Skip to main content
NCBI home page
eLife logoLink to eLife
. 2019 Jun 26;8:e46421. doi: 10.7554/eLife.46421

Imaging neuropeptide release at synapses with a genetically engineered reporter

Keke Ding 1,†, Yifu Han 2,3,†, Taylor W Seid 1, Christopher Buser 4, Tomomi Karigo 1, Shishuo Zhang 1, Dion K Dickman 2, David J Anderson 1,5,6,✉
Editors: Leslie C Griffith7, Eve Marder8
PMCID: PMC6609332  PMID: 31241464

Abstract

Research on neuropeptide function has advanced rapidly, yet there is still no spatio-temporally resolved method to measure the release of neuropeptides in vivo. Here we introduce Neuropeptide Release Reporters (NPRRs): novel genetically-encoded sensors with high temporal resolution and genetic specificity. Using the Drosophila larval neuromuscular junction (NMJ) as a model, we provide evidence that NPRRs recapitulate the trafficking and packaging of native neuropeptides, and report stimulation-evoked neuropeptide release events as real-time changes in fluorescence intensity, with sub-second temporal resolution.

Research organism: D. melanogaster

Introduction

Neuropeptides (NPs) exert an important but complex influence on neural function and behavior (Hökfelt et al., 2000; Insel and Young, 2000; Nässel and Winther, 2010; Bargmann and Marder, 2013). A major lacuna in the study of NPs is the lack of a method for imaging NP release in vivo, with subcellular spatial resolution and subsecond temporal resolution. Available techniques for measuring NP release include microdialysis (Kendrick, 1990), antibody-coated microprobes (Schaible et al., 1990) and GFP-tagged propeptides visualized either by standard fluorescence microscopy (van den Pol, 2012), or by TIRF imaging of cultured neurons (Xia et al., 2009). In Drosophila, a fusion between rat Atrial Natriuretic Peptide/Factor (ANP/F) and GFP was used to investigate neuropeptide trafficking at the fly neuromuscular junction (NMJ) (Rao et al., 2001). Release was measured indirectly, as a decrease in ANP-GFP fluorescence intensity at nerve terminals reporting residual unreleased peptide, on a time-scale of seconds (Wong et al., 2015). None of these methods combines NP specificity, genetically addressable cell type-specificity, high temporal resolution and applicability to in vivo preparations (Supplementary file 1). A major challenge is to develop a tool that encompasses all these features for direct, robust measurement of NP release in vivo.

Results

Neuropeptides are synthesized as precursors, sorted into dense core vesicles (DCVs), post-translationally modified and cleaved into active forms prior to release (Taghert and Veenstra, 2003). We reasoned that an optimal in vivo real-time NP release reporter should include (1) a reporter domain that reflects the physico-chemical contrast between the intravesicular milieu and the extracellular space (Figure 1—figure supplement 1A); and (2) a sorting domain that ensures its selective trafficking into DCVs (Figure 1—figure supplement 1b). The NP precursor may function as the sorting domain, suggested by studies of DCV fusion using pIAPP-EGFP (Barg et al., 2002) and NPY-pHluorin (Zhu et al., 2007) in cultured neurons, or ANP-GFP in Drosophila (Rao et al., 2001). We therefore developed a pipeline to screen various transgenes comprising NP precursors fused at different sites to fluorescent reporters, in adult flies (Figure 1—figure supplement 1B–C). A total of 54 constructs were tested. We found that optimal trafficking was achieved by substituting the reporter for the NP precursor C-terminal domain that follows the final peptide (Figure 1—figure supplement 1B). In order to maintain covalent linkage with the reporter domain, we removed the dibasic cleavage site C-terminal to the final peptide.

The DCV lumen has lower pH and free calcium (pH = 5.5–6.75, [Ca2+]~30 µM) compared to the extracellular space (pH = 7.3, [Ca2+]~2 mM) (Mitchell et al., 2001; Sturman et al., 2006). These differences prompted us to test validated sorting domains in a functional ex vivo screen using either pH-sensitive fluorescent proteins (Miesenböck et al., 1998) or genetically-encoded calcium indicators (GECIs) (Tian et al., 2012; Lin and Schnitzer, 2016) (Figure 1—figure supplement 1A–D). Reporters based on pHluorins (Miesenböck et al., 1998) did not perform well in our hands, therefore we focused on GCaMP6s (Chen et al., 2013). The calcium sensitivity threshold of GCaMP6s is below the calcium concentration in both DCVs and the extracellular space. However, GCaMP6s fluorescence is quenched in the acidic DCV lumen (Barykina et al., 2016), enabling it to function as a dual calcium/pH indicator (Figure 1A). These key properties should boost the contrast between GCaMP6s fluorescence in unreleased vs. released DCVs, potentially allowing us to trace NP release at the cellular level in vivo.

Figure 1. Design and Synaptic Localization of an NPRR.

(A) Schematic illustrating the principle of NPRRs (Neuropeptide Release Reporters). NPRR molecules in the DCV lumen (low pH/low calcium, left) exhibit increased fluorescence when released by fusion into the extracellular space (neutral pH/high calcium, right). NPRR fluorescent signal is expected to decay following diffusion into the synaptic cleft. New NPRR-containing DCVs are produced by synthesis and transport from the soma, not by recycling. NP: Neuropeptide. DCV: Dense Core Vesicle. SV: Synaptic Vesicle. (B) Distinct motor neuron subtypes at the Drosophila NMJ (muscle 12/13) have different proportions of DCVs vs. SVs. The GAL4 driver R57C10-Gal4 (nsyb-GAL4) labels all subtypes, while R20C11-GAL4 selectively labels only Type III neurons, which lack SVs (‘Type III-GAL4’). Light gray circles, black lines and dark gray shading represent boutons, inter-bouton intervals and subsynaptic reticulum respectively. The studies in this paper focus on Type Ib neurons and Type III neurons (in red rectangles). (C) Triple immunolabeling for GFP (green), Bursicon (blue) and vGluT (red), in flies containing nsyb-GAL4 driving UAS-GCaMP6s (upper), or NPRRANP (lower). Type Ib and Type III boutons are indicated. Scale bar, 5 µm. Inset image (NPRRANP, a-GFP channel) shows details of puncta distribution of NPRRANP in Type Ib neuron. Scale bar, 2 µm. (D) TEM images of boutons immunolabeled with anti-GFP (5 nm gold particle-conjugated) to detect nsyb>NPRRANP-GFP, which has an identical structure to NPRRANP, but is a GFP rather than GCaMP6s fusion to improve antigenicity (see Figure 1—figure supplement 4). Note strong labeling in DCVs (arrows) and the neuronal plasma membrane (arrowheads). Scale bar, 200 nm. Lower panel shows representative images of labeled DCVs. Scale bar,100 nm. (E) Quantification for TEM images in (D).

Figure 1—source data 1. Raw data for Immuno-EM experiments (ANP).
Figure 1—source data 2. Raw data for Immunno-EM experiments (control group).

Figure 1.

Figure 1—figure supplement 1. NPRR screening pipeline.

Figure 1—figure supplement 1.

A series of reporter-neuropeptide precursor fusions were designed, codon-optimized for Drosophila, cloned into expression vectors under the control of the GAL4 upstream activator sequence (UAS), and used to generate transgenic flies. (A) Candidate reporters interrogated included (constitutive) fluorescent reporters, genetically encoded calcium indicators (GECI) and pH indicators (pHluorins). (B) Sorting domain candidates included different truncated versions of rat Atrial Natriuretic Peptide (ANP; single-precursor-single-peptide) and Drosophila tachykinin (dTK; single-precursor-multiple-peptide) precursors. 52 constructs were built and injected. 44 of 54 were successfully integrated as transgenic lines, while eight were excluded due to lethality or unstable expression. (C–D) Candidate UAS-NPRR lines were crossed with an NPF-Gal4 driver line and selected based on their expression in NPF terminals in the adult fly brain. The raw fluorescence intensity of each NPRR candidate was measured using the same microscope parameters (laser power, HV, offset value). 14 candidates passed this screening. (C) We screened the performance of difference NPRRs (signal-to-noise contrast) by measuring fluorescence before and immediately after 70 mM high-potassium challenge in an ex vivo explant preparation of adult fly brains. The post/pre KCl fluorescence ratio is defined as ΔF/F. We arbitrarily set the threshold as 100%. 2 NPRRs with highest ΔF/F passed the final round of screening. Red asterisks indicate the candidates selected for the studies in Figure 2 and Figure 3. Blue asterisk indicates original ANP-GFP fusion) (Burke et al., 1997; Rao et al., 2001).

Figure 1—figure supplement 2. Exogeneous neuropeptide ANP dictates the expression pattern of NPRRANP.

Figure 1—figure supplement 2.

Membrane-bound mCD8::GFP fusion (A), cytosolic GCaMP6s (B) and NPRRANP (C) were expressed pan-neuronally in the larval NMJ and stained for both ANP (red) and NPRR (green, anti-GFP). (C) Note co-localization of ANP and GFP. Scale bar, 5 µm.

Figure 1—figure supplement 3. Expression of different reporters in Type III neurons in the larval NMJ.

Figure 1—figure supplement 3.

A GAL4 line (R20C11-GAL4, named Type III-GAL4 in this report) allows specific expression in Type III neurons. Expression patterns of (A) conventional GCaMP, (B) membrane-bound GFP, (C) NPRRdTK and (D) NPRRANP using Type III-GAL4. Arrows indicate boutons in Type III neurons, which contain the neuropeptide Bursicon. Note that anti-vGluT stains other types of motor neurons, which are not labeled by the Type III-specific driver used in this experiment. Scale bar, 5 µm.

