Synaptic structure and activity are sensitive to environmental alterations. Modulation of synaptic morphology and function is often induced by signals from glia. However, the process by which glia mediate synaptic responses to environmental perturbations such as hypoxia remains unknown. Here, we report that, in the mutant for Trachealess (Trh), the Drosophila homolog for NPAS1 and NPAS3, smaller synaptic boutons form clusters named satellite boutons appear at larval neuromuscular junctions (NMJs), which is induced by the reduction of internal oxygen levels due to defective tracheal branches. Thus, the satellite bouton phenotype in the trh mutant is suppressed by hyperoxia, and recapitulated in wild-type larvae raised under hypoxia. We further show that hypoxia-inducible factor (HIF) -1α/Similar (Sima) is critical in mediating hypoxia-induced satellite bouton formation. Sima upregulates the level of the Wnt/Wingless (Wg) signal in glia, leading to reorganized microtubule structures within presynaptic sites. Finally, hypoxia-induced satellite boutons maintain normal synaptic transmission at the NMJs, which is crucial for coordinated larval locomotion.
Animals need oxygen and food, not only to sustain life, but also for motility. In vertebrates, oxygen and nutrients are delivered through the vascular systems to organs and tissues throughout the body. To maintain proper nutrient and oxygen supply, and thus physiological functions, the vascular system is also highly coordinated with the nervous system during development. Indeed, the vascular and nervous systems resemble each other in terms of their anatomical structures and developmental processes [1]. In the brain, nerves and vessels, form close associations and are in physical contact through the third player astrocytes to form neurovascular units (NVU) [2]. Such organization is essential for controlling oxygen and glucose delivery through the blood vessels by neuronal activity, and this regulatory process is mediated through the coupled astrocytes [3]. However, some invertebrates lack the complex vascular systems [4]. In nematodes, oxygen is supplied simply by ambient diffusion to inner cells [5]. Insects such as Drosophila have evolved a prototype of the tracheal system to deliver oxygen and a primitive vascular system, the dorsal vessel, to facilitate nutrient delivery [6]. However, the physical association of nerves, trachea, and glial processes has also been demonstrated at the NMJs of adult Drosophila flight muscles [7]. Animals respond to changing oxygen levels by altering their oxygen delivery system. Insufficient oxygen levels (hypoxia) activate a broad range of genes to re-establish body homeostasis. One crucial regulator of these hypoxia-responsive genes is the sequence-specific DNA-binding transcription factor hypoxia inducible factor 1 (HIF-1) [8]. HIF-1 consists of α and β subunits that form heterodimers [9]. Whereas HIF-1β is expressed constitutively, HIF-1α protein levels are modulated by oxygen levels [10]. Under normal oxygen conditions (normoxia), oxygen-dependent prolyl hydroxylases (PHDs) catalyze hydroxylation of a conserved prolyl residue in the central oxygen-dependent degradation (ODD) domain of HIF-1α [11]. Hydroxylation of HIF-1α promotes interaction with Von Hippel Lindau (VHL), which is the substrate recognition subunit of the cullin2-based E3 ubiquitin ligase, leading to HIF-1α ubiquitination and proteasomal degradation. Under hypoxia, prolyl hydroxylation does not occur, HIF-1α proteins are stabilized and are translocated from the cytoplasm to the nucleus where they form heterodimers with HIF-1β to activate transcription of target genes. One major class of target genes encoding the Fibroblast Growth Factor (FGF) is involved in inducing angiogenesis in mammals. In Drosophila, the FGF member encoded by Branchless (Bnl) induces tracheal branching [12]. When oxygen levels are reduced, oxygen-starved cells express Bnl as a chemo-attractant to guide the growth tracheal terminal branches toward them [13]. In addition to adaptations of the respiratory system, the nervous system also responds to hypoxia. Oxygen levels modulate the survival, proliferation, and differentiation of radial glial cells (RGCs) in the human cerebral cortex. Interestingly, physiological hypoxia (3% O2) induces neurogenesis and differentiation of RGCs into glutamatergic neurons [14]. Hypoxia induces neurite outgrowth in PC12 cells through activation of A2A receptor [15]. Brief exposure