Introduction

The nervous system is highly plastic, with the capacity to undergo dynamic alterations in structure and strength in response to changing stimuli and environments (Lamprecht and LeDoux, 2004; Citri and Malenka, 2008). This activity-induced synaptic remodeling process is highly conserved across species and plays crucial roles in circuit formation during development, as well as in stabilizing existing connections post-development. Synaptic remodeling represents a finely orchestrated process, with communications across both the pre- and post-synapses to allow the growth of new synapses, and the stabilization and strengthening of existing ones. While much of the research on synaptic plasticity has concentrated on interactions between presynaptic axon terminals and postsynaptic cells, most synapses are tripartite synapses, with glial cells as the third cell type (Araque and Navarrete, 2010). Beyond simply providing metabolic support, glial cells are increasingly recognized for their roles as active modulators of synaptic plasticity (Sancho et al., 2021). However, how glial cells and neurons collaborate to coordinate activity-induced synaptic remodeling are not well understood.

The Drosophila larval neuromuscular junction (NMJ) is a genetically tractable system and a tripartite glutamatergic synapse that serves as an excellent model system to investigate mechanisms underlying activity-induced synaptic remodeling by glial cells (Banerjee and Bhat, 2008; Freeman, 2015; Kim et al., 2020). The peripheral glial cells at the fly NMJ perform some of the key functions similar to mammalian glia, including controlling neuronal excitability and conduction velocity (Kottmeier et al., 2020; Rey et al., 2023), recycling of neurotransmitters (Rival et al., 2004; Danjo et al., 2011), and engulfing and clearing debris during damage to allow the growth of new boutons (Fuentes-Medel et al., 2009). They also release proteins such as transforming growth factor (TGF-β) to support synaptic growth (Fuentes-Medel et al., 2012), TNF-α (eiger) to influence neuronal survival (Keller et al., 2011), Wingless to regulate GluR clustering (Kerr et al., 2014), and laminin to control animal locomotion (Petley-Ragan et al., 2016). However, the role of peripheral glia in regulating activity-induced stabilization and remodeling of the existing synapses at the NMJ remains unknown.

Previous studies on activity-induced synaptic remodeling at the fly NMJ demonstrated that neuronal activity leads to the enlargement of existing boutons, accompanied by increases in postsynaptic GluR abundance (Lee et al., 2017; Chang et al., 2024). Intense neuronal stimulation triggers the release of a protein called Shriveled (Shv) by presynaptic motoneurons, which activates βPS integrin bi-directionally to stimulate synaptic bouton enlargement and elevate GluR levels on the postsynaptic muscles (Lee et al., 2017). Consequently, shv mutants display defective post-tetanic potentiation (PTP), a form of functional synaptic plasticity similar to the early phase of long-term potentiation (LTP) seen in mammalian neurons. Here we demonstrate that the Shv protein is also expressed in glial cells and is released extracellularly by peripheral glial cells. Glial Shv not only regulates basal GluR clustering but is also required for activity-induced synaptic remodeling. We further demonstrate that while glial Shv is present extracellularly, it does not respond to neuronal activity nor does it activate integrin signaling, unlike Shv derived from neurons. Instead, glial Shv contributes to synaptic plasticity regulation by modulating the levels of Shv release from neurons and by controlling the levels of ambient glutamate concentration. Restoring ambient glutamate concentration could correct basal GluR abundance and defective synaptic plasticity caused by the loss of glial cells. These results further reveal that regulation of ambient extracellular glutamate concentration by glia is an important mechanism contributing to synaptic plasticity regulation.

Results

Shv is expressed in peripheral glia

To determine the role of Shv in glia, we first monitored its presence in different cell types. To this end, we knocked-in eGFP to the 3’-end of the full-length Shv protein using CRISPR/Cas9-catalyzed homology directed repair (Gratz et al., 2014). Western blots confirmed that the Shv protein is tagged with eGFP (Fig 1A), and immunostaining revealed its presence in neurons and glial cells (Fig 1B). Glial cells were identified using antibody against reverse polarity (Repo), a transcription factor expressed exclusively in glial cells (Xiong et al., 1994), and neuronal cells were marked either by Elav, a transcription factor expressed in neurons (Robinow and White, 1991), or HRP, which stains the neuronal cell membrane. We found that Shv-eGFP is present in both Repo and Elav positive cells in the larval brain, consistent with our previous report (Lee et al., 2017). Furthermore, Shv-eGFP can be detected at the NMJ, with weak signals in postsynaptic muscles and synaptic boutons, and stronger signals in peripheral glia (Fig 1C).