Figure 1—figure supplement 4. Subcellular distribution of NPRRANP and NPRRANP-GFP.

Figure 1—figure supplement 4.

NPRRANP (labeled as NPRRANP-GCaMP6s) (A) and NPRRANP-GFP (B) were expressed pan-neuronally in the larval NMJ and double immune-labeled with antibodies to vGluT (red) and GFP (green). Note that the distribution of GFP signal is similar whether the reporter fusion is GCaMP6s (A) or GFP (B). Scale bar, 5 µm.
Open in a new tab

We sought to test several NP precursor-GCaMP6s fusion proteins, called NPRRs (NeuroPeptide Release Reporters; unless otherwise indicated all NPRRs refer to fusions with GCaMP6s), in an intact preparation using electrical stimulation to evoke release. Initially for proof-of-principle experiments, we used the Drosophila larval NMJ to test NPRRANP, a GCaMP6s fusion with rat ANP (Burke et al., 1997). NMJ terminals are large, individually identifiable, and easy to image and record. In particular, boutons on muscle 12/13 are diverse -- Type Ib and Type Is boutons contain mostly synaptic vesicles and few DCVs, while Type III boutons contain an abundance of DCVs but no synaptic vesicles (Menon et al., 2013); moreover, Type III-specific GAL4 drivers are available (Koon and Budnik, 2012) (Figure 1B).

Expression of NPRRANP pan-neuronally (under the control of nsyb-GAL4) followed by double immuno-staining for ANP and GCaMP (anti-GFP) indicated that the sorting domain and the reporter domains showed a similar localization in Type III neurons (Figure 1—figure supplement 2). Moreover, the distribution of NPRRANP overlapped that of Bursicon (Figure 1—figure supplement 3D), an NP that is endogenously expressed in Type III neurons (Loveall and Deitcher, 2010). Both GCaMP and Bursicon immunoreactivity were strongest within boutons, consistent with the known subcellular localization of DCVs (Gorczyca and Budnik, 2006).

Glutamate is the only known canonical neurotransmitter used at the larval NMJ (Menon et al., 2013). This allowed visualization of the subcellular localization of small synaptic vesicles (SV) by immuno-staining for vGluT, a vesicular glutamate transporter (Fremeau et al., 2001; Kempf et al., 2013). In Type Ib neurons (which contain relatively few DCVs relative to SVs [Menon et al., 2013]), vGluT staining was observed as patches with a dim center, which may reflect clustered SVs, while NPRRANP immunoreactivity was seen in dispersed, non-overlapping punctae (Figure 1C, α-GFP, inset). In Type III neurons, NPRRs were strongly expressed but no vGluT immunoreactivity was detected (Figure 1C). The subcellular distribution of this NPRR in larval NMJ neurons, therefore, is similar to that of other DCV-targeted markers previously used in this system (Rao et al., 2001; Shakiryanova et al., 2006), and appears to reflect exclusion from SVs.

The diffraction limit of light microscopy precluded definitive co-localization of NPRRs in DCVs. Therefore, we employed Immuno-Electron microscopy (Immuno-EM) to investigate the subcellular localization of NPRRs at the nanometer scale. To maximize antigenicity for Immuno-EM, we generated constructs that replaced GCaMP6s with GFP (NPRRANP-GFP;). NPRRANP-GFP showed dense labeling in association with DCVs (Figure 1D, arrows), where the average number of gold particles/µm2 was substantially and significantly higher than in neighboring bouton cytoplasm (DCV/Bouton ~ 14.26) (Figure 1E, Supplementary file 2). Taken together, these data indicate that NPRRANP-GFP is localized to DCVs. By extension, they suggest that NPRRANP-GCaMP6s (which has an identical structure to NPRRANP-GFP except for the modifications that confer calcium sensitivity) is similarly packaged in DCVs. While these two reporters show indistinguishable distributions by immunofluorescence (Figure 1—figure supplement 4), we cannot formally exclude that the substitution of GCaMP for GFP may subtly alter subcellular localization of the NPRR in a manner undetectable by light microscopy.

To measure the release of NPRRs from DCVs, we next expressed NPRRANP in Type III neurons using a specific GAL4 driver for these cells (Koon and Budnik, 2012) (Figure 2E and Figure 1—figure supplement 3D). We delivered 4 trials of 70 Hz electrical stimulation to the nerve bundle, a frequency reported to trigger NP release as measured by ANF-GFP fluorescence decrease (Rao et al., 2001; Shakiryanova et al., 2006), and used an extracellular calcium concentration that promotes full fusion mode (Alés et al., 1999). This stimulation paradigm produced a relative increase in NPRRANP fluorescence intensity (ΔF/F), whose peak magnitude increased across successive trials (Figure 2A, red bars and 2D; Video 1; Figure 2—figure supplement 1, A1 vs. A7). Responses in each trial showed a tri-phasic temporal pattern: (1) In the ‘rising’ phase, NPRRANP ∆F/F peaked 0.5–5 secs after stimulation onset, in contrast to the virtually instantaneous peak seen in positive control specimens expressing conventional GCaMP6s in Type III neurons (Figure 2A–B). The NPRRANP latency to peak was similar to the reported DCV fusion latency following depolarization in hippocampal neurons (Xia et al., 2009). This delay is thought to reflect the kinetic difference between calcium influx and DCV exocytosis due to the loose association between DCVs and calcium channels (Xia et al., 2009). (2) In the ‘falling’ phase, NPRRANP ∆F/F began to decline 1–5 s before the termination of each stimulation trial, presumably reflecting depletion of the available pool of releasable vesicles. In contrast, GCaMP6s fluorescence did not return to baseline until after stimulation offset (Figure 2A–B). (3) Finally, unlike GCaMP6s, NPRRANP exhibited an ‘undershoot’ (∆F/F below baseline) during the post-stimulation intervals, followed by a ‘recovering’ phase (Figure 2A; Figures 2C,I1–4). This undershoot may reflect dilution of released fluorescent NPRR molecules by diffusion into the synaptic cleft (van den Pol, 2012), while recovery may reflect DCV replenishment in the boutons from vesicles proximal to the imaged release site.

Figure 2. NPRR specifically reports neuropeptide release.

(A) Trace from a representative experiment showing changes in NPRRANP fluorescence intensity (∆F/F) in Type III motor neurons at the larval NMJ evoked by electrical stimulation. BG: background. S1-S4: Stimulation trials 1–4. I1-I4: Inter-stimulation Intervals (ISIs) 1–4. Green line: ∆F/F averaged across all boutons in the field of view. Gray shading: s.e.m envelope. Red bar: electrical stimulation trials (70 Hz). The three typical phases of the response are indicated in S4. The peak height of the response on the first trial is characteristically lower (see also (D)), and may reflect competition with unlabeled DCVs in the readily releasable pool. (B) ΔF/F traces in control flies expressing cytoplasmic GCaMP6s in Type III neurons. (C) Integrated NPRRANP ΔF/F values during trials S1-4 and intervals I1-4. A.U.: arbitrary units. n = 8. ***, p<0.001. (D) Average NPRRANP ΔF/F peak heights for trials S1-4. n = 8. *, p<0.05. Plotted values in (C–D) are mean ± s.e.m. (E1–E2) Representative selection of ROIs (yellow). Details see Materials and methods. Scale bar, 5 µm. (F) NPRRANP ΔF/F response are abolished in Type III GAL4>UAS NPRRANP flies bearing UAS-TNT (F1) but not UAS-TNTimp (F2). (G) Average peak heights of NPRRANP ΔF/F in combined stimulation trials (S1-4) from (F). ****, p<0.0001. (H) Average ‘undershoot’, defined as the integrated ΔF/F during ISIs I1-4 (see (C)). In (C–D) and (G–H).

Figure 2.

Figure 2—figure supplement 1. Activation of NPRRANPin situ.

Figure 2—figure supplement 1.

Representative still frames (A1–A12) from video recordings of NPRRANP-expressing Type III neurons at the larval NMJ. ‘On’ (A2,4,7,10) represents the onset of electrical pulses. Color bar: Raw fluorescence intensity. Scale bar, 50 µm.

Figure 2—figure supplement 2. NPRR specifically reports neuropeptide release.

Figure 2—figure supplement 2.

(A) Left: Segmentation of Type III neurons into boutons (orange) and inter-bouton intervals (IBIs, red). Right: Schematic illustrating DCV distribution in Type III neurons, based on photomicrograph to the left. Green dots, DCVs. (B) Average time-integrated ratio of ΔF/F in boutons/IBIs (Materials and Methods), within each stimulation periods. n.s., not significant. *, p<0.05. ***, p<0.001. ****, p<0.0001.(C) TNT does not affect GCaMP6s ΔF/F kinetics. n = 6–7. GCaMP6s peak magnitudes were reduced slightly in TNT (C1) in comparison to TNTimp (C2) preparations, perhaps reflecting partial vulnerability of the cytosolic GCaMP6s reporter to TNT-mediated cleavage and degradation. NPRRs are expected to be protected from TNT by the DCV membrane.

Figure 2—figure supplement 3. Blocking DCV fusion using Tetanus Toxin.

Figure 2—figure supplement 3.

(A1, A2) Tetanus toxin (TNT) blocks vesicle fusion by cleavage of n-synaptobrevin (n-syb).
Open in a new tab

Figure 4. NPRR reveals distinct cell-type specific peptide release properties.