to anoxia and hypoglycemia caused axonal remodeling in hippocampal neurons, including presynaptic protrusion of filopodia and formation of multi-innervated spines [16]. Under hypoxia or upon depletion of PHD2, upregulation of the actin cross-linker Filamin A (FLNA) induces generation of more immature spines [17]. Astrocytes have been shown to play a crucial role in ischemic tolerance via the activation of P2X7 receptors, which trigger upregulation of HIF-1α [18]. Neuronal PAS (NPAS) proteins containing a DNA-binding Per-Arnt-Sim domain function in vascular and nervous system development. In mice, NPAS1 is responsible for cortical interneuron generation [19], whereas NPAS3 is required for adult neurogenesis [20]. NPAS1 and NPAS3 also play key roles in lung development [21,22]. The homolog of NPAS1/3 in Drosophila, Trachealess (Trh), has been well studied for its involvement in formation of the respiratory tracheal system. Trh is a master regulator of tracheal cell fates, activating gene expression to induce tracheal development [23,24]. However, the role of Trh in the development of other tissues, particularly the nervous system, is unknown. In this study, we found altered synaptic bouton morphology at the NMJs of trh1/trh2 mutants. By performing trh-RNAi knockdown and UAS-trh transgene rescue experiments, we show that trh is required in tracheal cells for normal bouton formation. Defective tracheal branching in the trh1/trh2 mutant mimics the effect of hypoxic conditions during larval development, and supplying higher than normal oxygen levels restored normal bouton morphology. We further show that glial cells respond to hypoxia by elevating Wnt/Wg expression to mediate synaptic bouton reorganization through HIF1-α/Sima in Drosophila. Finally, we reveal that this morphological change may be linked to normal synaptic transmission and locomotion of larvae.
To understand the possible role of Trh in synapse formation, we examined NMJ morphology in the trh mutant. Since both trh1 and trh2 loss-of-function alleles are homozygous lethal [23,25,26], we examined the trans-heterozygous trh1/trh2 mutant that survived to adult stages and compared it to wild-type (w1118) and heterozygous trh1/+ controls. Synaptic boutons of w1118 and trh1/+ NMJs were evenly spaced along the axonal terminals, displaying the typical “beads-on-a-string” pattern (Fig 1A, upper and middle panels, enlarged images at right). Strikingly, the trh1/trh2 mutant larvae exhibited aberrant NMJ morphology. Small synaptic boutons formed clusters or surrounded a normal large bouton (Fig 1A, bottom panel); a phenotype described as “satellite boutons” [27]. This satellite bouton phenotype in the trh1/trh2 mutant was detected at a high frequency; 20. 7 ± 4. 1% (n = 10) of total boutons were satellite ones, more than three-fold increases compared to 4. 3 ± 1. 7% (n = 10) for trh1/+ and 6. 3 ± 1. 7% (n = 10) for w1118 (Fig 1B). Although the percentage of satellite boutons in the trh1/trh2 mutant was greatly increased, the total bouton number was slightly higher than that observed in controls (74. 1 ± 4. 7, n = 10 in w1118; 89. 6 ± 5. 2, n = 10 in trh1/+; and 95. 5 ± 9. 1, n = 11 in trh1/trh2, bottom panel in Fig 1B). Also, the muscle areas were not significantly different from each other (S1A Fig). Given the small size and clustering of satellite boutons in the trh1/trh2 mutant, we examined whether these boutons express synaptic proteins normally. We found that the synaptic vesicle protein Synapsin (Syn in Fig 1A) was normally distributed relative to control, but the active zone protein Bruchpilot (Brp) was expressed at higher levels in the trh1/trh2 mutant (S1B Fig). The postsynaptic glutamate receptor, as revealed by GluRIII (S1B Fig) and GluRIIA (S1C Fig) signals, as well as dPAK (S1C Fig) were also localized in satellite boutons, which were surrounded by the subsynaptic reticulum protein Dlg (S1D Fig). Thus, although the Brp signal intensity in the trh1/trh2 mutant was stronger than in controls, the composition of synaptic proteins in satellite boutons was largely similar to that of normal-sized boutons. As trh is expressed in both tracheal and nervous systems in embryonic stages [28], altered bouton morphology in the trh1/trh2 mutant could be due to a lack of trh in neurons, tracheal cells or other cells/tissues. Therefore, we performed trh-RNAi knockdown by using tissue-specific GAL4 drivers for trachea (btl-GAL4), neurons (elav-GAL4), glia (repo-GAL4), and muscles (MHC-GAL4). We observed a dramatic increase in satellite boutons upon tracheal trh-RNAi knockdown using btl-GAL4 (10. 