Modeling of axonal endoplasmic reticulum network by spastic paraplegia proteins

Axons contain an endoplasmic reticulum (ER) network that is largely smooth and tubular, thought to be continuous with ER throughout the neuron, and distinct in form and function from rough ER; the mechanisms that form this continuous network in axons are not well understood. Mutations affecting proteins of the reticulon or REEP families, which contain intramembrane hairpin domains that can model ER membranes, cause an axon degenerative disease, hereditary spastic paraplegia (HSP). Here, we show that these proteins are required for modeling the axonal ER network in Drosophila. Loss of reticulon or REEP proteins can lead to expansion of ER sheets, and to partial loss of ER from distal motor axons. Ultrastructural analysis reveals an extensive ER network in every axon of peripheral nerves, which is reduced in larvae that lack reticulon and REEP proteins, with defects including larger and fewer tubules, and occasional gaps in the ER network, consistent with loss of membrane curvature. Therefore HSP hairpin-containing proteins are required for shaping and continuity of the axonal ER network, suggesting an important role for ER modeling in axon maintenance and function.


Introduction
Axons allow long-range bidirectional communication in neurons. They carry action potentials along their plasma membrane from dendrites and cell body to presynaptic terminals; and they transport cell components and signaling complexes using motor proteins, anterogradely and retrogradely. Another potential route for communication along axons is endoplasmic reticulum (ER); axons possess a network of ER tubules, which appears physically continuous both locally (Tsukita and Ishikawa, 1976;Villegas et al., 2014) and with ER throughout the neuron (Lindsey and Ellisman, 1985;Terasaki et al., 1994). The physical continuity of ER and its ensuing potential for long-distance communication has been likened to a "neuron within a neuron" (Berridge, 1998).
Axonal ER appears mostly tubular and smooth, with some cisternae (Tsukita and Ishikawa, 1976;Lindsey and Ellisman, 1985;Villegas et al., 2014). Some rough ER is likely to be present too: numerous mRNAs are found in both growing and mature axons (Zivraj et al, 2010;Shigeoka et al., 2016), and local axonal translation can occur in response to injury (Ben-Yaakov et al., 2012;Perry et al., 2012). However, rough ER sheets (Tsukita and Ishikawa, 1976;Villegas et al., 2014), and markers of protein export and folding that are characteristic of rough ER (Röper, 2007;O'Sullivan et al., 2012), are relatively sparse in mature axons. Axonal ER therefore likely has major roles other than protein export; these could include lipid biosynthesis (Tidhar and Futerman, 2013;Vance, 2015), calcium homeostasis and signaling (Ross, 2012), and coordination of organelle physiology (Phillips and Voeltz, 2016). The continuity of the ER network suggests that some of these roles might have long-range as well as local functions; indeed, an ER-dependent propagating calcium wave is seen after axotomy of Caenorhabditis elegans or mammalian dorsal root ganglion neurons (Ghosh-Roy et al., 2010;Cho et al., 2013).
A strong hint of the importance of ER in axons is found in Hereditary Spastic Paraplegia (HSP), a group of axon degeneration disorders characterized by progressive spasticity and weakness of the lower limbs (Blackstone et al., 2011;Blackstone, 2012).

Two widely expressed REEP proteins, ReepA and ReepB, localize to the endoplasmic reticulum
The reticulon and REEP families of double-hairpin-containing proteins are collectively responsible for formation or maintenance of most peripheral ER tubules in yeast (Voeltz et al., 2006). We previously showed that the Drosophila reticulon ortholog Rtnl1 was strongly localized in axons, and that its knockdown caused partial loss of a smooth ER marker in posterior larval segmental axons (O'Sullivan et al., 2012). To test the roles of Drosophila REEP proteins in axonal ER localization, we first dissected the ortholog relationships between the six Drosophila and six human REEP proteins. Multiple sequence alignment of mammalian and Drosophila REEP protein sequences suggested that CG42678 was the single Drosophila ortholog of mammalian REEP1-REEP4 (Fig. 1A). CG42678 has previously been designated Reep1 (http://flybase.org/reports/FBgn0261564.html), but we propose the name ReepA to reflect its orthology to the four mammalian genes REEP1-REEP4. Mammalian REEP5 and REEP6 appeared to share two Drosophila orthologs, CG8331 and CG4960 (Fig. 1). We designated CG8331 as ReepB because of its widespread expression (www.flyatlas.org; Chintapalli et al., 2007) and slower rate of evolutionary sequence divergence (Supplementary Table. 1). We excluded CG4690 and three additional Drosophila REEP genes from further study, since their expression was restricted to testes and larval fat body (www.flyatlas.org; Chintapalli et al., 2007), and their faster rate of evolutionary sequence divergence (Supplementary Table. 1, and reflected in longer branch lengths in Fig. 1A), suggesting poorly conserved function.