For each preparation, a series of stimulation trials were delivered at frequencies from 1 Hz to 70 Hz, as indicated. In-stimulation response peaks were normalized to 70 Hz. The normalized peaks of NPRRs or calcium responses (measured with cytosolic GCaMP6s) were pooled and plotted for both Type III (A-C) and Type Ib (D-F) neurons. Responses were compared to zero. n = 6–12. n.s., not significant. *, p<0.05. **, p<0.01. ***, p<0.001. ****, p<0.0001.

Figure 4.

Figure 4—figure supplement 1. Comparison of NPRR response at 30 and 50 Hz.

Figure 4—figure supplement 1.

Normalized ΔF/F peaks in at 30 Hz (A) and 50 Hz (B) electrical stimulation in Figure 4A,B,D,E are replotted and compared. n = 6–7. *, p<0.05. **, p<0.01. n.s., not significant.
Open in a new tab

Video 1. Activation of NPRRANPin situ.

Download video file (690.8KB, mp4)
Open in a new tab

Representative video recording of NPRRANP-expressing Type III neurons at the larval NMJ. In the preparation, the Type III neuron was stimulated as described in Figure 2A.

Because NPRRANP fluorescence was preferentially accumulated within boutons, we asked whether these regions contributed to ∆F/F peaks more significantly than the inter-bouton intervals (IBIs). To do this, we partitioned the processes into boutons and IBI fields (Figure 2—figure supplement 2A), and compared the ∆F/F in these regions during stimulation trials. The time-averaged ratio of bouton/IBI ΔF/F (see Materials and Methods) was significantly higher for NPRRANP than for GCaMP6s, particularly during later stimulation trials (Figure 2—figure supplement 2B, green bars, S2-4). This contrast indicates that NPRRANP signals are preferentially observed in boutons, where DCVs are located, and do not reflect differences in cytoplasmic free Ca2+ levels between these regions as detected by GCaMP6s.

To test definitively if NPRRANP ∆F/F signals are dependent upon NP release, we blocked vesicle fusion at terminals of Type III neurons using expression of tetanus toxin light chain (TNT) (Sweeney et al., 1995), a protease that cleaves n-synaptobrevin, a v-snare required for DCV fusion (Figure 2—figure supplement 3) (Xu et al., 1998). As a control, we used impotent TNT (TNTimp), a reduced activity variant (Sweeney et al., 1995). TNT expression completely abolished stimulation-induced ∆F/F increases from NPRRANP, while TNTimp did not (Figure 2F). Further analysis revealed that both the ∆F/F peaks and inter-stimulation undershoots were diminished by TNT (Figure 2G–H). In contrast, neither TNT nor TNTimp affected the kinetics of GCaMP6s signals in Type III neurons (Figure 2—figure supplement 2C), which report cytosolic Ca2+ influx. Taken together, these data support the idea that NPRRANP signals specifically reflect DCV release.

ANP is a rat NP that lacks a Drosophila homolog (Rao et al., 2001). To determine whether our method could be applied to detect the release of a specific, endogenous fly NP, we tested NPRRdTK, one of 6 different reporter variants we initially generated from the Drosophila neuropeptide precursor, DTK (Figure 1—figure supplement 1B). In contrast to ANP which encodes a single peptide, DTK yields multiple NP derivatives (Winther et al., 2003). Light microscopy (Figure 3A) and Immuno-EM (Figure 3B, arrows) confirmed that NPRRdTK, like NPRRANP, was localized to DCVs (DCV/bouton ~ 22.19, Figure 3C). Using the Type III-specific GAL4 driver to express NPRRdTK and the same stimulation protocol as used for NPRRANP, the basic tri-phasic response profile was also observed (Figure 3D). However, peak heights and baseline fluorescence fell progressively with successive stimulation trials (Figure 3E), in contrast to NPRRANP where the first peak and undershoot were lower (Figure 2C–D). The reason for this difference is currently unclear.

Figure 3. Application of the NPRR approach to a Drosophila neuropeptide.

Figure 3.

Open in a new tab

(A) Triple immunolabeling for GFP (green), Bursicon (blue) and vGluT (red) in Type III-GAL4> UAS NPRRdTK flies. Scale bar, 5 µm. (B) TEM images of boutons immunolabeled against GFP (5 nm gold) in nsyb-GAL4>UAS NPRRdTK-GFP flies. Note strong labeling in DCVs (arrows) and bouton plasma membrane (arrowheads). Scale bar, 200 nm. Lower panel shows representative images of labeled DCVs. Scale bar,100 nm. (C) Quantification of TEM images in (B). (D) NPRRdTK ΔF/F curve; stimulation conditions as in Figure 2A. (E) Average NPRRdTK ΔF/F peak height above pre-stimulation baseline (corrected; see Materials and methods) for stimulation trials S1-4. n = 6. **, p<0.01.

Figure 3—source data 1. Raw data for Immuno-EM experiments (dTK).

We next investigated the relationship between NPRR signal and stimulation intensity, by delivering to the Type III neurons a series of low to high frequency electrical stimuli (1–70 Hz; Levitan et al., 2007) while imaging the nerve terminals. For direct comparison of NPRR responses across different preparations, we applied a posteriori normalization of fluorescent peaks in each trial to the highest response obtained among all trials. For both NPRRANP and NPRRdTK (Figure 4A–B), the peak responses showed a positive correlation with stimulation frequency, analogous to that observed using cytosolic GCaMP6s (Figure 4C). In Type III neurons, the responses of both NPRRs to stimulation frequencies < 30 Hz (1,5,10,20 Hz) were not statistically significant from zero. NPRRANP showed a higher sensitivity to high stimulation frequencies (30 Hz: 18.14%, 50 Hz: 82.40% Normalized peak ∆F/F), while NPRRdTK showed a higher stimulation threshold and lower sensitivity (30 Hz: 3.57%, 50 Hz: 24.67% Normalized peak ∆F/F).

We next investigated whether the relatively high stimulation frequency required to observe significant responses with NPRRs was a function of the reporters, or rather of the cell class in which they were tested. To do this, we expressed both NPRRs in Type Ib neurons, a class of motor neurons that contains both SVs and DCVs (Figure 1B, Figure 4D–F), and performed stimulation frequency titration experiments. Strikingly, in Type Ib neurons, significant increases in ∆F/F could be observed at frequencies as low as 10 Hz (Figure 4D,E; NPRRANP @ 20 Hz: 12.50%, NPRRdTK @ 20 Hz: 17.67% normalized peak ∆F/F). The reason for the difference in NPRR threshold between Type III and Type Ib neurons is unknown, but parallels their difference in GCaMP6s response to electrical stimulation (Figure 4C vs. Figure 4F).

Notably, although NPRRANP and NPRRdTK presented distinct response profiles in Type III neurons, their performance in Type Ib neurons was more similar (Figure 4A vs. Figure 4B; cf. Figure 4D vs. Figure 4E). In summary, the differences in performance we observed between the two NPRRs appeared to be specific to Type III neurons, and were minor in comparison to the differences in performance of both reporters between the two cell classes. The reason for the differences between NPRRANP and NPRRdTK sensitivity and kinetics in Type III neurons is unknown but may reflect differences in how well these reporters compete with the high levels of endogenous neuropeptide (Bursicon) for packaging, transport or release.

Discussion

Here we present proof-of-principle for a method to detect the release of different neuropeptides in intact neural tissue, with subcellular spatial and sub-second temporal resolution. By exploiting the fluorescent change of GCaMP in response to a shift in pH and [Ca2+], we visualized the release of neuropeptides by capturing the difference between the intravesicular and extracellular microenvironment. NPRR responses exhibited triphasic kinetics, including rising, falling and recovering phases. In the falling phase, a post-stimulus ‘undershoot’, was observed in which the fluorescent intensity fell below pre-stimulation baseline. This undershoot presumably reflect the slow kinetics of DCV replenishment relative to release.

The molecular mechanisms of NP release are incompletely understood (Xu and Xu, 2008). It is possible that individual DCVs only unload part of their cargo during stimulation, in which case many DCVs that underwent fusion may still contain unreleased NPRR molecules following a stimulus pulse. Although we are convinced that NPRR signals do indeed reflect NP release, due to the presence of the recovering phase, we cannot formally exclude that unreleased NPRRs may contribute to the signal change due to their experience of intravesicular [Ca2+]/pH changes that occur during stimulation. To resolve this issue in the future, an ideal experiment would be to co-express an NPRR together with a [Ca2+]/pH-invariant NP-reporter fusion. Multiple attempts to generate such fusions with RFP were unsuccessful, due to cryptic proteolytic cleavage sites in the protein which presumably result in degradation by DCV proteases during packaging.

To test if NPRRANP ∆F/F signals are dependent on NP release, we expressed the light chain of tetanus toxin (TNT), a reagent shown to effectively block NP release in many (McNabb and Truman, 2008; Hentze et al., 2015; Zandawala et al., 2018), if not all (Umezaki et al., 2011), systems. We observed a striking difference in NPRR kinetics in flies co-expressing TNT vs. its proteolytically inactive ‘impotent’ control form TNTimp (Figure 2F). The strong reduction of NPRR signals by TNT-mediated n-syb cleavage is consistent with the idea that these signals reflect the release of NPRRs from DCVs.

We have tested the generalizability of the principles used to generate NPRRs by (1) constructing a surrogate NP reporter NPRRANP as well as a multi-peptide-producing endogenous Drosophila NP reporter NPRRdTK (Figures 2–3); (2) characterized NPRR signals in response to varying intensities of electrical stimulation; and (3) recorded NPRR signals in two different classes of NMJ motor neurons containing DCVs with or without SVs, respectively (Figure 4). These experiments revealed, to our surprise, that NPRR responses exhibit cell-type specific characteristics (Figure 4). As NPRRs are applied to other neuropeptides and cell types, a systematic characterization of neuropeptide release properties in different peptidergic neurons should become possible, furthering our understanding of neuropeptide biology.

