomato–Sec61β (D), the ER lumenmarker BiP–sfGFP–HDEL (E) and the mergedsignals (F). All transgenes were driven by themotor neuron driver OK371-Gal4. Scale bars:5 µm.

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responsiveness to transmitter by analyzing the frequency of synaptictransmission failures. In the low [Ca2+] bath utilized in Fig. 6, themotor neuron responds to nerve stimulation with release of eitherno transmitter, which is called a failure, or one or two vesicles oftransmitter, which is called a success. The decreased transmitterrelease in the atl knockdown larvae was accompanied by adecreased frequency of successes (Fig. 6A, right), indicating thatatl knockdown decreases transmitter release.Transmitter release at the Drosophila neuromuscular junction

can be affected by the target muscle or the neighboring peripheralglia (Huang and Stern, 2002; Kerr et al., 2014; McCabe et al., 2003;Schmidt et al., 2012). RNAi-mediated atl knockdown in muscle orglia did not affect transmitter release or frequency of synapticfailures (Fig. 6B, left, green squares; Fig. 6B, left, green diamonds).

Overexpression of atl in the motor neuron impairsneurotransmitter releaseExpression of UAS-atl+ in Drosophila motor neurons causesexcessive fusion and expansion of the nuclear envelope and ERmembrane, and defects in the secretory pathway in both theneuron cell body and presynaptic terminal (Orso et al., 2009).Using the BiP–sfGFP–HDEL imaging reagent described above,we found that atl+ overexpression caused accumulation of largeER punctae in motor neuron cell bodies and axons (Fig. S1A).To determine whether atl+ overexpression affected transmitterrelease, we compared EJP amplitudes in atl+ overexpressingand control larvae at both low (0.1 mM) and high (1.5 mM) bath[Ca2+]. We found that atl+ overexpression significantly decreasedEJP amplitudes and the frequency of synaptic successes at low

Fig. 3. Loss of atl disrupts the tubular ER network in motor axons and motor nerve terminals of third-instar larvae. (A) Schematic of the regions of themotor neuron from which images were collected. (B) Representative central confocal slices of motor neuron cell bodies in control [OK371>BiP–sfGFP–HDEL](Bi) and atl2 [atl2, OK371>BiP–sfGFP–HDEL] (Bii) larvae. z-projections of motor axonswithin the ventral nerve cord in wild-type (control) (Biii) and atl2 (Biv) larvae.(C) z-projections of neuromuscular junctions from muscle 4 at segment A6 in wild-type (Ci) and atl2 (Cii) larvae. Average frequency histograms of pixelintensities taken from regions of interest around boutons for control (Ciii) and atl2 (Civ) (w1118 n=5, atl2 n=6; red and blue dashed lines represent the mode andmedian, respectively, of the histogram and are color coded with the analysis in D). Boxed regions in Ci and Cii are expanded in Ci′ and Cii′, respectively. Dashedlines in Ci′ and Cii′ represent the positions used for the linescans of pixel intensities for wild-type (Cv) and atl2 (Cvi). (D) Mode and median of pixel intensityfrequency histograms (mean±s.e.m.) are shown for five control and six atl2 images. P-values shown represent an unpaired Student’s t-test performed withKaleidaGraph. Scale bars: 5 µm, except for Ci′ and Cii′ (2 µm).

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bath [Ca2+] (Fig. S1B), and decreased corrected quantal contentat high bath [Ca2+] (Fig. S1C). Rtnl1+ overexpression alsodecreased transmitter release but to a lesser extent than atl+. Weconclude that maximum levels of evoked transmitter releaserequire specific ratios of Atl and Rtnl1 protein.

Rtnl1 regulates transmitter release from multiple tissues atthe NMJWe used both tissue-specific transgenic rescue and RNAi-mediated knockdown to identify the tissue(s) within whichRtnl1 functions to control transmitter release. First, we foundthat driving Rtnl1 RNAi expression with the pan-neuronaldriver elav-Gal4 (Fig. 7A, left, blue squares) significantlydecreased both evoked EJP amplitude and percentage ofsynaptic successes (Fig. 7A, right). This result demonstratesthat Rtnl1 is required in the motor neuron to affect transmitterrelease. We then determined whether Rtnl1 is required in theother cell types of the tripartite synapse and found that RNAi-

mediated Rtnl1 knockdown in either muscle (Fig. 7A, left, bluediamonds) or glia (Fig. 7A, left, blue triangles) alsosignificantly decreased evoked EJP amplitude and synapticsuccess rate (Fig. 7A, right). Thus, Rtnl1 activity is required inall three cell types of the tripartite synapse for proper evokedtransmitter release.