Rtnl1and ReepA -ReepBlarval epidermal cells display an abnormal ER network
First, we asked whether Drosophila reticulon and REEP proteins contribute to ER network organization in third instar larval epidermal cells; Rtnl1 knockdown leads to a more diffuse ER network organization and expansion of ER sheets in these cells, compared to wild-type controls (O'Sullivan et al, 2012). Rtnl1mutant larvae also showed loss of ER network organization (Fig 2A). Intensity of KDEL labeling along a line from the nucleus to cell periphery displayed fluctuating intensity, reflecting the reticular distribution of KDEL in wild-type larvae, but less fluctuation in Rtnl1larvae; overall levels of KDEL remained unchanged ( Fig. 2A).
Similarly, loss of both ReepA and ReepB, but not loss of either gene alone, made KDEL levels in larval epidermal cells fluctuate less than in controls. Mean intensity of KDEL staining was decreased in all ReepA and ReepB mutant genotypes, and fluctuation in intensity was increased in ReepBlarvae (Fig. 2B). Ultrastructural analysis revealed that ReepA -ReepBdouble mutant epidermal cells showed longer ribosome studded sheet ER profiles, compared to controls, and to ReepAand ReepBsingle mutants (Fig. 2C). This mutant phenotype is similar to, although less severe than loss of Rtnl1 (O'Sullivan et al., 2012). ReepA -ReepBmutants also showed increased ER stress in epidermal cells ( Fig. 2D) but not in CNS (Fig.   2E). Therefore, Rtnl1 and REEP proteins shape the ER network in Drosophila, and their loss disrupts ER organization, causing longer ER sheet profiles.

Loss of Rtnl1 or ReepB causes partial loss of axonal ER marker from posterior axons
To understand the roles of reticulon and REEPs in axonal ER organization, we labeled axonal ER by expressing Acsl::myc (O'Sullivan et al., 2012) in two adjacent motor neurons using m12- GAL4 (Xiong et al. 2010). Loss of Rtnl1 caused partial loss of Acsl::myc from posterior (segment A6) axons, but not from anterior axons (segment A2); it also caused Acsl::myc staining to appear more irregular in posterior but not anterior axons, reflected in a higher coefficient of variation (SD/mean) of Acsl::myc staining intensity along the length of posterior axons lacking Rtnl1, compared to wild-type (Fig. 3A). These Rtnl1 loss-of-function phenotypes were found either on targeted knockdown of Rtnl1 in these motor axons or in Rtnl1mutant larvae, and the Rtnl1mutant phenotypes could be partially rescued by one copy of an Rtnl1 Pacman genomic clone (Fig. 3A).
ReepAmutants showed no loss of Acsl::myc from axons. Similar to Rtnl1 loss of function, ReepBmutants showed partial loss of Acsl::myc from posterior but not anterior motor axons (Fig. 3B); this phenotype was partially rescued with one copy of a genomic ReepB::GFP clone. A ReepA -ReepBdouble mutant showed loss of Acsl::myc from posterior axons, that was similar to that seen in ReepBmutants (Fig. 3B). ReepA -ReepBdouble mutant larvae, but not ReepAor ReepBsingle mutants, showed an increased coefficient of variation of Acsl::myc staining intensity in posterior but not in anterior axons (Fig. 3B). In summary loss of either Rtnl1 or at least ReepB alters axonal ER distribution in posterior motor axons, therefore supporting roles for these two protein families in axonal ER organization.