The method described here can, in principle, be extended to an in vivo setting. This would open the possibility of addressing several important unresolved issues in the study of NP function in vivo. These include the ‘which’ problem (which neuron(s) release(s) NPs under particular behavioral conditions?); the ‘when’ problem (when do these neurons release NPs relative to a particular behavior or physiological event?); the ‘where’ problem (are NPs released from axons, dendrites or both?); and the ‘how’ problem (how is NP release regulated?). The application of NPRRs to measuring NP release dynamics in awake, freely behaving animals may yield answers to these important long-standing questions.

Materials and methods

Key resources table.

Reagent type
(species) or resource		 Designation Source or reference Identifiers Additional
information
Genetic reagent (D. melanogaster) UAS-NPRRANP (attp2) this paper See Materials and methods, subsection Construction of transgenic animals.
Genetic reagent (D. melanogaster) UAS-NPRRdTK (attp2) this paper Same as above.
Genetic reagent (D. melanogaster) UAS-TNTimp Bloomington Drosophila Stock Center BDSC:28840; FLYB:FBti0038575; RRID:BDSC_28840 Flybase symbol: w[*]; P{w[+mC]=UAS TeTxLC.(-)V}A2
Genetic reagent (D. melanogaster) UAS-TNT Bloomington Drosophila Stock Center BDSC:28838; FLYB:FBti0038527; RRID:BDSC_28838 Flybase symbol: w[*]; P{w[+mC]=UAS TeTxLC.tnt}G2
Genetic reagent (D. melanogaster) w; +; UAS-GCaMP6s (su(Hw)attp1) Hoopfer et al., 2015
Antibody anti-GFP (chicken polyclonal) Aveslab Aveslab: GFP-1020; RRID:AB_2307313 (1:250:Immuno-EM, 1:1000: IHC)
Antibody anti-ANP (rabbit polyclonal) abcam abcam #14348 (1:500)
Open in a new tab

Fly strains

All experimental flies were reared on a 12/12 hr day-night cycle at 25°C. Standard chromosomal balancers and genetic strategies were used for all crosses and for maintaining mutant lines. Detailed genotypes used are summarized in Supplementary file 3. The following strains were obtained from Bloomington Stock Center (Indiana University): R20C11-Gal4 (#48887), R57C10-Gal4 (#39171), UAS-mCD8::GFP (#32185), UAS-TNT (#28838), UAS-TNTimp (#28840). UAS-opGCaMP6s was made by Barret Pfeiffer (Gerald Rubin’s lab, Janelia Farm) (Hoopfer et al., 2015).

Construction of transgenic animals

All PCR reactions were performed using PrimeSTAR HS DNA polymerase (Takara #R045Q). All constructs were verified via DNA sequencing (Laragen).

To construct UAS-NPRRANP, Drosophila codon-optimized ANP and GCaMP6s were synthesized using gBlocks service (Integrated DNA Technologies), and subcloned into pJFRC7 vector (from Addgene #26220) (Pfeiffer et al., 2010) using Gibson cloning. UAS-dTK-NPRR is built in a similar way except the dTK fragment was cloned from the Drosophila brain cDNA. NPRRdTK-GFP and NPRRANP-GFP were built similarly except Drosophila codon-optimized GFP was used for the subcloning. All the vectors were injected and integrated into attP2 or attp40 sites (Bestgene Inc; see Supplementary file 3 for attP sites used for each genotype employed).

Expression screening of NPRR candidates

Adult fly brains were dissected in chilled PBS and fixed in 4% formaldehyde for 55 min at room temperature. After three 10 min rinses with PBS, the brains were cleared with Vectashield (#1000, Vectorlabs), mounted, and used for native fluorescence measurements. We trace the NPF neuron somata and arborization as ROIs. We selected regions next to NPF neurons and measured its fluorescent intensity as a reference, which represents background autofluorescence. Candidates whose fluorescence reached at least 2-fold higher than reference were selected for functional screening.

Functional screening of NPRR candidates

For the baseline fluorescence measurement, we crossed NPF-Gal4 to the candidate lines and generated NPF-Gal4>NPRRx (x = candidate label) flies for tests. The dissected adult fly brains were mounted on a petri dish and immersed in Drosophila imaging saline (108 mM NaCl, 5 mM KCl, 2 mM CaCl2, 8.2 mM MgCl2, 4 mM NaHCO3, 1 mM NaH2PO4, 5 mM trehalose, 10 mM sucrose, 5 mM HEPES, pH 7.5). To deliver high potassium challenge, High-K imaging saline was perfused (43 mM NaCl, 70 mM KCl, 2 mM CaCl2, 8.2 mM MgCl2, 4 mM NaHCO3, 1 mM NaH2PO4, 5 mM trehalose, 10 mM sucrose, 5 mM HEPES, pH 7.5). Live imaging series were acquired using a Fluoview FV3000 Confocal laser scanning biological microscope (Olympus) with a 40×, 0.8 N.A. (Numerical Aperture) water immersion objective (Olympus). Candidates whose post-stimulation fluorescence reached at least 2-fold of baseline fluorescence (measured as averaged pre-stimulation fluorescence) were selected for in vivo tests at NMJ. For each candidate line, at least three brains were tested and fold-change of each was averaged.

Immunohistochemistry

Larval dissection was performed in chilled HL3 solution (70 mM NaCl, 5 mM KCl, 20 mM MgCl2, 10 mM NaHCO3, 115 mM sucrose, 5 mM trehalose, 5 mM HEPES and 1.5 mM CaCl2, pH 7.2). Dissected tissues were fixed in 4% formaldehyde or Bouin’s solution for 30 min at room temperature. After three 15 min rinses with PBS, tissues were incubated with primary antibodies overnight at 4°C. Following three 15 min rinses with PBS, tissues were incubated with secondary antibody for 2 hr at room temperature. Following three 15 min rinses, tissues were cleared with Vectashield (#1000, Vectorlabs) and mounted. Confocal serial optical sections were acquired using a Fluoview FV3000 Confocal laser scanning biological microscope (Olympus) with a 60×, 1.30 N.A. silicone oil objective (Olympus). All image processing and analyses were done using ImageJ (National Institute of Health).

The following primary antibodies were used: Chicken anti-GFP (1:250-1:1000, Aveslab #1020), Rabbit anti-ANP (1:500, abcam #14348), Guinea pig anti-vGluT (Goel and Dickman, 2018) (1:1500), Rabbit anti-syt1 (Littleton et al., 1993) (1:500) and Rabbit anti-Bursicon (1:2000, a gift from Dr. Benjamin White).

The following secondary antibodies were used: Alexa Fluor 488 Goat anti-Chicken IgY (#A11039, Invitrogen), Alexa Fluor 488 Goat anti-Rabbit IgG (#A11008, Invitrogen), Alexa Fluor 568 Goat anti-Rabbit IgG(H + L) (#A11011, Invitrogen), Alexa Fluor 633 Goat anti-Rabbit IgG(H + L) (#A21070, Invitrogen), Alexa Fluor 488 Goat anti-Guinea Pig IgG(H + L) (#A11073, Invitrogen), Alexa Fluor 568 Goat anti-Guinea Pig IgG(H + L) (#A11075, Invitrogen), Alexa Fluor 568 Goat anti-Mouse IgG(H + L) (#A11004, Invitrogen) and Alexa Fluor 633 Goat anti-Mouse IgG(H + L) (#A21050, Invitrogen).

Electron microscopy

Drosophila tissues were fixed in 4% formaldehyde in PBS and stored at 4°C until preparation by high-pressure freezing (HPF) and freeze-substitution (FS) (Buser and Walther, 2008; Buser and Drubin, 2013). Tissues were cryoprotected in 2.3 M sucrose for 45 min, transferred to 200 µm deep planchettes and high-pressure frozen in an EMPact2 with RTS (Leica, Vienna, Austria). FS was carried out in an AFS2 (Leica, Vienna, Austria) in methanol containing 5% water, 0.05% glutaraldehyde and 0.1% uranyl acetate (−90°C, 3 hr; −90 to −80°C, 10 hr; −80°C, 4 hr; −80°C to 4°C, 24 hr). Samples were washed once in methanol containing 5% water, infiltrated with hard grade LR White (Electron Microscopy Sciences, Hatfield, PA, USA) at 4°C ([LR White]: [methanol containing 5% water] 1:1, 24 hr; 100% LR White, 3 × 24 hr) and polymerized in a fresh change of LR White using a Pelco BioWave (Ted Pella, Inc, Redding, CA, USA) set to 750 W, 95°C for 45 min.

60 nm thin sections (UCT ultramicrotome, Leica, Vienna, Austria) were picked up on formvar-coated 50 mesh copper grids. The sections were blocked for 3 min in blocking buffer (PBS with 0.5% bovine serum albumin, which was used for all antibody dilutions), incubated in anti-GFP antibody (1:500, Aveslab #1020) for 5 min, washed 3 times in blocking buffer, incubated in rabbit anti chicken antibody (1:50, MP Biomedicals #55302) for 5 min, washed 3 times on blocking buffer, incubated on protein A - 5 nm gold (1:50, Utrecht, Netherlands), and washed 3 times in PBS and 3 times in distilled water. The sections were stained in uranyl acetate or uranyl acetate and Reynolds lead citrate depending on the desired contrast and imaged at 80 kV in a Zeiss EM10C (Zeiss, Oberkochen, Germany) using a CCD camera (Gatan, Pleasanton, CA, USA).