We next determined whether Rtnl1 activity in these threecell types was sufficient for proper evoked transmitter release.First, we verified effectiveness of the Rtnl1+ rescue construct bydetermining that ubiquitous expression of the Rtnl1+ transgene,driven by the Da-Gal4 driver, was sufficient to rescue bothevoked EJP amplitude (Fig. 7B, left) and frequency of synapticsuccesses of Rtnl11 (Fig. 7B, right). As expected from the RNAiresults described above, expression of Rtnl1+ driven pan-neuronally, in the motor neuron alone, the muscle alone, orboth the motor neuron and muscle, was not sufficient to rescuethe EJP amplitude or synaptic success phenotypes of Rtnl11

(Fig. 7B and data not shown). However, simultaneous expression

Fig. 4. Expression of UAS-atlRNAi or UAS-atlK51A disrupts the tubular ER network inmotor axons and presynaptic boutons.(A–C) ER in motor neuron (MN) cell bodies forcontrol [nSyb>BiP-sfGFP-HDEL] (A), atl RNAi[nSyb>BiP-sfGFP-HDEL, atlRNAi] (B), anddominant-negative Atl [nSyb>BiP-sfGFP-HDEL, atlK51A] (C). (D–F) ER in motor axons forcontrol (D), atl RNAi (E) and dominant-negative(DN) atlK51A (F). (G–I) ER in presynapticboutons for control (G), atl RNAi (H)and dominant-negative atlK51A (I).(G′–I′) Magnification of boxed regions for wild-type (G′), atl RNAi (H′), and dominant-negativeatlK51A (I′). Scale bars: 5 µm, except for G′,H′and I′ (2 µm).

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of Rtnl1+ in the motor neuron, muscle and peripheral glia wassufficient to significantly restore proper EJP amplitude andsynaptic success frequency to Rtnl11 (Fig. 7B). We conclude thatRtnl1 activity in the motor neuron, target muscle andneighboring peripheral glia is sufficient as well as necessary tomaintain proper synaptic transmission.

atl2 increases BMP signaling in larval motor neuronsBMPs are a family of secreted ligands that control a wide variety oforganismal functions. Following binding to membrane receptors,BMPs trigger the phosphorylation and activation of the Madtranscription factor (Miyazono et al., 2010), which is assayed withantibodies specific to the active phosphorylated (p)Mad. In addition

Fig. 5. atl2 and Rtnl11 decrease evokedneurotransmitter release. Average EJP traces for wild-type (w1118) (A) and atl2 (B) larvae at 0.4, 0.6, 1.0, and1.5 mM Ca2+. In ascending [Ca2+], for w1118 n=7, 17, 7,and 7 and for atl2 n=7, 23, 7, and 7. (C) Mean±s.e.m. EJPamplitude for w1118, atl2, Rtnl11, and Rtnl11; atl2. P-valuesshown represent a one-way ANOVA using a Fisher’s LSDpost hoc test performed with KaleidaGraph. (D) Mean±s.e.m. EJP amplitude for atl2; arm>UAS-atl+, Rtnl11;arm>UAS-Rtnl1+, and their respective controls using theubiquitous arm-Gal4 driver. P-values shown represent anunpaired Student’s t-test performed with KaleidaGraph.Recordings for C and D were performed at 0.6 mM Ca2+.(E) Mean±s.e.m. corrected quantal content for w1118,Rtnl11, atl2, and Rtnl11; atl2. In ascending [Ca2+], w1118

n=7, 17, 7, and 7; atl2 n=7, 23, 7, and 7; Rtnl11 n=7, 10, 7,and 7; Rtnl11; atl2 n=7, 7, 7, and 7. Resting membranepotentials (RMPs) were not significantly different amongthe four genotypes at bath [Ca2+] of 0.4 mM or 1.5 mM. Atbath [Ca2+] of 0.6 mM or 1.0 mM, RMPs of wild-type andatl2 were significantly different (P=0.0266, wild-type morehyperpolarized, at 0.6 mM [Ca2+], and P=0.0316, atl2