Rtnl1and Rtnl1 -ReepA -ReepBmutants disrupt axon ER integrity and axon transport
Since loss of both reticulon and REEP families in yeast removes most peripheral ER tubules (Voeltz et al., 2006), we tested whether loss of both protein families in Drosophila might have similarly severe effects on axonal ER. Rtnl1 -ReepA -ReepBtriple mutant larvae showed increased fluctuation of Acsl::myc staining intensity along motor axons compared to wild-type, mainly in middle parts (segment A4 and A5) of longer axons. At its most extreme, this manifested as fragmentation of Acsl::myc labeling, which was never seen in wild-type axons ( Fig. 4A). When gaps in labeling were found in the central regions of axons, labeling in the anterior and posterior parts of the same axons was usually continuous. Double labeling of plasma membrane (CD4::tdGFP) and axonal ER (Acsl::myc) showed no effect on axonal plasma membrane in Rtnl1 -ReepA -ReepBtriple mutant compared to control axons (Fig. 4B), implying that the phenotype was limited to ER and did not affect axon integrity. To compare genotypes and labels, we quantified irregular labeling along a 45-µm stretch of axon traversing the larval A4/A5 region in two ways: first we measured gaps in labeling, defined as intensity below a threshold that could consistently distinguish labeled axons above background labeling in the nerve; and second, we quantified the coefficient of variation of labeling intensity along the axon (Fig 4B,C). Rtnl1(RNAi) ReepA -ReepBlarvae also showed a fragmentation phenotype similar to Rtnl1 -ReepA -ReepB -, suggesting that loss of Rtnl1 is essential for it ( Fig.   4C). Rtnl1 loss-of-function axons, but not ReepA -ReepBmutant axons, also showed a mild ER fragmentation phenotype (Fig 4A,C). Therefore, loss of Rtnl1 causes a mild irregular organization of axonal ER, and this phenotype is exacerbated by loss of Reep proteins; however, loss of Reep proteins alone has no apparent effect.
We also tested for possible defects in axon transport by staining for abnormal accumulation of the synaptic vesicle protein CSP in axons. Rtnl1larvae and Rtnl1 -ReepA -ReepBtriple mutant larvae showed large accumulations of CSP in many peripheral nerves.
The large accumulations of CSP in Rtnl1larvae could be rescued by two copies of a Rtnl1 Pacman genomic clone (Fig. 5).

Rtnl1 -ReepA -ReepBmutant nerves show ER abnormalities in axons and glia
To better understand wild-type axonal ER organization, and the mutant phenotypes seen in confocal microscopy, we performed electron microscopy (EM) on 60-nm-thick serial sections of third instar peripheral nerves. Peripheral nerves contain both motor and sensory axons, arranged in fascicles, and wrapped in three main classes of glial cell (Stork et al., 2008;Matzat et al., 2015). We used ROTO staining (Tapia et al., 2012;Terasaki et al., 2013) to preferentially highlight cellular membranes including ER.
Wild type larvae showed a network of ER tubules, in every axon that could be observed  (Fig. 6M), and slightly more numerous (but not significantly when averaged across larvae; Fig. 6N) than in wild-type axons, resulting in nearly a four-fold increase in the length of affected axons that lacked ER tubules (Fig. 6O).
Wild-type peripheral nerves are surrounded by an outer perineurial glial cell, and just beneath 9 this a subperineurial glial cell; axons or axon fascicles are wrapped imperfectly by a wrapping glia cell (Stork et al., 2008;Matzat et al., 2015). All glial classes, but particularly subperineurial glia, showed a trend towards increased ER sheet profile length compared to control cells (