Figure 1.

Glial Shv is required for activity-induced synaptic remodeling

Loss of Shv in shv¹ mutant was previously shown to be essential for activity-induced synaptic remodeling (Lee et al., 2017). Given that Shv is observed in neurons, glia, and muscle (Fig 1C), we determined the tissue-specific requirement for Shv by systematically knocking down shv using RNAi and tissue-specific drivers (Fig 2A). We found that shv knockdown in neurons using the pan-neuronal driver, elav-GAL4, generated the same phenotype as shv¹ mutant (Lee et al., 2017), namely smaller bouton size, reduced basal GluR intensity, and defective synaptic remodeling in response to neuronal stimulation. However, knockdown of shv in glia using the pan-glial specific driver, repo-GAL4, resulted in normal bouton size but significantly elevated GluR levels, as well as abolished activity-induced synaptic remodeling. This increase in GluR is unexpected and further suggests that neuronal and glial Shv acts through different pathways to regulate GluR abundance. Lastly, knockdown of shv in post-synaptic muscles using the muscle specific driver 24B-GAL4 did not affect synaptic development or activity-induced synaptic changes. Collectively, these results are consistent with an earlier finding that Shv is predominantly released by neurons to maintain activity-induced synaptic remodeling (Lee et al., 2017), as well as reveal an unexpected requirement for glial Shv in sustaining activity-induced synaptic changes and basal GluR abundance.

Figure 2.

Given that reducing Shv in glia altered basal GluR levels during development, we used the GeneSwitch system to determine the temporal requirement for Shv in glia (Fig 2B). In the presence of RU486, the repo-GeneSwitch driver can undergo a conformational change to activate gene expression in glia (Osterwalder et al., 2001; Roman et al., 2001; Artiushin et al., 2018). When the 3^rd instar larvae were fed with RU486 to transiently drive shv-RNAi expression in glia, we observed defective synaptic remodeling and elevated GluR levels. These results suggest an acute requirement for the glial Shv protein in regulating synaptic plasticity and GluR abundance.

Next, we determined the spatial requirement for Shv in glial cells. The peripheral glia can be divided into three subtypes: wrapping glia (WG) is the innermost layer, wrapping and contacting the peripheral nerve bundle; the subperineurial glial (SPG) covers the WG, establishing the blood-brain barrier; the perineurial glia (PGs) is located on the outermost surface of the nerve that is also part of the blood-brain barrier (Awasaki et al., 2008; Stork et al., 2008; Brink et al., 2012; Freeman, 2015; Fernandes et al., 2024) (Fig 3A). Using GAL4 lines previously shown to drive expression in the specific glial subtypes (Stork et al., 2012), we found that reducing Shv in either SPG or PG was sufficient to block activity-induced synaptic remodeling (Fig 3B), phenocopying shv knockdown in all glial cells (Fig 2A). NMJs with shv knockdown in wrapping glia (nrv-Gal4) exhibited normal activity-induced synaptic remodeling (Fig 3B). We also tested whether the astrocyte-like glia located in the central nervous system also contributes by knocking down Shv using alrm-GAL4 (Doherty et al., 2009). We observed normal activity-induced synaptic remodeling (Fig 3). Together, these results suggest that Shv in PG and SPG glia, both in contact with synaptic boutons, extending into the muscles, and are part of the blood-brain barrier, are crucial for Shv function in glia and for synaptic remodeling.

Figure 3.

Glial expression of Shv is sufficient to rescue synaptic plasticity in shv mutants

While it has been demonstrated that Shv derived from neurons is sufficient to rescue shv¹ mutant in activity-induced synaptic remodeling (Lee et al., 2017), it is not known whether glial Shv can compensate for the loss of neuronal Shv. To test this, we expressed Shv (UAS-Shv) in shv¹ mutant using the repo-GAL4 driver. Figure 4A shows that glial expression of Shv was sufficient to restore basal GluR intensity and activity-induced synaptic remodeling but did not normalize bouton size. These data demonstrate that while neuronal Shv is necessary to regulate bouton size during development, Shv derived from glia is sufficient to maintain basal GluR levels and support activity-induced synaptic modifications.