more hyperpolarized, at 1.0 mM [Ca2+]). RMP of Rtnl11;atl2 was significantly different from other genotypes at abath [Ca2+] of 1.0 mM, Rtnl11; atl2 more hyperpolarized.(F) Mean± s.e.m. corrected quantal content for elav>dcr,elav>dcr, atlRNAi, and elav>dcr, Rtnl1RNAi at 1.5 mM[Ca2+]. (G) Mean±s.e.m. mEJP amplitudes for w1118, atl2,Rtnl11, and Rtnl11; atl2, pooled from all four [Ca2+].(H) Mean± s.e.m. mEJP amplitudes for the indicatedgenotypes collected at 1.5 mM bath [Ca2+]. P-values in D,F,G represent an unpaired Student’s t-test performed withKaleidaGraph. Recordings were performed in HL3 frommuscle 6 in segment A6. Each scattergram data point isthe average of the first 20 EJP responses (successes andfailures) recorded for each larva.

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to developmental functions,Drosophila BMP ligands released frommuscle, peripheral glia or the motor neuron itself regulatetransmitter release and synaptic growth in motor neurons(Fuentes-Medel et al., 2012; McCabe et al., 2003). Many HSPproteins, including Drosophila Spichthyin and Spartin, mammalianspastin and zebrafish atlastin, inhibit BMP signaling (Fassier et al.,2010; Nahm et al., 2013; Wang et al., 2007), possibly throughaltered BMP receptor trafficking. To determine whether Atl mightsimilarly inhibit BMP signaling, we measured nuclear pMad withinmotor neuron nuclei of atl2 third-instar larvae. We observed asignificant increase in nuclear pMad in atl2, which was rescued byboth weak ubiquitous (Fig. 8C, green squares) or motor neuron-specific expression of atl+ (Fig. 8C, green diamonds). Thus, Atlbehaves similarly to other HSP proteins as an inhibitor of BMPsignaling.

Neuronal loss of atl and Rtnl1 causes axon terminalovergrowthMutations in the Drosophila HSP orthologs described aboveincrease synaptic bouton number at the third-instar larval NMJ. Inaddition, Lee et al. (2009) have reported that bouton number is also

increased in atl2 NMJs, and that this increase is rescued by musclebut not neuronal atl+ expression. We found that pan-neuronalknockdown of either atl or Rtnl1 significantly increased boutonnumber, and that this increase was suppressed when both geneswere knocked down simultaneously (Fig. S2). These results suggestthat Atl and Rtnl1 are required within the motor neuron to restrainaxon outgrowth.

Neuronal knockdown of atl causes age-dependentlocomotor deficitsHSP patients share a characteristic increase in spasticity andweakening of the lower extremities with age.Drosophilamutant forany of several HSP orthologues similarly display behavioral deficits(Lee et al., 2008; Nahm et al., 2013; Sherwood et al., 2004). Weused two assays to determine whether pan-neuronal atl inhibitiongenerated locomotor deficits. First, we measured the time requiredfor elav>Dcr, atlRNAi adults to climb 6 cm, and second, we used theiFly tracking system (Kohlhoff et al., 2011) to measure walkingvelocity. We observed age-dependent locomotor deficits with eachassay (Fig. S3). It is unlikely that these locomotor deficits resultdirectly from the deficits in larval synaptic transmission that we have

Fig. 6. Neuronal expression of UAS-atlRNAi and UAS-atlK51A decreasesevoked neurotransmitter release.(A) Mean±s.e.m. EJP amplitude andcorresponding percentage success for wild-type (w1118), atl2, UAS-atlRNAi and UAS-atlK51A. Drivers include the pan-neuronalelav-Gal4 and nSyb-Gal4 and the motorneuronal D42-Gal4. (B) Mean±s.e.m. EJPamplitude and percentage success formuscle (Mef2-Gal4) and glial (Gli-Gal4)expression of UAS-atlRNAi and UAS-atlK51A.Recordings for A and B were made in HL3.1at 0.1 mM Ca2+ from muscle 6 in segmentA6. Each scattergram data point is theaverage of the first 20 EJP responses(successes and failures) recorded for eachlarva. P-values shown represent anunpaired Student’s t-test performed withKaleidaGraph.

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documented. Rather, these locomotor deficits might reflect age-dependent degeneration of specific populations of neurons as hasbeen reported previously for an atl hypomorph (Lee et al., 2008).