2 ± 1. 7%, n = 10) compared to those in the trh-RNAi (2. 7 ± 1. 3%, n = 10) or btl-GAL4 (3. 3 ± 0. 9%, n = 12) control (Fig 1C and 1D. In contrast, trh-RNAi knockdown by other tissue-specific GAL4 drivers failed to significantly increase satellite boutons, displaying similar amounts of satellite boutons to respective GAL4 drivers (2. 0 ± 0. 7%, n = 10 in elav-GAL4 v. s. 1. 4 ± 0. 6%, n = 10 in elav-GAL4/trh-RNAi; 2. 8 ± 1. 0%, n = 10 in repo-GAL4 v. s. 3. 4 ± 0. 9%, n = 10 in repo-GAL4/trh-RNAi; and 1. 3 ± 0. 8%, n = 10 in MHC-GAL4 v. s. 2. 1 ± 1. 0%, n = 10 in MHC-GAL4/trh-RNAi, Fig 1D). To further confirm the necessity of tracheal trh for normal bouton formation, we performed a rescue experiment for the trh1/trh2 phenotype. When UAS-trh expression was driven by tracheal btl-GAL4 in the trh1/trh2 mutant, the satellite bouton phenotype was suppressed (1. 8 ± 1. 4%, n = 13, Fig 1E and 1F). Controls bearing only btl-GAL4 (14. 3 ± 3. 3%, n = 10) or UAS-trh (21. 1 ± 3. 3%, n = 10) still contained large amounts of satellite boutons. These results indicate that trh is required in the trachea for normal bouton formation. Apart from specifying the tracheal cell fate, Trh is also involved in the branching of tubular structures during post-embryonic stages [24]. Therefore, we examined the tracheal phenotypes in the trh1/trh2 larvae (S2A Fig) and observed an increase in the number of terminal branches in the dorsal branch of the third segment (S2B Fig, 5. 7 ± 0. 15, n = 10 for trh1/+, and 7. 5 ± 0. 28, n = 11 for trh1/trh2). Furthermore, we identified morphological defects such as tracheal breaks and tangles, suggesting structural defects in the trh1/trh2 larvae (arrows in S2A Fig). Tracheal branching activity is enhanced under hypoxia [12]. Thus, the increased terminal branches in trh1/trh2 could be a compensatory mechanism for defective trachea formation. To understand whether trh1/trh2 mutant cells are under hypoxia, we used the hypoxia biosensor GFP-ODD, in which the GFP is fused to the ODD domain of Sima, under the control of the ubiquitin-69E (ubi) promoter [29]. We first confirmed that GFP-ODD signal was low under normoxia (21% O2) and enhanced under hypoxia (5% O2) in wild-type late-stage embryos when tracheal tubules are already formed and functioning [29]. Indeed, enhanced GFP signal was ubiquitous under hypoxia in wild-type embryos with some pronounced focal GFP signals (Fig 2A, upper row, and S2C Fig). The signal of mRFP-nls, also under the control of the ubi promoter as an internal control, remained constant under hypoxia (Fig 2A, bottom row, and S2D Fig). Quantification of the GFP/RFP ratio revealed a significant difference between normoxia and hypoxia (0. 18 ± 0. 05, n = 6 at 21% O2 and 0. 84 ± 0. 15, n = 6 at 5% O2, Fig 2B). We then examined whether oxygen supply is deficient in the trh1/trh2 mutant by measuring the GFP/RFP ratios. We detected a higher GFP/RFP ratio (0. 82 ± 0. 15, n = 6) in the mutant compared to that in heterozygous trh1/+ (0. 09 ± 0. 02, n = 6) in the 21% O2 condition, supporting that the trh1/trh2 mutant senses reduced oxygen levels internally (Fig 2A and 2B). Thus, formation of satellite boutons in the trh1/trh2 mutant could be caused by hypoxia. To test this hypothesis, we reared wild-type larvae under hypoxia (5%) and assessed synaptic bouton morphology at the third instar stage (Fig 2C). Consistently, small clustered boutons increased, mirroring the satellite bouton phenotype (Fig 2D, 4. 6 ± 1. 5%, n = 10 in 21% O2 v. s. 19. 4 ± 1. 2%, n = 11 in 5% O2). Furthermore, when the trh1/trh2 mutant was raised in a high oxygen level (50% O2), normal bouton morphology was restored (Fig 2C); the satellite bouton phenotype was almost completely suppressed (Fig 2D, 15. 2 ± 2. 5%, n = 10 in 21% O2 v. s. 0. 74 ± 0. 49%, n = 10 in 50% O2). These results suggest that hypoxia due to the defective tracheal system in the trh1/trh2 mutant induces the satellite bouton phenotype, and that this phenotype can be suppressed by extra oxygen supply. HIF-1α/Sima mediates the response to low oxygen supply [30,31]. The protein levels of Sima are increased in wild-type Drosophila embryos subjected to hypoxia [32], leading to transcriptional activation of downstream target genes and the induction of tracheal branching [12]. We overexpressed Sima in tracheal cells, neurons, glia, or muscle cells by tissue-specific drivers to investigate which types of cells may play a role in modulating synaptic bouton formation. Overexpressing Sima in trachea caused embryonic lethality, preventing us from observing NMJ phenotypes. Larvae in which Sima was overexpressed in muscles, neurons, or glia could survive to the third instar stage, allowing us to examine the bouton phenotypes. We found that Sima overexpression in glia gave the highest number of satellite boutons (Fig 3A, and 11. 