Figure 4.

In addition to structural rescue, we next tested whether glial Shv can rescue functional plasticity. Electrophysiological recordings showed shv¹mutants displayed reduced miniature excitatory potential (mEPSP) amplitude but normal evoked EPSP (Fig 4B), consistent with a previous report (Lee et al., 2017). Selective expression of shv in glia was sufficient to restore mEPSP amplitude, in line with the rescue in GluR levels. Glial-specific expression of shv also rescued post-tetanic potentiation (PTP), an activity-dependent plasticity in Drosophila that is functionally similar to the initial stages of long-term potentiation (LTP) (Fig 4C). Collectively, these results suggest that glial Shv is sufficient to support functional and structural plasticity.

Glial Shv does not activate integrin signaling

Shv was previously shown to be released by neurons to trigger synaptic remodeling through integrin activation (Lee et al., 2017); we thus asked whether glial Shv restores synaptic plasticity in shv¹ mutants by activating integrin signaling similar to its neuronal counterpart. Figure 5A shows that glia can indeed release Shv, as glial expression of HA tagged Shv (driven by repo-GAL4) can be detected extracellularly when stained using non-permeabilizing conditions. Next, we determined the effects of glial Shv on integrin signaling by monitoring the levels of phosphorylated focal adhesion kinase (pFAK), as its levels strongly correlate with integrin activation (Mitra et al., 2005; Tsai et al., 2008). To our surprise, glial expression of shv in shv¹ mutant did not restore pFAK levels to normal (Fig 5B). Furthermore, shv knockdown in glial cells exhibited higher pFAK staining compared to the control (Fig 5C), a result that is opposite to what was observed with shv knockdown in neurons (Lee et al., 2017). shv overexpression in glia, although did not change basal pFAK levels, blocked the activity-induced pFAK increases normally seen in control (Fig 5C). Collectively, these results suggest that glial Shv does not activate integrin and appears to play an inhibitory role in activity-induced integrin activation.

Figure 5.

To elucidate whether glial Shv directly inhibits activity-induced integrin activation, we monitored the amount of Shv released by glia during neuronal activity by expressing Shv-HA in either glia or neurons of shv¹and measured its extracellular levels. Fig 5D demonstrates that while stimulation significantly increased extracellular Shv released by neurons, the amount of extracellular Shv released by glial cells was reduced. These data indicate that glial Shv does not directly inhibit integrin signaling. Next, we tested the hypothesis that glial Shv instead influences integrin signaling by regulating Shv release from neurons. To this end, we monitored Shv released by neurons or glia using the endogenously tagged Shv-eGFP line and tissue-specific knockdown of shv. When Shv was selectively knocked down in glia, an increase in extracellular Shv-eGFP level was observed at the NMJ (Fig 5E), which likely originated from neurons. This finding is consistent with a role for glial Shv in suppressing neuronal Shv release and further explains the higher pFAK level observed in case of glial shv knockdown (Fig 5C). A similar compensatory upregulation from glia was obtained when Shv was knocked down in neurons (Fig 5E). We also monitored the levels of Shv-eGFP in the larval brain, confirming that the shv-RNAi approach successfully reduced Shv levels in the selective cell types (Sup Fig 1). Taken together, these results reveal that the amount of Shv released by neurons is homeostatically regulated by Shv produced in glia. Furthermore, unlike neurons, glial Shv release is independent of neuronal activity and does not directly alter integrin signaling.

Drosophila peripheral glia and Shv control ambient extracellular glutamate levels to regulate activity-induced synaptic remodeling

If Shv derived from glia does not activate integrin signaling, how does it restore synaptic plasticity in shv¹? One plausible explanation is that glial Shv is required for normal glial growth or survival. We therefore monitored glial morphology when Shv is knocked down in glia by co-expressing membrane targeted CD4-tdGFP using the repo-GAL4 driver. Sup Fig 2 shows that there is no observable change in gross glial morphology, with glial processes extending and associating with proximal synaptic boutons, indicating glial Shv is not required for glial maintenance. This finding further bolster support for the claim that glial Shv does not activate integrin, which is essential for maintaining the integrity of peripheral glial layers (Xie and Auld, 2011).