DISCUSSIONThe role of ER morphology in nervous system function andanatomyMutations in two genes that affect ER morphology, atlastin (ATL1)and reticulon 2 (RTN2), cause two forms of hereditary spasticparaplegia (HSP), which result in progressive limb weakness,spasticity and degeneration of the longest motor axons (Blackstoneet al., 2010). These observations suggest that altered ERmorphology is causal for motor axon dysfunction, but themechanisms underlying these dysfunctions are unclear. Here, we

use Drosophila to evaluate the nervous system deficits caused byaltered atl and Rtnl1 activity. Using a new fluorescent ER imagingreagent, we show that the ER in wild-type motor nerve terminals ispresent as a network of tubules, termed ‘baskets’, underlying theplasma membrane, and that these baskets are eliminated in larvaelacking or overexpressing atl. We also show that loss of either atl orRtnl1 increases arborization and decreases evoked transmitterrelease, and that evoked release is restored to normal by elevatedbath [Ca2+]. Finally, we show that atl loss increases signalingthrough the bone morphogenetic protein (BMP) pathway and causesage-dependent declines in adult locomotion. This set of phenotypesis also exhibited by Drosophila mutant for the HSP orthologs ofspartin, spastin and spichthyin, as well as for spinster and nervouswreck, which encode regulators of receptor trafficking through

Fig. 7. Rtnl1 affects neurotransmitter release frommultiple tissues at the NMJ. (A) Mean±s.e.m. EJP amplitude and corresponding percentage success forwild-type (w1118), Rtnl11, and UAS-Rtnl1RNAi expression in neurons, muscles and glia using elav-Gal4, Mef2-Gal4 and Gli-Gal4, respectively. (B) Mean±s.e.m.EJP amplitude and corresponding percentage success for w1118, Rtnl11 and expression of UAS-Rtnl1+ in Rtnl11 mutants. UAS-Rtnl1+ was expressedubiquitously, in motor neurons and inmuscles using da-Gal4,D42-Gal4 andMef2-Gal4, respectively.OK371,Mef2 andGli-Gal4 denote concurrent expression ofUAS-Rtnl1+ from motor neurons, muscles and glia. Recordings for A and B were made in HL3.1 at 0.1 mM Ca2+ from muscle 6 in segment A6. Each scattergramdata point is the average of the first 20 EJP responses (successes and failures) recorded for each larva. P-values shown represent an unpaired Student’s t-testperformed with KaleidaGraph.

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endosomes (Nahm et al., 2013; O’Connor-Giles et al., 2008;Ozdowski et al., 2011; Sherwood et al., 2004; Sweeney and Davis,2002;Wang et al., 2007). The possible involvement of the ER in thisreceptor trafficking pathway will be discussed.

Effects of atl loss or overexpression on ER morphology inmotor neuronsWe made two adjustments to improve visualization of the ER. First,we introduced into flies a transgene that encoded an ER-localizedsuperfolder GFP, which was optimized for efficient folding in theER. Second, based on previous results indicating that the ER as wellas the lysosomal tubule network is labile to fixation (Johnson et al.,2015), we imaged ER in live tissues. Using these approaches, weshowed that the ER is present within the axon initial segments ofmotor neurons as a polygonal structure with numerous crossbridges(three-way junctions), and in motor nerve terminals as a network oftubules that we term ‘baskets’, which underlie the plasmamembrane. We also showed that these structures are disruptedby either loss of or overexpression of atl. In particular, atloverexpression causes the aberrant appearance of large punctae inmotor neuron cell bodies or axon initial segments. In contrast, atlloss decreases the number of crossbridges in the axon initialsegment, leading to excessively long tubules. A similar appearancewas noted previously (Hu et al., 2009; Orso et al., 2009) andattributed to deficits in fusion of orthogonal ERmembranes. Loss ofatl also disrupts nerve terminal baskets and appears to cause ERfragmentation. It is possible that the transition from tubules tobaskets as the ER moves from the interbouton region to boutonsoccurs through ER fragmentation followed by Atl-dependentreassembly. In this view, loss of atl would prevent thisreassembly, thus causing the fragmented ER that we observe.

Deficits in evoked transmitter release in larvae lacking atl orRtnl1 are rescued by elevated bath [Ca2+]Evoked transmitter release deficits in both atl2 and Rtnl11 mutants,and in pan-neuronal atl or Rtnl1 knockdown larvae, were rescuedpartially or completely by elevated bath [Ca2+]. These resultsindicate that loss of atl or Rtnl1 decreases evoked transmitter releaseat low bath [Ca2+] through causing deficits in evoked increases incytoplasmic [Ca2+]. Insufficient Ca2+ influx could result fromattenuated action potentials, which would decrease the opening ofvoltage-gated Ca2+ channels, decreases in number of plasmamembrane Ca2+ channels or decreased Ca2+ release from the ER.