9 ± 2. 3%, n = 10 for repo-GAL4/UAS-sima in Fig 3B, v. s. 2. 8 ± 1. 0%, n = 10 for repo-GAL4 in Fig 1D). However, Sima overexpression in neurons by elav-GAL4 failed to induce satellite boutons (2. 6 ± 0. 8%, n = 8 in elav-GAL4/UAS-sima in Fig 3B v. s. 2. 0 ± 0. 7%, n = 10 for elav-GAL4 in Fig 1D). Also, muscle expression of Sima also maintained a basal level of satellite boutons (1. 7 ± 1. 0%, n = 10 for MHC-GAL4/UAS-sima in Fig 3B v. s. 1. 3 ± 0. 8%, n = 10 for MHC-GAL4 in Fig 1D). This result shows that the hypoxia-responding factor Sima is capable of inducing satellite bouton formation when overexpressed in glia. The Drosophila PHD protein Fatiga (Fga) promotes prolyl hydroxylation and degradation of Sima in normaxia [33]. Consistently, glial knockdown by expressing fga-RNAi led to a significant increase of satellite boutons (S3A and S3B Fig, 8. 3 ± 1. 3%, n = 10 for repo-GAL4/fga-RNAi) from the control (4. 3 ± 0. 7%, n = 10 for fga-RNAi). Instead, tracheal knockdown by btl-GAL4 (3. 8 ± 0. 7%, n = 9 for btl-GAL4/fga-RNAi) showed no apparent difference to the control. This result is consistent with that Sima upregulation in glia induces satellite bouton formation. If glial Sima is the factor responding to hypoxia in the trh1/trh2 mutant, reducing Sima level in glia would suppress satellite bouton formation. Accordingly, we expressed the sima-RNAi transgene, which could reduce sima expression in both transcript and protein levels (S3C and S3D Fig), using repo-GAL4 in the trh1/trh2 mutant. As our prediction, the satellite bouton phenotype was suppressed upon glial sima knockdown (3. 6 ± 1. 0%, n = 10 in repo-GAL4/sima-RNAi trh1/trh2), as compared to controls of the trh1/trh2 mutant carrying either the UAS-sima-RNAi transgene (9. 2 ± 1. 5%, n = 10 in sima-RNAi trh1/trh2) or the repo-GAL4 driver (12. 2 ± 2. 2%, n = 10 in repo-GAL4 trh1/trh2) that displayed high percentages of satellite boutons (Fig 3C and 3D). We also tested whether low oxygen level-induced satellite bouton formation is mediated through Sima in glia. Satellite bouton phenotypes were detected in controls carrying either UAS-sima-RNAi (12. 5 ± 2. 7%, n = 11) or repo-GAL4 (16. 4 ± 2. 2%, n = 10) when raised in 5% O2. However, almost no satellite boutons were detected in larvae carrying both repo-GAL4 and UAS-sima-RNAi (0. 45 ± 0. 45%, n = 11 in repo-GAL4/sima-RNAi) when raised in the same condition (Fig 3E and 3F). We also examined whether the Sima protein levels are changed in hypoxia or in the trh mutant. We found ubiquitous increases in the Sima levels in the wild-type control under the 5% O2 condition or in the trh1/trh2 mutant (S3E and S3F Fig). The increases could be identified in glial processes along the peripheral nerves and in different subtypes of glia. Thus, glial Sima could play the role to mediate hypoxia in the trh1/trh2 mutant and in the low O2 condition to modulate synaptic bouton formation. Next, we explored possible signals transduced from glia to neurons in response to hypoxia. The glia-secreted Wingless (Wg) signaling molecule regulates synaptic growth at Drosophila NMJs [34,35]. Therefore, we examined whether Wg can be induced under hypoxia in the trh1/trh2 mutant. Wg signals were enriched around the synaptic boutons of wild-type NMJs (Fig 4A). Whereas the pattern of Wg signals at trh1/+ NMJs was similar to that of w1118, we detected much higher levels of Wg signals at the trh1/trh2 NMJ (Fig 4A). Quantification of Wg immunofluorescence intensities normalized to co-stained HRP in trh1/trh2 (Wg/HRP: 0. 49 ± 0. 07, n = 9, Fig 4B) revealed ~3-fold increases relative to w1118 (0. 18 ± 0. 03, n = 8) and trh1/+ (0. 15 ± 0. 02, n = 9). We then examined whether glial Sima is required for the enhanced Wg expression in the trh1/trh2 mutant (Fig 4C). The Wg level relative to the HRP level in the trh1/trh2 mutant carrying repo-GAL4 (0. 68 ± 0. 16, n = 8 for repo-GAL4 trh1/trh2) was also about 3 folds to the repo-GAL4 control (0. 23 ± 0. 