An alternate possibility is that Shv modulates glial function to rescue activity-induced synaptic remodeling. Glial cells have an established role in providing support and maintaining glutamate homeostasis in the nervous system (Augustin et al., 2007). The Drosophila larval NMJ is a glutamatergic synapse with high ambient glutamate concentration in the hemolymph, with an average in the range of 1-2 mM glutamate (Chen et al., 2009). Surprisingly, the larval NMJ does not contain the excitatory amino acid transporter 1 protein (Eaat1), which is essential for removing extracellular glutamate, suggesting that high extracellular glutamate is better tolerated and removed by diffusion through the hemolymph (Rival et al., 2006; Chen et al., 2009). The high ambient glutamate concentration is maintained by the cystine/glutamate antiporter (Cx-T), which imports cystine and exports glutamate into the extracellular milieu (Augustin et al., 2007; Grosjean et al., 2008). The purpose of the high extracellular glutamate is not well understood, but it is thought to control GluR clustering and maintain a reserved pool of GluR intracellularly (Chen et al., 2009). Given that we observed significantly upregulated GluR clustering when Shv was reduced in glia, we hypothesized that glial Shv may influence the levels of ambient extracellular glutamate. To detect glutamate level at the synapse, we took advantage of the GAL4/UAS and LexA/LexAop systems to express the glutamate sensor (iGluSnFR) in neurons while simultaneously knocking down or upregulating Shv in glia (Marvin et al., 2013) (Fig 6A). Knockdown of Shv in glia significantly reduced iGluSnFR signal, whereas upregulating Shv increased it, suggesting that glial Shv critically controls ambient extracellular glutamate levels. As a control, we also expressed Shv in neurons using Elav-LexA (Fig 6A). We found neuronal expression of Shv did not alter iGluSnFR signal at the synapse, confirming that the change in ambient glutamate level is selectively caused by Shv from glia.

Figure 6.

Next, we set out to determine whether the reduced extracellular ambient glutamate is responsible for the defective synaptic remodeling observed when Shv is depleted in glia. To this end, we incubated the NMJ with HL-3 solution containing 2 mM glutamate for 1 hour before stimulating the NMJ with high KCl. Strikingly, this treatment condition not only corrected basal GluR levels, but also fully rescued the activity-induced synaptic remodeling defect seen in glial knockdown of shv (Fig 6B). In contrast, incubating the NMJ with 2 mM glutamate did not correct basal GluR or activity-induced synaptic remodeling defect when Shv is knocked down in neurons (sup Fig 3). These data are consistent with the iGluSnFR imaging results and further confirm that glial and neuronal Shv acts through distinct pathways to jointly regulate synaptic remodeling.

To further understand the function of glial cells in synaptic plasticity regulation, we ablated glia by using the inducible repo-GeneSwitch driver to express reaper (rpr) in early third instar larvae. This method allows us to examine the acute requirement for glia while avoiding lethality associated with chronic glial ablation. Following RU486 treatment for 24 hours, fragmentation of the glial cell membrane and a reduction in GFP intensity was observed (Fig 7A), indicating degeneration of glial cells. This is consistent with an earlier report demonstrating that Rpr expression is sufficient to induce apoptosis and trigger cell death (White et al., 1996). Fig 7B shows that glial ablation caused the same phenotype as shv knockdown in glial cells (Fig 2A), with elevated basal GluR levels and blocked activity-induced synaptic remodeling. We hypothesized that if a primary role of glial cells in synaptic plasticity regulation is to maintain ambient extracellular glutamate levels, similar to glial Shv, then correcting extracellular glutamate concentration should be sufficient to restore synaptic plasticity, even in the absence of functional glia. Strikingly, incubating NMJs with 2 mM glutamate not only restored basal GluR levels but also rescued activity-induced synaptic remodeling caused by glial ablation (Fig 7B, 7C). Together, these data confirm that the ability of peripheral glial cells to maintain high ambient extracellular glutamate concentrations at the NMJ is crucial for synaptic plasticity.

Figure 7.