Given the role of atlastin and the reticulons as ER-shapingmolecules, effects on ER Ca2+ release would be the most directexplanation for this Ca2+ phenotype. ER-localized Ca2+ releasechannels such as the inositol 1,4,5-trisphosphate (IP3) receptor, theryanodine receptor and the TRPV1 channel play key roles in evokedneurotransmitter release (Emptage et al., 2001; Liang et al., 2002;Llano et al., 2000; Wong et al., 2014). In addition, dominant-negative mutations in the Drosophila ER-localized Ca2+ pumpSERCA decrease evoked transmitter release by∼50% (Sanyal et al.,2005), which is consistent with the possibility that ER-derived Ca2+

contributes significantly to the Ca2+ required to trigger transmitterrelease.

Rtnl1 affects evoked neurotransmitter release frommultipletissuesUnlike Atl, which appears to affect evoked transmitter release fromneurons alone, Rtnl1 is required in neurons, muscles and peripheralglia for correct evoked transmitter release. This finding is consistentwith previous data demonstrating that proper synaptic transmissionrequires intercellular signaling among these three cell types. Inparticular, loss of activity within the peripheral glia of the kinesinheavy chain gene or the inebriated-encoded neurotransmittertransporter alters evoked transmitter release (Huang and Stern,2002; Schmidt et al., 2012). In addition, the peripheral glia secrete atleast two proteins, the TGF-β ligand Maverick and Wingless/Wnt,that regulate synaptic function (Fuentes-Medel et al., 2012; Kerret al., 2014). The muscle, in turn, secretes the BMP ligand Gbb toregulate both evoked transmitter release and motor neuronarborization (McCabe et al., 2003). It is possible that loss of Rtnl1affects transmitter release from glia or muscle by perturbingsecretion of these or other regulators.

Pan-neuronal atl knockdown causes progressive adultlocomotor deficits during agingThe most prominent clinical symptom in HSP patients isprogressive, age-dependent locomotor difficulties. Drosophilamutant for any of several HSP orthologs, including spartin,spastin, atl and Rtnl1, as well as spinster, exhibit similar age-dependent locomotor deficits or lifespan deficits (Dermaut et al.,2005; Lee et al., 2008; Nahm et al., 2013; O’Sullivan et al., 2012;Orso et al., 2009; Sweeney and Davis, 2002). Here, we showlocomotor impairment in adults with neuronal-specific atlknockdown. These results indicate a requirement for atl in

Fig. 8. atl2 increases BMP signaling in motor neurons. Representative confocal images of w1118 (A) and atl2 (B) motor neuron nuclei from third-instar larvaelabeled with anti-pMad antibody. Scale bar: 10 µm. (C) Mean±s.e.m. pMad levels for w1118, atl2, atl2; arm>+, atl2; arm>atl+, atl2; OK6>+, and atl2; OK6>atl+ inmotor neuron nuclei. Each scattergram data point is the average nuclear signal intensity per area for a single larva. P-values shown represent an unpairedStudent’s t-test performed with KaleidaGraph.

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neurons for proper locomotion but do not rule out crucial roles foratl in other tissues as well.

A potential role for the ER in endocytic receptor traffickingMutants in Drosophila orthologs of several HSP genes, includingspartin, spastin and spichthyin, and the additional related genesspinster and nervous wreck share a common set of nervous systemphenotypes, including increased arborization and BMP signaling atthe larval NMJ, decreased evoked transmitter release and locomotordeficits (Nahm et al., 2013; O’Connor-Giles et al., 2008; Ozdowskiet al., 2011; Sherwood et al., 2004; Sweeney and Davis, 2002;Wang et al., 2007) (note that not all phenotypes have been reportedfor each mutant). The encoded proteins localize to variouscompartments within the endocytic receptor trafficking pathway(Allison et al., 2013; Edwards et al., 2009; O’Connor-Giles et al.,2008; Sweeney and Davis, 2002; Wang et al., 2007). In fact, theincreased BMP signaling in several of these mutants has beenattributed to trafficking defects of the BMP receptor Wishfulthinking (Wit). We have shown that atl loss confers these samephenotypes, raising the possibility that atl acts in the endocyticreceptor trafficking pathway as well. Although the ER is not knownto play prominent roles in this pathway, a recent report hasdemonstrated that the ER is required for endosome fission in COScells, and, in fact, the ER selects the location of fission (Rowlandet al., 2014). In addition, it has been found that this process isinhibited by overexpression of Rtn4a, which, similarly to atl loss,elongates ER tubules and inhibits formation of crossbridges(Rowland et al., 2014). Thus, loss of atl could impact on thereceptor trafficking pathway in Drosophila nerve terminals bysimilarly preventing endosome fission.The variety of phenotypes exhibited in common by the mutants