Labeling density was estimated using stereological methods (Griffiths and Hoppeler, 1986). Cross-sections through boutons were recorded and the following parameters were measured: total image area, total number of gold particles, number of visible dense core vesicles (DCV), number of gold particles within a 50 nm radius of the DCV center, bouton area (grid intersection estimate), gold within the bouton cytoplasm, gold within 20 nm of the bouton plasma membrane, gold outside of the bouton (mainly sER). Background labeling was estimated using internal controls (labeling on blank resin and on muscle fibers) and a biological control (non-GFP expressing genotype). Occasional obvious, large gold aggregates were disregarded. Background was consistently below 0.6 gold/µm2 in independently repeated labeling experiments.

Electrical stimulation

The dissection of third-instar larvae was performed in zero-calcium HL3 saline. The CNS was removed to avoid spontaneous motor neuron activity. To minimize muscle contraction induced by electrical stimulation of motor neurons, the larval body walls were slightly stretched and incubated in HL3 saline supplemented with 10 mM glutamate for 5 mins after dissection to desensitize postsynaptic glutamate receptors. Samples were then shifted to HL3 saline containing 1 mM glutamate and 1.5 mM Ca2+. Motor nerves were sucked into a glass micropipette with a stimulation electrode. In Figure 2 and Figure 3, to induce maximum dense core vesicle release at type III motor neuron terminals, four repetitive bursts (70 Hz stimulation for 18–20 s with pulse width of 1 ms) with intervals of 40–42 s were programmed and triggered with a Master-9 stimulator (A.M.P.I., Israel) connected to an iso-flex pulse stimulator (A.M.P.I., Israel). The stimulation intensity was tested and set to double the intensity required to induce muscle contraction by a single pulse stimulation.

In Figure 4, stimulation trials were delivered with the same duration, but with a series of frequencies spanning 1 Hz to 70 Hz.

Calcium imaging

A Nikon A1R confocal microscope with resonant scanner and NIS Element software were used to acquire live Ca2+ imaging on third instar larvae, bathed with 1 mM glutamate added in 1.5 mM Ca2+ HL3 saline. Type III motor neuron terminals in abdominal segments from A2 to A5 were imaged using a 60x APO 1.4 N.A. water immersion objective with 488 nm excitation laser. A 5 min period was used for time-lapse imaging at a resonance frequency of 1 fps (512 × 512 pixels or 1024 × 1024 pixels), with z-stacks (step length varying from 1 to 1.5 μm) covering the depth of entire type III motor neuron terminals. The repetitive electrical stimulation of 70 Hz was delivered during the imaging session. Samples with severe muscle contractions were abandoned due to imaging difficulties. Maximum intensity projection (MIP) and image registration were conducted using Image J. Plugins including Image Stabilizer (K. Li, CMU) and Template Matching (Q. Tseng) were used for compensating drifting and correcting movement induced by electrical stimulations. ROIs were manually selected by tracing the outer edge of each neuron based on the baseline fluorescence. If the fluorescence was too weak to trace, we established a reference stack by empirically adjusting the contrast on a duplicate of the raw image stack. We used the reference stack for ROI selection and projected the selected ROIs back onto to the raw image stack for measurement. For frames in which the sample movement could not be automatically corrected, we manually outlined the ROIs used for measurements. Preparations with severe movement or deformation artifacts were abandoned to avoid unreliable measurements. Each ROI represent a traceable neuronal branch except Figure 2—figure supplement 2B, in which the ROIs were further manually partitioned into boutons and IBIs (Inter-Bouton Intervals) based on morphology. Fluorescence change were normalized to the pre-stimulation background except for Figure 3E, for which the data in each trial was normalized to the average ∆F/F during a 5 s period just before stimulation was initiated. No sample size is predetermined based on statistics. Ca2+ imaging data were acquired from at least six independent NMJs from at least five animals.

Statistical analysis

Data are presented as mean ± s.e.m. All data analysis was performed with Graphpad Prism 6, Microsoft Excel and custom Matlab codes (Source code 1). Mann-Whitney U test was used for comparison except in Figure 4, where One-sample T test was used for comparison with a specified value (0).

Acknowledgements

We thank T Südhof for advice and encouragement at the inception of this project, T Littleton for suggesting GCaMP as a potential reporter for DCV release, B Pfeiffer for advice on construct design, members of the Anderson laboratory for critical feedback, K Zinn for helpful comments on the manuscript, P Goel for the information on antibodies, B Kiragasi for Master-9 programming, K Chen and D Ma for advice on imaging analysis and programming, G Mancuso and C Chiu for administrative assistance and lab management, and A Sanchez for maintenance of fly stocks. This work was supported by NIH BRAIN Initiative grant R21EY026432 and R01 DA031389 (to DJA) and NS091546 (to DD). DJA is an Investigator of the Howard Hughes Medical Institute.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

David J Anderson, Email: wuwei@caltech.edu.

Leslie C Griffith, Brandeis University, United States.

Eve Marder, Brandeis University, United States.

Funding Information

This paper was supported by the following grants:

  • National Institutes of Health R21EY026432 to David J Anderson.

  • National Institutes of Health R01DA031389 to David J Anderson.

  • National Institutes of Health NS091546 to Dion K Dickman.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing—original draft, Writing—review and editing.

Data curation, Formal analysis, Investigation, Methodology, Writing—review and editing.

Validation, Investigation, Methodology.

Data curation, Formal analysis, Investigation, Methodology.

Investigation, Methodology.

Validation, Investigation.

Resources, Formal analysis, Supervision, Funding acquisition, Methodology, Writing—review and editing.

Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing—original draft, Project administration, Writing—review and editing.

Additional files

Source code 1. Code for data analysis.
elife-46421-code1.zip (1.8KB, zip)
Supplementary file 1. Current techniques for neuropeptide release measurement.

Summary of current techniques used for neuropeptide release, including microdialysis, antibody-coated microprobes, GFP-tagged propeptide imaging and NPRR. NA, ‘not applicable’

elife-46421-supp1.xlsx (10.6KB, xlsx)
Supplementary file 2. Stereological labeling estimates.

Stereological labeling estimations of NPRRANP-GFP and NPRRdTK-GFP, respectively, in Type Ib neurons, or in Type Ib and Type III neurons. Biological controls and internal controls are described in Materials and methods. SNR: Signal-to-Noise Ratio.

elife-46421-supp2.docx (15.9KB, docx)
Supplementary file 3. Complete genotypes of transgenic flies used in this study.

Summary of complete genotypes of transgenic flies used in this study.

elife-46421-supp3.docx (17.4KB, docx)
Transparent reporting form

Data availability

Source data of EM for Figure 1 and 3, and codes used for Figure 2 and 3 have been provided.