04, n = 11, Fig 4D). When we reduced sima levels in the trh1/trh2 mutant by repo-GAL4-driven UAS-sima-RNAi, Wg signals were suppressed to a level equivalent to that in the repo-GAL4 control (0. 25 ± 0. 05, n = 9 for repo-GAL4/sima-RNAi trh1/trh2). Interestingly, sima-RNAi knockdown in glia of the repo-GAL4 control had no effect on the Wg level (0. 25 ± 0. 02, n = 10 for repo-GAL4/sima-RNAi), suggesting that Sima is induced in the trh1/trh2 mutant to upregulate Wg expression but has no role in basal Wg expression in the wild-type. Taken together, we suggest that glial Sima is required for Wg upregulation at the NMJs of the trh1/trh2 mutant. If glia-secreted Wg is responsible for satellite bouton induction in the trh1/trh2 mutant, then glia-specific wg knockdown should phenocopy sima knockdown in glia to suppress satellite bouton formation. In control, satellite boutons were still prominent in the trh1/trh2 mutant bearing only UAS-wg-RNAi (Fig 4E and 4F, 17. 0 ± 2. 8%, n = 8 for wg-RNAi trh1/trh2). The satellite bouton phenotype was suppressed in the trh1/trh2 mutant bearing both repo-GAL4 and UAS-wg-RNAi (4. 4 ± 1. 4%, n = 10 for repo-GAL4/wg-RNAi trh1/trh2). Also, overexpression of Sima by repo-GAL4 induced satellite bouton formation (Fig 3B, 11. 9 ± 2. 3%, n = 10 for repo-GAL4/UAS-sima), which was suppressed by co-expressing the wg-RNAi transgene (Fig 3B, 2. 3 ± 1. 3%, n = 7 for repo-GAL4/UAS-sima UAS-wg-RNAi). Taken together, these results strongly suggest that glia-secreted Wg mediates Sima activity in promoting satellite bouton formation in the trh mutant. As Wg signals are secreted from both glia and presynaptic neurons [34,36], we found that reduction of Wg signals from neurons also suppressed satellite bouton formation in the trh1/trh2 mutant (Fig 5A and 5B, 10. 1 ± 1. 5%, n = 10 for elav-GAL4 trh1/trh2; 6. 0 ± 1. 0%, n = 8 for elav-GAL4/wg-RNAi trh1/trh2). Also, neuronal and glial but not muscle overexpression of Wg induced satellite bouton formation (S4A and S4B Fig). Glial and neuronal knockdown of Wg reduced total Wg levels at the trh1/trh2 NMJ, although the reduction by neuronal knockdown was not significant (S4C and S4D Fig). Thus, neuronal Wg also contributes to satellite bouton formation. However, only glial overexpression of Sima induced higher levels of Wg, but not neuronal or muscle overexpression of Sima (S4E and S4F Fig). Thus, while neuronal expression of Wg contributes to the overall level at NMJs in the trh mutant, Sima-induced Wg expression is likely glial-specific. At larval NMJs, Wg signaling in pre-synaptic and post-synaptic sites through distinct signaling pathways [35–37]. However, the receptor Frizzled2 (Fz2) is involved in transducing the signaling pathway activities in both sites. We tested whether pre- or post-synaptic Wg signaling is involved in satellite bouton formation by fz2 knockdown in the trh mutant. Neuronal expression of fz2-RNAi by elav-GAL4 suppressed satellite bouton formation in the trh1/trh2 mutant (Fig 5A and 5B, 11. 8 ± 2. 7%, n = 9 for fz2-RNAi trh1/trh2; 10. 1 ± 1. 5%, n = 10 for elav-GAL4 trh1/trh2; and 2. 5 ± 1. 1%, n = 9 for elav-GAL4/fz2-RNAi trh1/trh2). In contrast, muscle expression of fz2-RNAi by MHC-GAL4 had no significant difference to the controls (6. 4 ± 1. 6%, n = 10 for MHC-GAL4 trh1/trh2 and 7. 3 ± 1. 6%, n = 8 for MHC-GAL4/fz2-RNAi trh1/trh2). Thus, the elevated Wg level in the trh mutant is mainly transduced in presynaptic sites to modulate satellite bouton formation. Inactivation of Wg signaling leads to a reduction of the more stabilized microtubule loops within synaptic boutons, which could be visualized by immunostaining for the microtubule-binding protein Futsch [35,36]. We examined whether elevated Wg signaling alters microtubule loops in presynaptic boutons of the trh1/trh2 mutant by Futsch immunostaining and found significantly more Futsch-positive loops within the boutons of the trh mutant (Fig 5C and 5D, 2. 92 ± 0. 29, n = 12 for trh1/+; and 9. 56 ± 1. 11, n = 9 for trh1/trh2), supporting the elevation of presynaptic Wg signaling. Glial processes invade synaptic boutons to match the growth of NMJs [38], which intrigued us to assess whether glia in the trh1/trh2 mutant exhibits morphological change. In a live fillet preparation for imaging NMJs, we found that glial processes labeled by GFP invaded the area of synaptic boutons in the trh1/trh2 mutant, whereas glial processes were relatively restrained from the bouton areas in the control (Fig 5E). Quantification of the glial process overlaying the synaptic bouton area revealed significantly greater area of overlap in the trh1/trh2 mutant relative to control (Fig 5F, 6. 