described above raises the possibility that certain phenotypes mighthave causal relationships with others. The subcellular locations ofthese proteins suggest that they might directly affect receptortrafficking. If so, then the increased BMP signaling, as aconsequence of altered Wit trafficking, might be the direct cause ofthe increased arborization and locomotor deficits. The phenotypesconferred by direct activation of the BMP pathway in neurons areconsistent with this possibility (McCabe et al., 2003; Nahm et al.,2013).However, the increasedBMP signaling is unlikely to cause thedecreased transmitter release, as decreased BMP signaling, ratherthan increased BMP signaling, decreases evoked transmitter release(McCabe et al., 2003). We suggest that trafficking of receptors inaddition toWit are altered in themutants described above, and it is thealtered signaling of these additional receptors that is at least partlyresponsible for the transmitter release phenotype. Drosophila motornerve terminals express a cholecystokinin-like receptor (CCKLR), atoll-like receptor, a metabotropic glutamate receptor (mGluRA) andlikely the insulin receptor (Ballard et al., 2014; Bogdanik et al., 2004;Chen and Ganetzky, 2012; Howlett et al., 2008). Loss of mGluRAincreases evoked transmitter release (Bogdanik et al., 2004; Howlettet al., 2008), raising the possibility that increased mGluRA signalingmight decrease transmitter release,which could explain the decreasedtransmitter release observed in these receptor trafficking mutants.

MATERIALS AND METHODSDrosophila stocks and mediumThe following lines were obtained from the Bloomington Drosophila StockCenter at Indiana University: w1118 (#3605), arm-Gal4 (#1560), elav-Gal4(#458), elav-Gal4; UAS-Dcr-2 (#25750), nSyb-Gal4 (#51635), OK371-Gal4 (#26160), Mef2-Gal4 (#27390), UAS-Dcr-2 (#24650), UAS-Dcr-2(#24651), and UAS-myr::tdTomato (#32221). The da G32 (#108252) Gal4

driver line was provided by the Drosophila Genetic Resource Center. UAS-Rtnl1RNAi (#7866) was obtained from the Vienna Drosophila ResourceCenter. OK6-Gal4 (Aberle et al., 2002) was provided by Hermann Aberle(Heinrich Heine University, Düsseldorf, Germany). D42-Gal4 (Yeh et al.,1995) was provided by Thomas Schwarz (Children’s Hospital Boston, F. M.Kirby Neurobiology Center, Boston, MA).Gli-Gal4 (Sepp and Auld, 1999)was provided by Vanessa Auld (The University of British Columbia,Department of Zoology, Vancouver, British Columbia, Canada). UAS-atl+,UAS-atlK51A, and UAS-atlRNAi were as described previously (Orso et al.,2009). atl2 (Lee et al., 2009) and Rtnl11 (Wakefield and Tear, 2006) were asdescribed previously. UAS-Rtnl1+ was provided by Andrea Daga.

All fly stocks were maintained on cornmeal and agar medium (6%dextrose, 6.8% cornmeal, 1.2% yeast, 0.72% agar, 2% methyl 4-hydroxybenzoate) at room temperature (∼23°C) or 25°C.

Plasmid constructionTo construct pJM952 (pAc5/BiP-sfGFP-HDEL), Superfolder GFP (sfGFP)in pEGFP-N1 (Aronson et al., 2011)] was provided by Erik Snapp (AlbertEinstein College of Medicine, Department of Anatomy & StructuralBiology, Bronx, NY). The Drosophila BiP signal sequence(MKLCILLAVVAFVGLSLG-RS) was then fused to sfGFP with a C-terminal ER retention signal (-HDEL) fromDrosophila Calreticulin (Smith,1992) in pAc5.1/V5-His A (Invitrogen). To construct pJM1033 (UAS-BiP-sfGFP-HDEL), pUASTattB (Bischof et al., 2007) was provided by KonradBasler (University of Zürich, Institute of Molecular Life Sciences, Zürich,Switzerland). The BiP-sfGFP-HDEL cassette from pJM952 was thenmoved to pUASTattB. To construct pJM1072 (UAS-tdTomato-dSec61β),tdTomatowas fused toDrosophila Sec61β (DrosophilaGenomics ResourceCenter cDNA clone #RE18615) in a 20X-UAS-IVS vector derived frompJFRC7 (Addgene #26220).