References

  1. Alés E, Tabares L, Poyato JM, Valero V, Lindau M, Alvarez de Toledo G. High calcium concentrations shift the mode of exocytosis to the kiss-and-run mechanism. Nature Cell Biology. 1999;1:40–44. doi: 10.1038/9012. [DOI] [PubMed] [Google Scholar]
  2. Barg S, Olofsson CS, Schriever-Abeln J, Wendt A, Gebre-Medhin S, Renström E, Rorsman P. Delay between fusion pore opening and peptide release from large dense-core vesicles in neuroendocrine cells. Neuron. 2002;33:287–299. doi: 10.1016/S0896-6273(02)00563-9. [DOI] [PubMed] [Google Scholar]
  3. Bargmann CI, Marder E. From the connectome to brain function. Nature Methods. 2013;10:483–490. doi: 10.1038/nmeth.2451. [DOI] [PubMed] [Google Scholar]
  4. Barykina NV, Subach OM, Doronin DA, Sotskov VP, Roshchina MA, Kunitsyna TA, Malyshev AY, Smirnov IV, Azieva AM, Sokolov IS, Piatkevich KD, Burtsev MS, Varizhuk AM, Pozmogova GE, Anokhin KV, Subach FV, Enikolopov GN. A new design for a green calcium indicator with a smaller size and a reduced number of calcium-binding sites. Scientific Reports. 2016;6:34447. doi: 10.1038/srep34447. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Burke NV, Han W, Li D, Takimoto K, Watkins SC, Levitan ES. Neuronal peptide release is limited by secretory granule mobility. Neuron. 1997;19:1095–1102. doi: 10.1016/S0896-6273(00)80400-6. [DOI] [PubMed] [Google Scholar]
  6. Buser C, Drubin DG. Ultrastructural imaging of endocytic sites in saccharomyces cerevisiae by transmission electron microscopy and immunolabeling. Microscopy and Microanalysis. 2013;19:381–392. doi: 10.1017/S1431927612014304. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Buser C, Walther P. Freeze-substitution: the addition of water to polar solvents enhances the retention of structure and acts at temperatures around -60 degrees C. Journal of Microscopy. 2008;230:268–277. doi: 10.1111/j.1365-2818.2008.01984.x. [DOI] [PubMed] [Google Scholar]
  8. Chen TW, Wardill TJ, Sun Y, Pulver SR, Renninger SL, Baohan A, Schreiter ER, Kerr RA, Orger MB, Jayaraman V, Looger LL, Svoboda K, Kim DS. Ultrasensitive fluorescent proteins for imaging neuronal activity. Nature. 2013;499:295–300. doi: 10.1038/nature12354. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Fremeau RT, Troyer MD, Pahner I, Nygaard GO, Tran CH, Reimer RJ, Bellocchio EE, Fortin D, Storm-Mathisen J, Edwards RH. The expression of vesicular glutamate transporters defines two classes of excitatory synapse. Neuron. 2001;31:247–260. doi: 10.1016/S0896-6273(01)00344-0. [DOI] [PubMed] [Google Scholar]
  10. Goel P, Dickman D. Distinct homeostatic modulations stabilize reduced postsynaptic receptivity in response to presynaptic DLK signaling. Nature Communications. 2018;9:1856. doi: 10.1038/s41467-018-04270-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Gorczyca M, Budnik V. Anatomy of the larval body wall muscles and NMJs in the third instar larval stage. The Fly Neuromuscular Junction: Structure and Function. 2006;75:367–373. doi: 10.1016/S0074-7742(06)75016-4. [DOI] [Google Scholar]
  12. Griffiths G, Hoppeler H. Quantitation in immunocytochemistry: correlation of immunogold labeling to absolute number of membrane antigens. Journal of Histochemistry & Cytochemistry. 1986;34:1389–1398. doi: 10.1177/34.11.3534077. [DOI] [PubMed] [Google Scholar]
  13. Hentze JL, Carlsson MA, Kondo S, Nässel DR, Rewitz KF. The neuropeptide allatostatin A regulates metabolism and feeding decisions in Drosophila. Scientific Reports. 2015;5:11680. doi: 10.1038/srep11680. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Hökfelt T, Broberger C, Xu ZQ, Sergeyev V, Ubink R, Diez M. Neuropeptides--an overview. Neuropharmacology. 2000;39:1337–1356. doi: 10.1016/S0028-3908(00)00010-1. [DOI] [PubMed] [Google Scholar]
  15. Hoopfer ED, Jung Y, Inagaki HK, Rubin GM, Anderson DJ. P1 interneurons promote a persistent internal state that enhances inter-male aggression in Drosophila. eLife. 2015;4:e11346. doi: 10.7554/eLife.11346. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Insel TR, Young LJ. Neuropeptides and the evolution of social behavior. Current Opinion in Neurobiology. 2000;10:784–789. doi: 10.1016/S0959-4388(00)00146-X. [DOI] [PubMed] [Google Scholar]
  17. Kempf C, Staudt T, Bingen P, Horstmann H, Engelhardt J, Hell SW, Kuner T. Tissue multicolor STED nanoscopy of presynaptic proteins in the calyx of held. PLOS ONE. 2013;8:e62893. doi: 10.1371/journal.pone.0062893. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Kendrick KM. Microdialysis measurement of in vivo neuropeptide release. Journal of Neuroscience Methods. 1990;34:35–46. doi: 10.1016/0165-0270(90)90040-M. [DOI] [PubMed] [Google Scholar]
  19. Koon AC, Budnik V. Inhibitory control of synaptic and behavioral plasticity by octopaminergic signaling. Journal of Neuroscience. 2012;32:6312–6322. doi: 10.1523/JNEUROSCI.6517-11.2012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Levitan ES, Lanni F, Shakiryanova D. In vivo imaging of vesicle motion and release at the Drosophila neuromuscular junction. Nature Protocols. 2007;2:1117–1125. doi: 10.1038/nprot.2007.142. [DOI] [PubMed] [Google Scholar]
  21. Lin MZ, Schnitzer MJ. Genetically encoded indicators of neuronal activity. Nature Neuroscience. 2016;19:1142–1153. doi: 10.1038/nn.4359. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Littleton JT, Bellen HJ, Perin MS. Expression of synaptotagmin in Drosophila reveals transport and localization of synaptic vesicles to the synapse. Development. 1993;118:1077–1088. doi: 10.1242/dev.118.4.1077. [DOI] [PubMed] [Google Scholar]
  23. Loveall BJ, Deitcher DL. The essential role of bursicon during Drosophila development. BMC Developmental Biology. 2010;10:92. doi: 10.1186/1471-213X-10-92. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. McNabb SL, Truman JW. Light and peptidergic eclosion hormone neurons stimulate a rapid eclosion response that masks circadian emergence in Drosophila. Journal of Experimental Biology. 2008;211:2263–2274. doi: 10.1242/jeb.015818. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Menon KP, Carrillo RA, Zinn K. Development and plasticity of the Drosophila larval neuromuscular junction. Wiley Interdisciplinary Reviews. Developmental Biology. 2013;2:647–670. doi: 10.1002/wdev.108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Miesenböck G, De Angelis DA, Rothman JE. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature. 1998;394:192–195. doi: 10.1038/28190. [DOI] [PubMed] [Google Scholar]
  27. Mitchell KJ, Pinton P, Varadi A, Tacchetti C, Ainscow EK, Pozzan T, Rizzuto R, Rutter GA. Dense core secretory vesicles revealed as a dynamic ca(2+) store in neuroendocrine cells with a vesicle-associated membrane protein aequorin chimaera. The Journal of Cell Biology. 2001;155:41–52. doi: 10.1083/jcb.200103145. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Nässel DR, Winther AM. Drosophila neuropeptides in regulation of physiology and behavior. Progress in Neurobiology. 2010;92:42–104. doi: 10.1016/j.pneurobio.2010.04.010. [DOI] [PubMed] [Google Scholar]
  29. Pfeiffer BD, Ngo TT, Hibbard KL, Murphy C, Jenett A, Truman JW, Rubin GM. Refinement of tools for targeted gene expression in Drosophila. Genetics. 2010;186:735–755. doi: 10.1534/genetics.110.119917. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Rao S, Lang C, Levitan ES, Deitcher DL. Visualization of neuropeptide expression, transport, and exocytosis in Drosophila melanogaster. Journal of Neurobiology. 2001;49:159–172. doi: 10.1002/neu.1072. [DOI] [PubMed] [Google Scholar]
  31. Schaible HG, Jarrott B, Hope PJ, Duggan AW. Release of immunoreactive substance P in the spinal cord during development of acute arthritis in the knee joint of the cat: a study with antibody microprobes. Brain Research. 1990;529:214–223. doi: 10.1016/0006-8993(90)90830-5. [DOI] [PubMed] [Google Scholar]
  32. Shakiryanova D, Tully A, Levitan ES. Activity-dependent synaptic capture of transiting peptidergic vesicles. Nature Neuroscience. 2006;9:896–900. doi: 10.1038/nn1719. [DOI] [PubMed] [Google Scholar]
  33. Sturman DA, Shakiryanova D, Hewes RS, Deitcher DL, Levitan ES. Nearly neutral secretory vesicles in Drosophila nerve terminals. Biophysical Journal. 2006;90:L45–L47. doi: 10.1529/biophysj.106.080978. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Sweeney ST, Broadie K, Keane J, Niemann H, O'Kane CJ. Targeted expression of tetanus toxin light chain in Drosophila specifically eliminates synaptic transmission and causes behavioral defects. Neuron. 1995;14:341–351. doi: 10.1016/0896-6273(95)90290-2. [DOI] [PubMed] [Google Scholar]
  35. Taghert PH, Veenstra JA. Drosophila neuropeptide signaling. Advances in Genetics. 2003;49:1–65. doi: 10.1016/S0065-2660(03)01001-0. [DOI] [PubMed] [Google Scholar]
  36. Tian L, Hires SA, Looger LL. Imaging neuronal activity with genetically encoded calcium indicators. Cold Spring Harbor Protocols. 2012;2012:647–656. doi: 10.1101/pdb.top069609. [DOI] [PubMed] [Google Scholar]
  37. Umezaki Y, Yasuyama K, Nakagoshi H, Tomioka K. Blocking synaptic transmission with tetanus toxin light chain reveals modes of neurotransmission in the PDF-positive circadian clock neurons of Drosophila melanogaster. Journal of Insect Physiology. 2011;57:1290–1299. doi: 10.1016/j.jinsphys.2011.06.004. [DOI] [PubMed] [Google Scholar]
  38. van den Pol AN. Neuropeptide transmission in brain circuits. Neuron. 2012;76:98–115. doi: 10.1016/j.neuron.2012.09.014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Winther AM, Siviter RJ, Isaac RE, Predel R, Nässel DR. Neuronal expression of tachykinin-related peptides and gene transcript during postembryonic development of Drosophila. Journal of Comparative Neurology. 2003;464:180–196. doi: 10.1002/cne.10790. [DOI] [PubMed] [Google Scholar]
  40. Wong MY, Cavolo SL, Levitan ES. Synaptic neuropeptide release by dynamin-dependent partial release from circulating vesicles. Molecular Biology of the Cell. 2015;26:2466–2474. doi: 10.1091/mbc.E15-01-0002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Xia X, Lessmann V, Martin TF. Imaging of evoked dense-core-vesicle exocytosis in hippocampal neurons reveals long latencies and kiss-and-run fusion events. Journal of Cell Science. 2009;122:75–82. doi: 10.1242/jcs.034603. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Xu T, Binz T, Niemann H, Neher E. Multiple kinetic components of exocytosis distinguished by neurotoxin sensitivity. Nature Neuroscience. 1998;1:192–200. doi: 10.1038/642. [DOI] [PubMed] [Google Scholar]
  43. Xu T, Xu P. Differential Regulation of Small Clear Vesicles and Large Dense-Core Vesicles. In: Wang Z. W, editor. Molecular Mechanisms of Neurotransmitter Release. Totowa, NJ: Humana Press; 2008. pp. 327–399. [Google Scholar]
  44. Zandawala M, Yurgel ME, Texada MJ, Liao S, Rewitz KF, Keene AC, Nässel DR. Modulation of Drosophila post-feeding physiology and behavior by the neuropeptide leucokinin. PLOS Genetics. 2018;14:e1007767. doi: 10.1371/journal.pgen.1007767. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Zhu D, Zhou W, Liang T, Yang F, Zhang RY, Wu ZX, Xu T. Synaptotagmin I and IX function redundantly in controlling fusion pore of large dense core vesicles. Biochemical and Biophysical Research Communications. 2007;361:922–927. doi: 10.1016/j.bbrc.2007.07.083. [DOI] [PubMed] [Google Scholar]

Decision letter

Editor: Leslie C Griffith1
Reviewed by: Leslie C Griffith2, Paul Taghert3

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Thank you for submitting your article "Imaging neuropeptide release at synapses with a genetically engineered reporter" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Leslie C Griffith as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Eve Marder as the Senior Editor. The following individual involved in review of your submission has also agreed to reveal their identity: Paul Taghert (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This paper develops a new set of tools for looking at release of peptides in real time. The idea is a good one and there is a need in the field for such tools. The authors describe a genetically encoded reporter system for dense core vesicle (DCV) fusion in Drosophila. They generate a host of fusions of GCaMP6 to ANF and DTK neuropeptides and identify a specific fusion event that localizes well to DCVs by both immunocytochemistry and immuno-EM. The authors show the sensor can respond to strong stimulation and compare the response to cytosolic GCaMP6 as a read-out of intra-terminal calcium changes. Overall, it's a potentially useful tool for the field, but the study could add a bit more quantification to demonstrate how well the sensor is actually working.