2 ± 1. 0%, n = 10 for trh1/+; and 16. 7 ± 3. 7%, n = 10 for trh1/trh2). This increased extent of glial processes in the synaptic area may facilitate signal transduction from glia to synaptic boutons for structural reorganization. Taken together, these results suggest that Wg plays a prominent role in the trh1/trh2 mutant to transduce the hypoxia signal from glia to modify presynaptic bouton structure. Given the evident morphological changes at trh1/trh2 NMJs, we wondered if locomotion is affected in mutant larvae. We observed larvae crawling under free-movement conditions and found that wild-type control and trh1/+ heterozygous larvae presented smooth crawling paths (Fig 6A), with an average speed of 0. 64 ± 0. 09 mm/s (n = 13) in w1118 and 0. 45 ± 0. 08 mm/s (n = 12) in trh1/+ (Fig 6B). However, the trh1/trh2 larvae had much shorter paths and a slower speed of 0. 14 ± 0. 02 mm/s (n = 14). The head turning angle of the trh1/trh2 mutant was comparable to both controls, not contributing to the slow movement (Fig 6C, 7. 8 ± 0. 6 degree/s, n = 13 for w1118; 9. 7 ± 1. 2 degree/s, n = 12 for trh1/+; and 11. 3 ± 1. 9 degree/s, n = 14 for trh1/trh2). Larval crawling is a rhythmic behavior involving a series of periodic strides (S1 Movie) [39]. We noticed uncoordinated crawling in the trh1/trh2 larvae, with their posterior body segments failing to follow the rhythmic movement (S2 Movie). We recorded larval forward crawling and constructed kymographs to represent the stride cycle. In wild-type larvae, normal and consistent periodic strides were apparent with regular head and tail displacements (Fig 6D, left panel). Similar to the wild-type, head movements of trh1/trh2 larvae were smooth and periodic, albeit slower. However, tail movements of trh1/trh2 larvae were abrupt (Fig 6D, right panel). While wild-type larvae crawled completely normal (n = 10), 69. 2 ± 9. 6% (n = 10) of the strides of trh1/trh2 larvae were uncoordinated (Fig 6E), which might contribute to the slower crawling of the trh1/trh2 mutant. This uncoordinated stride cycle prompted us to examine the bouton morphology in A2-A6 segments of the trh1/trh2 larvae (Fig 7A and 7B). Strikingly, large numbers of satellite boutons were detected in segments A2 (18. 0 ± 4. 3%, n = 10) and A3 (20. 7 ± 4. 1%, n = 10) and an intermediate level of satellite boutons was detected in segment A4 (9. 2 ± 2. 6%, n = 9). The numbers of satellite boutons were low in more posterior segments A5 (4. 9 ± 2. 3%, n = 10) and A6 (3. 2 ± 1. 7%, n = 9). In wild-type larvae, all segments had relatively normal bouton morphology except the A3 segment (0. 6 ± 0. 4%, n = 10 in A2,6. 3 ± 1. 7%, n = 10 in A3,0. 7 ± 0. 4%, n = 10 in A4,0. 5 ± 0. 5%, n = 10 in A5, and 0. 9 ± 0. 6%, n = 9 in A6). While high numbers of satellite boutons appeared in the trh mutant, the total numbers of boutons in individual segments were comparable between wild-type and trh1/trh2 (Fig 7C, w1118: A2,176. 6 ± 15. 0, n = 10; A3,76. 1 ± 4. 8, n = 9; A4,85. 8 ± 4. 1, n = 10; A5,66. 1 ± 5. 4, n = 10; A6,40. 0 ± 3. 7, n = 9; trh1/trh2: A2,173. 2 ± 8. 8, n = 10; A3,106. 6 ± 15. 3, n = 10; A4,74. 1 ± 9. 9, n = 9; A5,60. 7 ± 9. 1, n = 10; A6,33. 1 ± 3. 6, n = 9). Therefore, these analyses suggest that satellite boutons are prone to appear in more anterior than posterior segments. We further examined whether glial Sima and Wg have any role on modifying bouton morphology in the posterior A6 segment of the trh1/trh2 larvae. With the satellite boutons at a basal level in the A6 segment of trh1/trh2 (Fig 7B), we tested whether overexpression of Sima or Wg could induce satellite boutons in the trh1/trh2 mutant. Overexpression of Sima by repo-GAL4 in trh1/trh2 induced some satellite boutons (S5A and S5B Fig, 6. 8 ± 2. 3%, n = 10), which showed no significant difference to the trh1/trh2 mutant carrying repo-GAL4 (2. 8 ± 1. 5%, n = 8). Overexpression of Wg by repo-GAL4 in trh1/trh2 displayed a basal level of satellite boutons (2. 6 ± 1. 5%, n = 9). Also, glial wg-RNAi knockdown suppressed satellite bouton formation in the A3 segment of trh1/trh2 (Fig 4F), but had no effect on the morphological phenotype in the A6 segment (S5C and S5D Fig). Thus, the analysis of these data suggests that Wg and Sima might have relatively specific roles to induce satellite bouton formation in the anterior A3 segment. Given the satellite bouton phenotype in the trh1/trh2 larvae, we assessed basal synaptic transmission properties, firstly at muscle 6 of the A3 segment. The amplitude of spontaneous release, or the miniature evoked junctional potential (mEJP), was slightly but non-significantly reduced (Fig 8A), from 1. 