Drosophila stock constructionpJM1033 and pJM1072 were injected into attP2 and VK37 embryos byGenetivision (Houston, TX). Injected flies were backcrossed twice to w1118

flies and stocks carrying insertions on chromosome two (VK37) and three(attP2) were established. The UAS-Rtnl1 line was produced with the Rtnl1-PB isoform cDNA (LD14068). The cDNAwas subcloned in-frame with anHA tag in the pUAST vector, and transgenic lines were generated bystandard microinjection.

Live larval imagingWandering third-instar larvae, reared at 25°C, were dissected in HL6 buffer(Macleod et al., 2002), a complex buffer designed for imaging metabolicallyactive fly larvae, with 0 mM CaCl2 and 7 mM monosodium glutamate on35-mm petri dish lids completely filled with Sylgard 184 (ElectronMicroscopy Sciences). Minutien pins (0.1-mm diameter, Fine ScienceTools), bent to 90° and trimmed, were used to secure larvae to the Sylgard.Round 25-mm coverslips (#1 thickness) were cut in half with a diamondknife and attached to the Sylgard on either side of dissected larvae to form achannel. Square 22-mm coverslips (#1.5 thickness) were placed over thelarvae and secured to the bottom coverslips with nail polish. Larvae wereimaged on a Zeiss LSM 710 inverted confocal microscope using a Plan-Apochromat 63× (1.40 NA) oil immersion objective. GFP was excited witha 488-nm argon laser and emitted light between 493 and 522 nm wascollected. tdTomato was excited with a 543-nm helium neon laser andemitted light between 552 and 691 nm was collected. Images were adjustedfor brightness and contrast in ImageJ. Regions of interest (ROIs) weremanually drawn around boutons in ImageJ. ROI pixel intensities wereextracted and processed with a custom Python script (available uponrequest). Pixel intensities were scaled by dividing each value by the meanpixel intensity. These data were binned into 50 equally spaced bins rangingfrom 0 to 8 to generate a frequency histogram. Themedian andmode of pixelintensities were calculated for each image and graphed with KaleidaGraph.

Fixed larval imagingWandering third-instar larvae were reared and dissected as described for liveimaging. Larvae were then fixed with 4% formaldehyde in HL6 buffer for

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5 min, washed with HL6 and mounted in Vectashield (Vector Labs). Slideswere imaged as described for live larvae.

S2 cell imagingpJM952 was transfected into S2R+ cells (Drosophila Genomics ResourceCenter) using Fugene HD (Promega) according to the manufacturer’sprotocols. Cells were adhered to Concanavalin-A-coated glass-bottomdishes (Mat-tek) overnight. Fixed cells were exposed to 4% formaldehyde inHL3 for 5 min and washed with PBS. Cells were imaged on a Nikon A1-Rsilaser scanning confocal microscope using a CFI Plan Apo VC 60× (1.4 NA)oil immersion objective. GFP was excited with a 488-nm argon laser andemitted light between 500 and 550 nm was collected.

ElectrophysiologyWandering third-instar larvae were dissected in HL3 (Stewart et al., 1994)for data collected at 0.4 mM, 0.6 mM, 1.0 mM and 1.5 mM [Ca2+], orHL3.1 (Feng et al., 2004) for data collected at 0.1 mM [Ca2+]. Recordingswere performed as described previously (Howlett et al., 2008). Evokedresponse traces for larvae were recorded for at least 20 s. Quiescent traces of60 s were recorded for mEJP frequency and amplitude analysis. EJP andmEJP analysis was accomplished using Synaptosoft Mini AnalysisProgram. Frequency and amplitude of mEJPs were analyzed using the‘non-stop analysis’ function. For this analysis, the mEJP amplitudethreshold was set at 0.7 mV with a LoPass Butterworth filter and a cut-offfrequency of 200 Hz. EJP amplitude was determined by manually selectingthe peaks, which corresponded to the first 20 stimulations within a trace.Failures within the first 20 stimulations of a trace were denoted as zeroes andcontributed toward average EJP data. No threshold minimum was employedfor EJPs. Corrected quantal content was calculated usingm=ν/ν1(1-ν/V0)

−1,where ν=EJP amplitude, ν1=mEJP amplitude and V0=resting potentialminus a correction factor of 15 mV (Martin, 1955).