Discussion
The existence of a tubular axonal ER network has been known for decades. Nevertheless, the cellular mechanisms that organize a compartment that is usually distributed throughout cells, along the great lengths of axons, are until now largely unknown. The finding that several causative genes for the axon degenerative disease HSP encode ER modeling proteins, suggests a link between ER modeling and axon function or maintenance, and provides candidate proteins that may be instrumental in structure and function of the axon ER network.
These candidates include several hairpin-loop-containing HSP proteins, of the spastin, atlastin, reticulon, REEP, and Arl6IP1 families, that influence ER structure in situations including yeast, mammalian cultured cells, and neuronal cell bodies in vivo (Shibata et al., 2006;Voeltz et al., 2006;Hu et al., 2008;Shibata et al., 2009;Park et al., 2010;Shibata et al., 2010). The HSPrelated protein families that model ER, and some other proteins that interact with them, share a common feature of one or two intramembrane hairpin loops that can insert into the cytosolic face of the ER membrane, thereby recognizing or inducing curvature. This property makes the reticulon and REEP(DP1) families together responsible for most peripheral ER tubules in yeast, and contribute to the curved edges of ER sheets (Voeltz et al., 2006;Hu et al., 2008). ortholog of human REEP5-REEP6. We monitored phenotypes of these mutants, singly and in combination, by visualizing ER in individual axons in situ: confocal microscopy of axonal ER markers in small numbers of motor neurons, and EM using membrane-specific staining. 10 Confocal microscopy revealed a partial loss of ER marker in distal but not in more proximal motor axons in Rtnl1 and in ReepB mutants. Although the only REEP genes identified as causative for HSP are REEP1 and REEP2, loss of their ortholog ReepA had no effect on axonal ER on its own. However, some role for ReepA in modeling axonal ER is suggested by a ReepA -ReepBdouble mutant having a more severe axonal ER phenotype than a ReepBmutant (Fig. 4B). The less severe phenotype of ReepAcompared to ReepBmutants is consistent with the lower levels of ReepA expression, judged by transcriptomics (www.flyatlas.com), and the relative expression of their GFP fusions (Fig. 2).
Given the joint and partly redundant requirement of the reticulon and REEP families for ER tubule formation in yeast (Voeltz et al., 2006), we tested whether this was also true for axonal ER. Flies lacking Rtnl1, ReepA and ReepB -equivalent to mammals lacking all four reticulons and all six REEPs, and homozygous viable -indeed showed more extreme ER fragmentation ( Fig. 4)  EM examination of wild-type axonal ultrastructure revealed that ER tubules were effectively ubiquitous in Drosophila peripheral nerve sections, as seen previously in mammalian neurons (Tsukita and Ishikawa, 1976;Villegas et al., 2014), albeit with fewer tubules, presumably reflecting the smaller diameters of the axons examined here.
Reconstruction over several µm showed a continuous network of ER tubules in most axons examined, in agreement with the ER continuity found in neurons by lipid dye labeling (Terasaki et al., 1994). However, in a few axons, we found short lengths of axon with no detectable ER ( Fig. 6J-O). There could be several reasons for this: Terasaki et al. (1994) only assessed continuity in dendrites, cell body and proximal axon; some of the gaps we observe in EM could be short transient gaps in a dynamic network; larval axons with low diameters might be 11 intrinsically more susceptible to occasional gaps in the ER network, than wider axons with more tubules; and we might occasionally miss an ER tubule due to weaker staining, or close proximity to other structures like plasma membrane. We also observed occasional small ER sheet-like structures in wild-type axons ( Fig. 6H; Supplementary Movie 3). Although we do not see ribosomes on these, this could be due to lack of staining by the ROTO protocol. As discussed above, rough ER and translation are relatively sparse in axons, but low levels of rough ER are possible, and consistent with the occasional sheet structures observed here.
EM also showed phenotypes consistent with loss of ER membrane curvature in Rtnl1 -ReepA -ReepBtriple mutant axons (Fig. 6). Mutant axons had ER tubules of larger diameter, fewer tubules per axon cross-section, and consistent with our confocal data (Fig. 4), longer gaps in the ER network than wild-type, although most parts of most mutant axons examined still had a continuous ER network (Fig. 8). These mutant phenotypes could potentially have physiological consequences. Larger tubules could potentially store and release more calcium than thinner ones, while the reduced network could make the role of the ER in calcium buffering or release more localized. The less extensive ER network in mutants might also reduce the amount of contact between ER and other organelles, with consequences for calcium and lipid homeostasis that require these contacts, or for regulation of mitochondrial fission (Friedman et al., 2011) -although the continuing proximity of ER to mitochondria and plasma membrane in mutants means that any effects are presumably quantitative rather than qualitative. The reduced curvature of ER membrane in mutants might also influence their protein composition, since many membrane proteins have mechanisms for recognizing differential membrane curvature (Antonny, 2011). The occasional lack of continuity could prevent propagation of ER-dependent Ca 2+ signals like those seen in injured mammalian sensory neurons (Cho et al., 2013); it could also cause local impairments in Ca 2+ or lipid homeostasis that could lead to local transport inhibition, as is the case for mitochondrial transport (Wang and Schwarz, 2009). Sporadic lack of ER continuity might explain the preferential sensitivity of distal longer axons to HSPs, since these would be more likely to suffer from a gap in ER continuity to the cell body, compared to proximal or shorter axons. In this model, disease-causing alleles in single hairpin-encoding genes could promote degeneration in distal motor axons by increasing the probability of such gaps, dependent on factors such as age, axon length or diameter, and ER tubule density and dynamics.
The apparent ubiquity of ER in axons, the extent of its continuity over long distances, and the preferential susceptibility of distal longer axons to mutations that affect ER-modeling proteins, all point to important physiological roles of this compartment and of its continuity. In this work we have begun to reveal the mechanisms that determine its organization. We have shown roles for two protein families that contain HSP disease gene products, in influencing the shape of individual tubules and the axonal ER network, with potential physiological 12 consequences that would also be affected by mutations in these genes. Understanding these physiological consequences, both in the genotypes we have described here, and in new genotypes that might also affect axonal ER organization and continuity, will provide models for the potential physiological defects in HSP and other axon degeneration diseases.