Discussion

In this study, we show that the peripheral glial cells at the Drosophila NMJ play an important role in regulating synaptic plasticity. We demonstrate that neurons and glia jointly orchestrate activity-induced synaptic remodeling at the NMJ, with Shv playing a pivotal role. While neurons release Shv in an activity-dependent manner to regulate synaptic remodeling through integrin signaling (Lee et al., 2017), release of Shv by peripheral glial cells does not rely on neuronal activity and does not activate integrin signaling (Fig 5A-C). Instead, glial Shv influences activity-induced synaptic remodeling by keeping the levels of Shv released by neurons in check (Fig 5D-F) and controlling ambient extracellular glutamate levels (Fig 7).

Our data suggest that one mechanism underlying activity-induced synaptic remodeling by glia is through indirect control of Shv release from neurons, thereby maintaining minimal integrin signaling under ambient conditions. We propose that this low baseline integrin signaling enables neurons to respond with high sensitivity to enhanced integrin activation by Shv release from neurons following neuronal activity, leading to rapid synaptic remodeling. Conversely, knocking down Shv in glia removes this suppression, resulting in increased Shv release from neurons, higher basal integrin signaling, GluR levels, and pathway saturation, thereby inhibiting activity-induced synaptic remodeling. Supporting this, overexpression of Shv in neurons elevated basal pFAK and abolished synaptic plasticity (Lee et al., 2017). These findings also raise several intriguing questions for future research, including how neurons and glia distinguish different sources of Shv, and how they sense and communicate this information to regulate extracellular Shv levels from different cells.

Another main function of glial Shv is to regulate ambient extracellular glutamate concentration. We report that overexpression of shv elevated ambient glutamate levels at the synapse measured using a genetically encoded glutamate sensor, whereas knockdown of shv in glia reduced its level (Fig 6B). Furthermore, ablating glia recapitulated the phenotypes of shv knockdown in glia, and transiently restoring extracellular ambient glutamate concentration efficiently rescued synaptic plasticity. It has been shown that the Drosophila larval NMJ maintains a surprisingly high ambient extracellular glutamate concentration, with an average in the range of 1-2 mM (Augustin et al., 2007). How Shv regulates extracellular glutamate concentration remains to be explored, but a likely mechanism is by affecting the levels or functions of the cystine/glutamate (Cx-T), which imports cystine and exports glutamate into the extracellular matrix. While Shv has been shown to activate integrin, it encodes a homolog of the mammalian DNAJB11protein (Lee et al., 2016), which functions as a molecular chaperone vital for proper protein folding in the endoplasmic reticulum (Shen et al., 2002). Shv thus could potentially be required for the proper folding and the function of the Cx-T, which is located on glial cell membrane at the fly NMJ (Augustin et al., 2007; Grosjean et al., 2008). Aligned with this, mutations in Cx-T also resulted in elevated basal GluR levels at the fly NMJ (Augustin et al., 2007). Future studies examining the functional interactions between Cx-T and Shv will shed light on mechanisms for Shv-dependent regulation of ambient extracellular glutamate levels at the NMJ.

How does extracellular glutamate regulate GluR levels and synaptic plasticity? Changes in glutamate levels has been shown to directly impact neurotransmission, glutamate receptor activity, and influence GluR clustering by internalizing the desensitized GluR (Featherstone and Shippy, 2008; Chen et al., 2009; Yao et al., 2018). We have also shown that basal GluR level is homeostatically regulated by extracellular glutamate concentration (Fig 7). It is plausible that a high extracellular glutamate concentration enables the NMJ to maintain a reserved pool of GluRs that can be readily mobilized to the surface upon neuronal activity. Additionally, activation of GluR and downstream signaling pathways could trigger local protein translation machineries to prime the synapses to respond rapidly to changes in neuronal activity. Future studies examining intracellular mechanisms controlling activity-induced GluR increases will lead to better insights on synaptic plasticity regulation.