Essential revisions:

1) The major issue the reviewers had with the feasibility of this approach for dissecting DCV release is the conditions needed to trigger these changes – 70 Hz stimulation for 18 secs. These motor neurons would never experience such extreme stimulation in vivo. Peptidergic neurons typically fire in the 1-5 Hz range and while motor neurons fire at higher rates, natural stimulation is not of the duration used here. The signal-to-noise ratio (δ F changes) of this DCV sensor (compared to GCaMP alone which appears to be at least 5-10x more sensitive) is also rather poor. As such, it would be informative to have more information on the dynamics of their new reporter.

a) Provide a dose response curve to demonstrate both the required stimulation frequency (only 70 Hz is reported), the required length of stimulation (only 18 seconds is reported) and the calcium response curve. What does 1 Hz look like? Or 5 Hz?

b) The authors only report changes in the Type 3 terminals. What did the authors see in the Type 1 terminals that have fewer DCVs? Was DCV release detectable in these neurons, as has been reported in the field for the classical ANF-GFP DCV sensor? These data may be more useful than Type III data for investigators contemplating using this tool in neurons that do not have huge numbers of DCVs.

c) It would be helpful to use a benchmark to compare the methodology described here to that currently used in the field – the loss of ANF-GFP signal following stimulation. Is the current system actually an upgrade over that? There is unique value here in the potential to endogenously target specific neuropeptides for this technique, but if the sensitivity is far below that of ANF-GFP, the older system might still be the one of choice for dissecting the biology of DCV fusion overall. Without these data it is hard to claim an advance.

2) It would be helpful for the authors to further clarify in discussion what they actually think the sensor is reporting over time. It was not clear to reviewers exactly what the rise/falling/undershoot/recovery phases actually represent. The recovery phase is especially mysterious since the soma is gone so it cannot be replenishment via transport. DCVs are known to only release part of their cargo during stimulation, so presumably many of these DCVs still contain GCaMP-tether neuropeptide within them. Do the DCVs that have fused become less acidic and take up come extracellular calcium and thus activate the remaining GCaMP within them? Or do the authors think that the DCVs release all their content, or not take up calcium or lose their acidity during partial fusion events? This seems a potential confound – the authors are measuring a transient rise in calcium influx and pH change within the lumen of the DCV, not the actual release of the neuropeptide-GCaMP.

While the reviewers did not feel that it was required that this point be addressed with new experiments, it would be incredibly informative to assay their sensor in animals concurrently expressing an ANF-mCherry, ANF-mOrange (or other red-shifted tagged neuropeptide) where loss of the neuropeptide is directly reported in addition to the neuropeptide-GCaMP signal. That would really help clarify what is likely to be happening with their sensor during these phases of DCV release or partial fusion.

eLife. 2019 Jun 26;8:e46421. doi: 10.7554/eLife.46421.sa2

Author response

Essential revisions:

1) The major issue the reviewers had with the feasibility of this approach for dissecting DCV release is the conditions needed to trigger these changes – 70 Hz stimulation for 18 secs. These motor neurons would never experience such extreme stimulation in vivo. Peptidergic neurons typically fire in the 1-5 Hz range and while motor neurons fire at higher rates, natural stimulation is not of the duration used here. The signal-to-noise ratio (δ F changes) of this DCV sensor (compared to GCaMP alone which appears to be at least 5-10x more sensitive) is also rather poor. As such, it would be informative to have more information on the dynamics of their new reporter.

a) Provide a dose response curve to demonstrate both the required stimulation frequency (only 70 Hz is reported), the required length of stimulation (only 18 seconds is reported) and the calcium response curve. What does 1 Hz look like? Or 5 Hz?

b) The authors only report changes in the Type 3 terminals. What did the authors see in the Type 1 terminals that have fewer DCVs? Was DCV release detectable in these neurons, as has been reported in the field for the classical ANF-GFP DCV sensor? These data may be more useful than Type III data for investigators contemplating using this tool in neurons that do not have huge numbers of DCVs.

We share the reviewers’ concern. In response to points 1a and 1b, we have added a new Figure 4, where we present stimulation frequency titrations of NPRRANP, NPRRdTK and GCaMP6 responses in both Type III neurons and Type Ib neurons. Because 70 mM potassium or 70 Hz electric stimulation were reported necessary for inducing maximum dense core vesicle release events in Drosophila type III motor neurons 1, we covered stimulation frequencies from 1 Hz to 70 Hz as the reviewer requested (1a). To compare and contrast the results between preparations and genotypes, for each preparation we normalized the ∆F/F peaks to responses obtained at 70Hz. We did not change the duration of stimulation, as it adds an additional dimension of complexity to the experiments.

Type III motor neurons contain primarily DCVs and exhibit a distinct morphology compared to type Ib motor neurons. As reviewer requested (1b), we have applied the same electric stimulation series to the glutamatergic type Ib motor neurons, which contain both SVs and DCVs 2.

To answer the reviewers’ questions in reverse order:

1b) NPRR responses can be observed in Type Ib as well as in Type III neurons (new Figure 4).

1a) In Type Ib neurons, no statistically significant NPRR responses could be observed at 1 Hz (NPRRANP: P=0.2817; NPRRdTK: P=0.0723) or 5 Hz (NPRRANP: P=0.0727; NPRRdTK: P=0.3467). However, responses at 10 Hz were statistically significant (NPRRANP: P=0.0108; NPRRdTK: P=0.0124). GCaMP responses were statistically significant at 5 Hz (P=0.0152) but not at 1 Hz (P=0.1194). Importantly, unlike the case in Type III neurons, we did not observe significant differences in the performance of the NPRRANP and NPRRdTK reporters in Type Ib neurons (Figure 4—figure supplement 1). In Type III neurons, no statistically significant responses were observed at 1 Hz (NPRRANP: P=0.3304; NPRRdTK: P=0.0719), 5 Hz (NPRRANP: P=0.3699; NPRRdTK: P=0.3467) and 10 Hz (NPRRANP: P=0.3084; NPRRdTK: P=0.5660). We observed a statistically significant increase in ∆F/F at 30 Hz for NPRRANP but not NPRRdTK; at 50 Hz and above we observed significant responses with both reporters. (NPRRANP: P=0.0021; NPRRdTK: P=0.034). The difference between both NPRRs at 30 Hz or 50 Hz was also statistically significant ((Figure 4—figure supplement 1).

(*Statistical methods: For in-sample comparison, one-sample T-test was used to compare response at a given stimulation frequency to 0. For between-sample comparison at the same stimulation frequency, Mann-Whitney U test was used.)

These results reveal two important new findings: (1) the difference between the performance of the two NPRRs we tested in Type III neurons is not a general property of these reporters, but a peculiarity of Type III neurons; (2) the high threshold for detection of NPRR responses in Type III neurons (30-50 Hz) is also not a general property of these reporters, since significant responses can be observed at frequencies as low as 10 Hz in Type 1b neurons. 10 Hz is well within the range of physiological rates for many neurons. These observations should allay concerns that NPRRs will have limited applicability.

Indeed, these new results reveal a novel application for NPRRs: they can be used to compare the relationship between stimulation and neuropeptide release between different types of neurons. In the present case, the difference in NPRR release detection thresholds between Type Ib and Type III neurons appears to be correlated with differences in the overall responses of these two cell types to electrical stimulation, as revealed by conventional GcaMP imaging (Figure 4C, F). However further studies will be necessary to substantiate this. To our knowledge, such a comparison of NP release thresholds between different neuronal cell types has not been performed previously. These data therefore illustrate a new area of investigation that is opened up by this technology.

In response to the reviewer’s comment that the signal-to-noise ratio of this sensor is rather poor, in comparison to that of GCaMP6, we would respectfully point out that the signal-to-noise ratio of the first-generation GCaMPs were also rather poor (max ∆F/F ≪100%). It took a managed team of protein engineers at Janelia 5 years and $10 million to optimize GCaMP from version 1.3, to version 6 (the first version with single-spike sensitivity). We would respectfully submit that the expectation that a first-generation reporter of a new class would perform as well as the 6th generation GcaMP is unrealistic. We anticipate that there will be multiple iterations of NPRRs to optimize their performance, but such optimization is beyond the scope of this manuscript.

c) It would be helpful to use a benchmark to compare the methodology described here to that currently used in the field – the loss of ANF-GFP signal following stimulation. Is the current system actually an upgrade over that? There is unique value here in the potential to endogenously target specific neuropeptides for this technique, but if the sensitivity is far below that of ANF-GFP, the older system might still be the one of choice for dissecting the biology of DCV fusion overall. Without these data it is hard to claim an advance.