Materials and Methods
Drosophila genetics. ReepA 541 (referred to as ReepA -), and ReepB 48 (referred as ReepB -) mutants were generated by imprecise excision of P elements shown in Fig. 1. One of the precise excisions generated in these experiments, ReepA +C591 (referred to as ReepA + ) was used as a genetic background control where feasible. Rtnl1 1 , referred as Rtnl1 -, was a gift from G. Tear (Wakefield and Tear, 2006). ReepA -ReepBand Rtnl1 -ReepA -ReepBwere generated by meiotic recombination on the second chromosome, and recombinants were screened for using PCR primers (Supplementary Table 2 Supplementary Table 2. BLAST sequence searches were used to define genome coordinates of P-element excisions, and Pfam domain coordinates in coding regions, and compare protein divergence rates. They were performed at the National Center for Biotechnology Information (www.ncbi.nlm.nih.gov). REEP dendrograms were drawn from a ClustalW alignment (Larkin et al., 2007) using the neighbor-joining algorithm in MEGA 5.05 (Tamura et al., 2011).
Analysis of ER structure. Confocal images were analyzed blind to genotype using ImageJ (imagej.nih.gov/ij/). Images of entire epidermal cells were obtained as z-projections of three consecutive sections. Using the line tool of ImageJ a 12-µm line was drawn from the nuclear envelope towards the periphery of each cell analyzed. Pixel intensity along the line was recorded in an Excel file. Local variance of intensity was calculated by dividing the rolling variance of the intensity (in 10-pixel windows), by the rolling mean intensity, all along the line.
Proximal (anterior) axons were imaged from segment A2, middle images were from the end of segment A4 and A5, distal (posterior) axons were imaged from segment A6 of third instar larvae. Mean gray intensity for single axon images were measured by drawing a 45-µm line, either along both M12-GAL4-expressing axons (where they could not be separated), or along the most strongly labeled axon (where they appeared as separate axons), and quantifying gray intensity (0-255) by ImageJ; occasional images with saturated pixels were excluded from analysis after blinding. Coefficient of variation was calculated by dividing the standard deviation of staining intensity by the mean; occasional images with faint staining throughout the axon were excluded from analysis after blinding. Gaps were defined as regions where staining intensity was less than 20 (out of 255), after background subtraction. 14 Electron microscopy. For epidermal cell EM, larvae were prepared and fixed as described by O'Sullivan et al. (2012). Transverse sections were cut on a Leica Ultracut UCT ultramicrotome at 70 nm, using a diamond knife, and contrasted with uranyl acetate and lead citrate (for epidermal cells). Sections were viewed using a Tecnai G2 electron microscope operated at 120 kV, and an AMT XR60B camera running the Deben software in the Multi-Imaging Centre, School of Biology, University of Cambridge.
For EM of peripheral nerves, we used a ROTO protocol (Tapia et a., 2012, Terasaki et al., 2013 to highlight membranes. Third instar larvae were dissected in HL3 solution and fixed in 0.05 M sodium cacodylate (pH 7.4) containing 4% formaldehyde, 2% vacuum distilled glutaraldehyde, and 0.2% CaCl 2 ), at 4 o C for 6 hours. Larvae were dissected as for confocal analysis, but leaving overlying organs such as gut and fat body attached, to reduce loss of peripheral nerves during processing. Preparations were then washed 3 times for 10 minutes each at 4 o C using cold cacodylate buffer with 2 mM CaCl 2 . A solution of 3% potassium ferricyanide in 0.3 M cacodylate buffer with 4 mM CaCl 2 was mixed with an equal volume of 4% aqueous osmium tetroxide; larval preparations were incubated in this solution at 4 o C for 1-12 hours, then rinsed with deionized water at room temperature 5 times for 3 minutes each.
Thiocarbohydrazide solution was prepared by adding 0.1 g thiocarbohydrazide to 10 ml deionized water, kept in a 60 o C oven in a secondary embedding pot for 1 hour, swirled every 10 minutes to facilitate dissolution, and filtered through two 9 cm filter papers just before use.
Larval preparations were incubated in thiocarbohydrazide solution for 20-30 minutes at room temperature and covered with foil to protect from light. Then they were rinsed with deionized water at room temperature 5 times for 3 minutes each, incubated in 2% osmium tetroxide for 30-60 minutes at room temperature, and rinsed with deionized water at room temperature 5 times for 3 minutes each. Preparations were incubated in 1% uranyl acetate (maleate-buffered to pH 5.5) at 4 o C overnight and rinsed with deionized water at room temperature 5 times for 3 minutes each. Then they were incubated in lead aspartate solution (0.66 g lead nitrate dissolved in 100 ml 0.03 M aspartic acid, pH adjusted to 5.5 with 1 M KOH) at 60 o C for 30 minutes and rinsed with deionized water at room temperature 5 times for 3 minutes. Then they were dehydrated twice with 50%, 70%, 90% and 100% ethanol, twice with dried ethanol, twice with dried acetone and twice with dry acetonitrile. Preparations were incubated in 50/50 acetonitrile/Quetol 651 overnight at room temperature, three times for 24 hours each in Quetol epoxy resin 651 (Agar Scientific, Stansted, UK) and three times for 24 hours each in Quetolepoxy resin 651 with BDMA (dimethylbenzylamine). They were then incubated at 60 o C for at least 48 hours.
Serial 60-nm-thick transverse sections were cut in the larval abdominal region, visualized using scanning EM, and images were aligned for analysis of serial sections and reconstruction, as described by Terasaki et al. (2013).