4 ± 0. 1 mV (n = 10) in w1118 to 1. 2 ± 0. 1 mV (n = 13) in trh1/trh2 (Fig 8C). Comparable frequencies were detected between w1118 (1. 9 ± 0. 1 Hz, n = 10) and trh1/trh2 (2. 4 ± 0. 3 Hz, n = 13) (Fig 8D). The amplitudes of EJP were also comparable between w1118 (54. 6 ± 2. 1 mV, n = 10) and trh1/trh2 (50. 9 ± 4. 2 mV, n = 13) (Fig 8B and 8E). The quantal content, calculated by dividing the EJP amplitude with that of mEJP, was slightly but non-significantly increased, from 40. 2 ± 3. 3 (n = 10) in w1118 to 46. 9 ± 4. 5 (n = 13) in trh1/trh2 (Fig 8F). We then evaluated the synaptic transmission properties of muscle 6 for the A6 segment. Between the w1118 control and the trh1/trh2 mutant, the mEJP amplitudes (1. 2 ± 0. 1, n = 9 v. s. 1. 5 ± 0. 2 mV, n = 8), the mEJP frequencies (2. 0 ± 0. 3 Hz, n = 9, v. s. 1. 4 ± 0. 2 Hz, n = 8), and the EJP amplitudes (46. 2 ± 3. 7 mV, n = 9 v. s. 41. 6 ± 5. 7 mV, n = 8) remained similar without significant difference (Fig 8A–8E). However, the slight increase in mEJP and the slight decrease in EJP in trh1/trh2 lead to a significant reduction in the quantal content, from 38. 9 ± 3. 7 (n = 9) in w1118 to 27. 9 ± 1. 5 (n = 8) in trh1/trh2 (Fig 8F). Thus, the impaired synaptic activity of the A6 segment in the mutant larvae may underlie the defective stride cycles of the posterior segments.
Our results suggest that Trh has a late developmental role in tracheal morphogenesis, in addition to its well-characterized role in early tracheal cell fate specification [23,24]. We observed defective tracheal structure in the trh mutant (S2A Fig), which may result in hypoxic conditions inside the larval body. The increase in terminal branch number (S2A and S2B Fig) may be a response to oxygen supply deficiency [12]. Moreover, the increases in Sima protein levels and ODD-GFP reporter expression indicate reduced internal oxygen levels (Fig 2A and 2B and S3E and S3F Fig). Finally, the satellite bouton phenotype in the trh mutant was recapitulated by rearing larvae under hypoxia, and it was suppressed by rearing larvae under hyperoxia. Taken together, these observations suggest that cells in the trh mutant sense low oxygen levels caused by the defective tracheal system and respond by elevating Sima protein levels. It is not clear how profound this effect is for other types of larval cells. Based on our ODD-GFP and Sima immunostaining patterns (Fig 2A and 2B and S3E and S3F Fig), many types of cells are likely to be affected [40]. We suggest that glia is the major cell type mediating satellite bouton formation in the trh mutant under hypoxia. While Sima was increased ubiquitously, manipulating the levels of Sima or Ftg, the negative regulator of Sima, in glia modulates satellite bouton formation (Fig 3 and S3A and S3B Fig). Elevated Sima levels induce tracheal sprouting in tracheal cells, as well as protrusions in non-tracheal cells [12]. Interestingly, we also observed protrusion of glial processes into synaptic area in the trh mutant, indicative of a glial response (Fig 5E and 5F). Several types of cells in Drosophila have been shown to respond to hypoxia [32,41]. For instance, under hypoxia, elevated Sima levels induce the expression of Breathless (Btl, the FGF receptor) in tracheal cells that branch out seeking cells that express Branchless (Bnl) /FGF, with this latter process also being partially dependent on Sima [12,13]. In an alternative pathway, atypical soluble guanylyl cyclases can mediate graded and immediate hypoxia responses mainly in neurons [42,43]. Drosophila glia have not been reported to sense and respond to hypoxia, but mammalian astrocytes in the central nervous system have been shown to be involved in these processes. In a mouse model for middle cerebral artery occlusion, astrocyte activation was shown to play a crucial role in ischemic tolerance, which is mediated through P2X7 receptor-activated HIF-1α upregulation [18]. Under physiological hypoxia, reduced mitochondrial respiration leads to the release of intracellular calcium and exocytosis of ATP-containing vesicles, thereby signaling the brainstem to modulate animal breathing [44]. Our results reveal a role for Drosophila larval glia in sensing hypoxia via the conventional HIF-1α/Sima pathway. We also