pMad quantificationWandering third-instar larvae, reared at room temperature, were dissected inS2 medium (Invitrogen) and fixed immediately (4% formaldehyde in S2medium) for 15 min. Larvae were washed with PBS-T (PBS plus 0.1%Triton X-100) and transferred to Sylgard-coated 24-well plates. Larvae wereincubated with anti-p-Smad1/5 (Ser463/465) rabbit monoclonal antibody(Cell Signaling, 41D10, 1:50 in PBS-T) overnight (4°C) with gentle orbitalshaking, washed with PBS-T, incubated with goat Alexa-Fluor-488-conjugated anti-rabbit IgG antibody (Invitrogen, A-11008, 1:1000 inPBS-T) for 90 min at room temperature, washed with PBS, and mounted invectashield. For each experiment, at least three w1118 larvae were included,and all larvae were exposed to the same antibody dilution. Ventral gangliafrom at least eight larvae from two independent experiments were analyzed.Larvae were imaged as described for live larvae. ROIs were drawn aroundmotoneuronal nuclei in the ventral ganglia using ImageJ. Mean pixelintensity was divided by area. ROIs were summed and averaged for eachlarva, and normalized to the w1118 internal control.

ArborizationWandering third-instar larvae reared at 25°C were dissected in HL3,fixed in 4% formaldehyde (in HL3) and washed with PBS-T. Peltswere placed in sylgard-coated 24-well plates and exposed to goat Alexa-Fluor-488-conjugated anti-horseradish-peroxidase antibody (JacksonImmunoResearch, 123-545-021, 1:500) overnight at 4°C with gentleorbital shaking, washed in PBS, and mounted in Vectashield. Boutons atmuscles 6 and 7 in segment A3 were imaged as described for live larvae.Boutons were counted in ImageJ.

Climb testsCrosses were made with ten mating pairs in half-pint bottles at 25°C.Progeny were collected 2 days after eclosion, and flies were aged for 0 and2 days post eclosion. Males were placed into vials in groups of 10–12 fliesand aged at 25°C. Adult male flies were separated into individual emptyvials, tapped to the bottom and timed to see how long it took them to climb 6cm vertically. Flies experienced three time trials with 5 min of rest between

trials. If a fly failed to cross the 6-cm mark in 60 s, the time was recorded as60 s. The minimum time for each fly was compiled and averaged as the timefor that day. Results represent a minimum of ten flies.

Velocity testsAdult flies that were 0–2 days old were reared at 25°C and anesthetized withcarbon dioxide. Groups of 6–10 males were aged until 3, 10, 15, 25 and30 days (±1 day). Flies were transferred, without anesthesia, into vials with10 ml of 1% agar and allowed to rest for 10 min. Flies were placed in awhite, translucent Plexiglas chamber with a digital video camera focused onthe vial. Two mirrors placed behind the vial at 40° angles produce tworeflections visible in the video. Vials were banged to knock flies to the agarsurface and climbing-behavior videos were captured. Three trials of 30 seach were recorded in succession for each group. Average velocity for eachvial was measured using the iFly system (Jahn et al., 2011; Kohlhoff et al.,2011).

AcknowledgementsThe authors would like to thank various members of the Drosophila community aswell as the Bloomington Stock Center for providing fly lines. Damian Crowther andEleonora Khabirova provided software and assistance with the iFly system. EnriqueMartinez assisted with the S2 imaging, Justin Lee assisted in determining EJPamplitudes with Synaptosoft, Zach Wright assisted with imaging active zoneproteins, Miguel Betancourt-Solis assisted with the sfGFP constructions and JustinVincent assisted with analysis of synaptic bouton number.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsJ.A.M., M.S., J.B.S. and J.E.F. designed the experiments. M.S., J.B.S. and J.E.F.generated reagents, D.P. constructed the UAS-Rtnl1 line while in the lab of A.D.,J.B.S. and J.E.F. performed the experiments. J.B.S., J.E.F., E.F. and J.F. analyzeddata. J.A.M., M.S., J.B.S. and J.F. wrote the manuscript.

FundingE.F. was supported by a George J. Schroepfer, Jr. Undergraduate SummerFellowship; M.S. was supported by a grant from the Hamill Foundation; J.A.M. issupported by the National Institutes of Health [grant number GM101377]; theSimmons Family Foundation; and the Hamill Foundation. Deposited in PMC forrelease after 12 months.

Supplementary informationSupplementary information available online athttp://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.184929/-/DC1

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