Materials and methods

Fly stocks

Flies were maintained at 25°C on a standard fly food consisting of cornmeal, yeast, sugar, and agar, under a 12-hour light/dark cycle unless otherwise specified. The following fly lines were obtained from the Bloomington Drosophila Stock Center at Indiana University (BDSC, stock numbers in parentheses), the Vienna Drosophila Resource Center and Drosophila Genomics Resource Center (VDRC, specified before stock number), or Kyoto stock center (Kyoto, stock number in parentheses): repo-GAL4 (BDSC, 7415), 24B-GAL4 (BDSC, 1767), elav-GAL4 (BDSC, 458), nSyb-GAL4 (BDSC, 51635), Gliotactin-GAL4 (BDSC, 9030), Nrv-GAL4 (BDSC, 6800), alrm-GAL4 (BDSC, 67032), TRiP-RNAi control (BDSC, 35788), iGluSnFR (BDSC, 59611), UAS-CD4-tdGFP (BDSC, 35836), UAS-IVS-myr::tdTomato (BDSC, 32221), Elav-LexA (BDSC, 52676), UAS-shv-RNAi (VDRC 108576), PG-GAL4 (also known as NP6293-GAL4, Kyoto, 105188), and, repo-GeneSwitch-GAL4 (Artiushin et al., 2018), repo-LexA (a gift from Dr. Henry Y. Sun), UAS-Shv and shv¹ (Lee et al., 2017).

To visualize the endogenous levels of Shv, Shv with eGFP inserted into the 3’-end of the endogenous Shv gene (Shv-eGFP) was generate using the CRISPR/Cas9 genome editing, and homology directed repair (HDR) as described (Gratz et al., 2014). Briefly, a target Cas9 cleavage site in Shv was selected at the 3’ end of Shv without obvious off-target sequence in the Drosophila genome using CRISPR optimum Target Finger. The sgRNA target sequence: 5’-GCCGGCGAGCACTTTTATTG was cloned into the pU6-BbsI-chiRNA vector (Addgene, #45946). The vector for HDR contains the 5’ homology arm containing Shv genomic region (1000 base pairs at the 3’ end of the Shv gene with the stop codon removed) plus eGFP sequence, and the 3’ homology arm (1020 base pair downstream of the Cas9 cleavage site in the Shv gene region) was cloned into pHD-DsRed (Addgene, # 51434). Both cloned vectors were injected into fly stocks containing Cas9 (BDSC 55821) and positive genome editing screen by the presence of DsRed by BestGene Inc. Insertion was confirmed by sequencing and Western blots.

LexAop-Shv RNAi and LexAop-Shv fly lines were generated by subcloning Shv-RNAi hairpin sequence based on VDRC Shv-RNAi design or the coding regions of Shv which contains a V5 tag into the pJFRC19-13XLexAop2-IVS-myr::GFP vector (Addgene, 26224). Both lines were inserted into the X chromosome specific position by microinjecting the plasmid into a P{CaryP}attP18 fly line (BDSC, 32107).

RU486 feeding

repo-GeneSwitch embryos were collected in 3 hours windows and early 3^rd instar larvae (48 hours after larval hatching), were fed with food containing 10 μM RU486 (1:1000, 10 mM stock) or 0.1% EtOH (vehicle control) for 24 hours before dissection (Gatto and Broadie, 2008).

Immunochemistry

Dissection and stimulation of the third-instar larvae fillets were done as described (Lee et al., 2017). Briefly, dissected NMJs were fixed for 3 minutes at room temperature (RT) with Boutin’s fixative for GluR staining. For all other antibodies, 4% paraformaldehyde was used (25 minutes at RT). The fixed samples were subsequently washed with either 0.1% Triton X-100 in PBS (PBST) or with PBS alone for a detergent-free condition. The samples were then blocked with 5% normal goat serum in either PBST or PBS, as specified. Primary antibodies used were: Rabbit anti-pFAK, 1:250 (Invitrogen, 44-624G), Rabbit anit-HA,1:1000 (Sigma, H6908), Rabbit anti-GluRIIC, 1:1000 (Chang et al., 2024), Cy3/Alexa-647 conjugated anti-HRP, 1:100 for non-permeabilized staining or 1:250 for permeabilized staining (Jackson ImmunoResearch); rat anti-Elav, 1:500 (7E8A10, Developmental Studies Hybridoma Bank at the University of Iowa (DSHB)); mouse anti-Repo, 1:50 (8D12, DSHB); mouse anti-GFP, 1:100 (4C9, DSHB). The secondary antibodies used were Alexa Fluor 488 or 405 conjugated, diluted at 1:250 (Invitrogen). For experiments with glutamate incubation, dissected preps were incubated with 2mM Glutamate solution for 1hr or 10 min before performing stimulation protocol as indicated. Mock controls were treated the same way but without 2mM glutamate.