We took the reviewers’ point and attempted to benchmark our method to ANP-GFP. While both methods showed comparable decreases in fluorescence in response to 70 mM KCl (see below), in response to electrical stimulation, we failed to observe any decrease in ANP-GFP signals in Type Ib or Type III neurons (Author response image 1A1,2). In contrast, under the same stimulation conditions NPRRANP clearly exhibited an increase in dF/F (Author response image 1A3,4). In other words, in our hands the ANPGFP method yielded no signal (decreased fluorescence) in response to electrical stimulation, whereas the NPRR clearly did so. Therefore rather than it being the case that the sensitivity of NPRRs is “far below that of ANF-GFP,” it appears that NPRRs are far more sensitive than ANF-GFP, in the context of electrical stimulation.

Author response image 1. Benchmarking NPRR with ANP-GFP.

Author response image 1.

Open in a new tab

Trace from a representative experiment showing changes in fluorescence intensity (∆F/F) of ANP-GFP and NPRRANP evoked by electrical stimulation (Red bar, frequency shown in A1). No significant ANP-GFP fluorescent change was observed within stimulations, in both Type Ib (A1) and Type III neurons (A2), while NPRRANP displays distinct characteristics in Type Ib (A3) and Type III(A4). Green line: mean ∆F/F; Grey shading: s.e.m envelope.

Details: We measured the native fluorescent intensity of ANP-GFP or of NPRRANP directly in fixed specimens following stimulation with 70 mM KCl. In this case, we observed a significant decrease in ANP-GFP fluorescence (36.04% decrease, N=7) after 10 mins, consistent with published work from the Levitan laboratory 1,3. In parallel, NPRRANP underwent a comparable level of fluorescent loss (33.57% decrease, N=7), suggesting a similar sensitivity of the two reporters under these conditions of bath depolarization.

We cannot explain our failure to observe any fluorescent loss of ANP-GFP in response to tenable electrical stimulation, in contrast to published results from the Levitan laboratory 1,3. Whatever the explanation, our results demonstrate the superiority of the NPRR methodology. In addition to being able to visualize specific neuropeptides, the ultimate objective of generating NPRRs is to be able to measure NP release in behaving animals over time. For this purpose, a positive signal would be more optimal than a negative signal (ANP-GFP), as fluorescent loss could be confounded by other technical issues, such as photobleaching or movement.

2) It would be helpful for the authors to further clarify in discussion what they actually think the sensor is reporting over time. It was not clear to reviewers exactly what the rise/falling/undershoot/recovery phases actually represent. The recovery phase is especially mysterious since the soma is gone so it cannot be replenishment via transport. DCVs are known to only release part of their cargo during stimulation, so presumably many of these DCVs still contain GCaMP-tether neuropeptide within them. Do the DCVs that have fused become less acidic and take up come extracellular calcium and thus activate the remaining GCaMP within them? Or do the authors think that the DCVs release all their content, or not take up calcium or lose their acidity during partial fusion events? This seems a potential confound – the authors are measuring a transient rise in calcium influx and pH change within the lumen of the DCV, not the actual release of the neuropeptide-GCaMP.

We appreciate the reviewers’ request to clarify the temporal kinetics we observed. As we stated in the original submission, we interpret the rising phase as reflecting epochs of NP release. The falling phase where an undershoot was observed suggests that the overall decrease of GCaMP fluorescence represents a reduction of total DCV NPRR cargo in the bouton area following vesicle fusion.

The reviewers state that the recovery phase is mysterious, because replenishment via transport cannot occur in the absence of the soma, which is removed in these preparations. The reviewer’s statement is partially incorrect: recent studies have demonstrated that axonal transport indeed occurs during short-term presynaptic protein remodelling with4 or without5 the ventral verve cord (i.e., somata) removed, over a time course ranging from 10 – 30 min 6,7, a timeframe similar to that in our experimental design. Thus, replenishment of DCVs via continuous transport from unstimulated axon segments contained within our preparation seems the most parsimonious explanation for the recovery phase.

DCV release occurs in several modes. At this point, we are unable to determine whether an individual DCV unloads all or a portion of its content in our experiments, although as mentioned above (response to point 1c) we detected an ~36% decrease in overall GCaMP fluorescence following high potassium stimulation. Furthermore, the ∆F/F undershoot observed in the recovery phase supports the idea that at least some, if not all, NPRR molecules are released to the extracellular space, since a purely intravesicular change in calcium/pH would not cause such an undershoot. Nevertheless, we cannot fully exclude the possibility that some fraction of NPRR increased ∆F/F signal is contributed by unreleased NPRR molecules, and now acknowledge this in the Discussion (second paragraph).

While the reviewers did not feel that it was required that this point be addressed with new experiments, it would be incredibly informative to assay their sensor in animals concurrently expressing an ANF-mCherry, ANF-mOrange (or other red-shifted tagged neuropeptide) where loss of the neuropeptide is directly reported in addition to the neuropeptide-GCaMP signal. That would really help clarify what is likely to be happening with their sensor during these phases of DCV release or partial fusion.

We agree with the reviewers’ suggestion to have an additional channel to report neuropeptide loss directly. In fact, we initially generated sixteen different neuropeptide precursor-Red Fluorescent Protein fusions (Figure 1—figure supplement 1). Unfortunately, all 16 red NPRRs exhibited 1) nearly invisible baseline fluorescence; and 2) no fluorescence change following high potassium challenge (unlike GFP fusions). We tried to investigate the reason for this failure using Neuropred (http://stagbeetle.animal.uiuc.edu/cgi-bin/neuropred.py), an online algorithm that predicts potential peptidergic cleavage sites within a given protein. This analysis revealed that RFP and tdTomato have 10 and 25 predicted cleavage sites, respectively, as opposed to 3 sites in GCaMP6s, mainly reflecting the high abundance of positively-charged residue repeats (Lys-Lys, Lys-Arg) in RFPs. If these cleavage sites are accessible to DCV proteases that processes packaged NP precursors, it would explain why no red fluorescence was observed with these reporters. We have now mentioned this in the text. Apparently, generating such RFP-based reporters would require mutagenizing all such sites to avoid cleavage, while retaining red fluorescence. Such protein engineering is desirable, but outside the scope of the present manuscript.

We appreciate the suggestion to use orange fluorescent proteins such as mOrange or mKO as a fusion alternative. Although it was not included in our first round of design, we would be glad to try it in the future.

References:

1) Levitan, E. S., Lanni, F. and Shakiryanova, D. In vivo imaging of vesicle motion and release at the Drosophila neuromuscular junction. Nat Protoc 2, 1117-1125, doi:10.1038/nprot.2007.142 (2007).

2) Rao, S., Lang, C., Levitan, E. S. and Deitcher, D. L. Visualization of neuropeptide expression, transport, and exocytosis in Drosophila melanogaster. J Neurobiol 49, 159-172 (2001).

3) Shakiryanova, D., Tully, A. and Levitan, E. S. Activity-dependent synaptic capture of transiting peptidergic vesicles. Nat Neurosci 9, 896-900, doi:10.1038/nn1719 (2006).

4) Goel, P. et al. Homeostatic scaling of active zone scaffolds maintains global synaptic strength. J Cell Biol 218, 1706-1724, doi:10.1083/jcb.201807165 (2019).

5) Bohme, M. A. et al. Rapid active zone remodeling consolidates presynaptic potentiation. Nat Commun 10, 1085, doi:10.1038/s41467-019-08977-6 (2019).

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Supplementary Materials

    Figure 1—source data 1. Raw data for Immuno-EM experiments (ANP).
    elife-46421-fig1-data1.zip (95MB, zip)
    Figure 1—source data 2. Raw data for Immunno-EM experiments (control group).
    elife-46421-fig1-data2.zip (26.7MB, zip)
    Figure 3—source data 1. Raw data for Immuno-EM experiments (dTK).
    elife-46421-fig3-data1.zip (49.3MB, zip)
    Source code 1. Code for data analysis.
    elife-46421-code1.zip (1.8KB, zip)
    Supplementary file 1. Current techniques for neuropeptide release measurement.

    Summary of current techniques used for neuropeptide release, including microdialysis, antibody-coated microprobes, GFP-tagged propeptide imaging and NPRR. NA, ‘not applicable’

    elife-46421-supp1.xlsx (10.6KB, xlsx)
    Supplementary file 2. Stereological labeling estimates.

    Stereological labeling estimations of NPRRANP-GFP and NPRRdTK-GFP, respectively, in Type Ib neurons, or in Type Ib and Type III neurons. Biological controls and internal controls are described in Materials and methods. SNR: Signal-to-Noise Ratio.

    elife-46421-supp2.docx (15.9KB, docx)
    Supplementary file 3. Complete genotypes of transgenic flies used in this study.

    Summary of complete genotypes of transgenic flies used in this study.

    elife-46421-supp3.docx (17.4KB, docx)
    Transparent reporting form
    elife-46421-transrepform.pdf (203KB, pdf)

    Data Availability Statement

    Source data of EM for Figure 1 and 3, and codes used for Figure 2 and 3 have been provided.

    Articles from eLife are provided here courtesy of eLife Sciences Publications, Ltd

    RESOURCES