Axonal EM analyses.
To quantify axonal ER tubule diameter, non-axonal staining was removed manually, and ER tubule profiles were identified based on the local threshold in a single cross-section, and the presence of signals at the same position in adjacent sections.
The minimum Feret diameter of each tubule was measured using ImageJ Fiji (https://fiji.sc) via the Analyze Particles command. ER numbers per axon were counted manually for all axons detected in the nerve. 3D reconstruction was carried out using the Fiji TrakEM2 plug-in. To quantify gaps in the tubule network, each axon was analyzed throughout the entire stack of sections. To allow for occasional lightly stained or blurred sections, only complete loss of ER tubules from three or more sections in an axon was defined as a gap. Continuous ER tubules were identified as the presence of signals at the same position for three or more sections.
Given the varying brightness and contrast of EM sections, faint staining that coincided with a tubule signal in adjacent sections was also considered as an ER tubule. Color shading and sketches drawn in Fig. 7 were processed in Adobe Photoshop CS6. For quantification of glial ER sheet length, individual ER sheet profiles were measured using Fiji via the line tool and Measure command. Wrapped axons were defined as one-axon or two-axon fascicles which were completely wrapped by glial cells and isolated from other neighboring axons.
Statistical analysis. Statistical analyses were performed in GraphPad Prism 6. Data were analyzed by either two-tailed Student's t-test or Mann-Whitney U tests (for data that were not normally distributed). Bar graphs and scatter plots show mean ± SEM; box plots show median with interquartile range, and the 5% and 95% percentiles as whiskers. Sample sizes are reported in figures. Confocal mean intensity datapoints from separate cells or axons were pooled across larvae or across experiments when ANOVA analysis indicated no differences among samples of the same genotype; when ANOVA analysis indicated differences among larvae or repeated experiments, mean intensity was averaged to yield datapoints for each larva or experiment, respectively. P levels are indicated as ns P > 0.05, *P < 0.05, **P < 0.01, ***P < 0.001, or ****P < 0.0001, except where indicated, when criteria could be defined more stringently.

Fig. 5. Loss of Rtnl1 causes mild accumulation of synaptic vesicles in axons
Peripheral nerves of Rtnl1and Rtnl1 -ReepA -ReepBtriple mutant larvae show larger accumulations of synaptic vesicle protein CSP (e.g. yellow arrows), and smaller elongated CSP puncta (e.g. yellow arrowheads). In contrast, control larvae and ReepA -ReepBdouble mutant larvae show an even distribution of small round CSP puncta. CSP accumulations are not significantly bigger in Rtnl1 -ReepA -ReepBtriple mutants than in Rtnl1mutants. The CSP accumulations in Rtnl1larvae can be rescued by two copies of a Rtnl1 Pacman genomic clone.