Imaging and image analysis

Images of synaptic boutons from muscle 6/7, A2 or A3 were taken using Olympus FV3000 confocal microscope at 60X with 1.6 zoom. Images showing expression patterns were taken at lower magnifications with either 10X or 60X objectives. To establish changes in bouton size and GluR level, mock controls (unstimulated larvae) were always dissected, immunostained, and imaged in parallel with stimulated genotypes for each experiment using the same conditions. Image analyses for GluRIIC, bouton size and extracellular staining of Shv and pFAK done using the same protocol as described (Lee et al., 2017).

Electrophysiology

Third-instar larvae were dissected and immersed in a modified HL-3 solution containing NaCl (70 mM), KCl (5 mM), MgCl2 (10 mM), sucrose (115 mM), HEPES (5 mM), trehalose (5 mM), and NaHCO3 (10 mM), at pH 7.2, with specified concentrations of CaCl₂ (0.25 or 0.5 mM). Current-clamp recordings were performed on muscle area 6 in abdominal segments A2 or A3, using suction electrodes to stimulate the severed ventral nerves with a 0.3 ms stimulus duration. The recording electrode, filled with 3 M KCl and having a resistance between 15 and 40 mΩ, was used to acquire data. Data with resting potentials more hyperpolarized than -60 mV were analyzed, while datasets were excluded if resting potentials deviated by more than 10 mV during recording or if there was a sudden drop in EPSP amplitude, indicating incomplete nerve function. The experimental setup included an Axopatch 200B Amplifier, a Digidata 1440A for digitization, and pClamp 10.3 software (Molecular Devices) for control. Data analysis was conducted using MiniAnalysis (Synaptosoft), Clampfit (Molecular Devices), and Microsoft Excel, with nonlinear summation applied to correct the average EPSP.

Western blot

Drosophila larval brain extract was obtained by homogenizing 6-10 larval brains collected on ice in RIPA buffer with EDTA (50 mM Tris-HCl, pH7.5, 1% NP-40, 0.5% NaDoc, 150 mM NaCl, 0.1% SDS, 10 mM EDTA, 50 mM NaF, 1 mM Na3VO4, 250 nM cycloporin A, protease inhibitor cocktail (Roche) using mortar and pestle. Protein homogenate was separated by 10% SDS-PAGE and transferred to nitrocellulose membranes. Primary antibodies were diluted in blocking solution as following: rabbit anti-Shv, 1:400 (Lee et al., 2017); Rabbit Anti-GFP,1:1000 (NOVUS, NB600-308), Anti-β-tubulin, 1:500 (E7, DSHB).

Statistical Analysis

All data are shown as mean ± SEM. Sample sizes, indicated in the graphs or figure legends, represent biological replicates and adhere to established standards in the literature. Comparisons between unstimulated and stimulated samples of the paired genotype were made using Student’s t-test. For comparisons involving multiple samples, one-way ANOVA followed by Tukey’s multiple comparison test was employed to determine statistical significance. To minimize bias, all samples were randomized during dissection, image collection, and data analysis.

Acknowledgements

We would like to thank Dr. Amita Seghal and Dr. Henry Y. Sun for the generous gift of repo-GeneSwitch-GAL4 and repo-LexA, respectively. K.T.C. is supported by NIH grants R01NS102260 and R01NS080946.

Additional information

Author contributions

Y-C.C. performed most of the experiments and data analyses. Y-J.P. performed the electrophysiology experiments. J.Y.L. designed and helped to generate the Shv-eGFP line using CRISPR/Cas9, K.T.C. conceived, designed, and supervised all aspects of the project. Y-C.C. and K.T.C. wrote the manuscript with inputs from Y-J.P. All authors reviewed and edited the manuscript.

Competing interest

The authors declare no competing interests.

Data availability

Additional information and requests for reagents used in this study are available upon request.

List of Supplementary Materials

Supplementary Figure 1.

Supplementary Figure 2.

Supplementary Figure 3.