demonstrate that under hypoxia, glia modulate the formation of synaptic boutons (Fig 3E and 3F). These results clearly place the glia-modulated morphology of synaptic boutons in the context of hypoxia responses. Our study further establishes that in response to hypoxia, Wg is a glial signal that modulates synaptic bouton formation. Two sources of Wg, presynaptic motor neurons and glia, are involved in synaptic growth and remodeling [34,36]. Our results suggest that Sima upregulates the level of Wg secreted from glia to modulate synapse formation in the trh mutant or in control larvae grown under hypoxia. In hypoxic macrophages, HIF-1α ediates the induction of Wnt11, which is a mammalian homolog of Wg [45]. It is likely wg is a direct target of Sima in Drosophila. We found the HIF1-α binding motif (CGTG) at the -269 nucleotide sequence in the wg promoter. Also, direct binding of Sima to this consensus site was reported in a systematic ChIP-seq experiment [46]. We further show that the level of Wg is controlled by glial Sima in the trh mutant (Fig 4C and 4D) and that Wg mediates Sima-induced satellite bouton formation at trh NMJs (Fig 3B). Also, glial overexpression of Sima upregulates Wg levels at NMJs (S4E and S4F Fig). Wg is also expressed from presynapses [35]. Thus, neuronal wg knockdown partially suppressed satellite bouton phenotypes in the trh mutant (Fig 5A and 5B), while neuronal wg overexpression in wild-type induced the phenotypes (S4A and S4B Fig). These results are consistent with the idea that presynaptic Wg contributes to the overall pool of Wg at NMJs of the trh mutant. As a secreted morphogen, Wg functions in both pre-synaptic and post-synaptic sites [35–37]. At presynaptic terminals, the canonical Wg pathway induces microtubule loop formation to regulate synaptogenesis. We also detected an increase in microtubule loops in the trh mutant (Fig 5C and 5D), consistent with a role for Wg signaling in modulating synaptic reorganization. Postsynaptic Wg signaling leads to subsynaptic reticulum differentiation [35], which was not apparent in the trh mutant (S1D Fig), suggesting that Wg might be a component of the complex hypoxia response that induces synaptic bouton reorganization. Brief exposure to hypoxia induces immature spines and impaired synaptic function in hippocampal neurons [17]. The morphological change to satellite boutons at the A3 segment of the trh NMJs was not accompanied by altered synaptic transmission (Fig 8), which may be compensated during long-term hypoxia. The satellite boutons, also named as bunch boutons, have been described in spastin mutants [47,48]. As an AAA ATPase, Spastin severs microtubules to facilitate transport to distal axon segments [49]. Accordingly, the spastin mutant also exhibits a lack of microtubules at terminal boutons [48]. In contrast, the trh mutant presented an increase of the more stabilized microtubule loops (Fig 5C and 5D). Microtubule loops have been linked to synaptic bouton stabilization, and an excess of microtubule loops has been associated with increased synaptic bouton formation [50,51]. The altered morphology of satellite boutons may be part of the structural changes necessary to maintain normal synaptic transmission under hypoxia. The trh and spastin mutants also exhibit differences in synaptic function, with loss of spastin function slightly enhancing spontaneous synaptic transmission release but reducing evoked synaptic transmission [48]. Thus, although the morphology of synaptic boutons at trh NMJs resembles that of spastin mutants, satellite boutons at trh NMJs retain synaptic functions, unlike the impaired synaptic transmission of spastin mutant boutons. The size of NMJs in muscles 6/7 decreases from the anterior to posterior segments, which could represent a coupling with muscle growth [52,53], thereby maintaining similar electrophysiological efficacy at anterior and posterior NMJs (Fig 8). Interestingly, our findings show that synaptic responses in the trh mutant differ, with satellite boutons only appearing in anterior segments (Fig 7A and 7B). Furthermore, synaptic transmission at trh NMJs remained normal in the anterior A3 segment but was impaired in the posterior A6 segment (Fig 8). These observations are consistent with the idea that satellite bouton formation is a part of a homeostatic response to restore synaptic activity. Why synapses are not reorganized in the posterior segments remains elusive. We failed to detect an upregulation of Wg in the A6 segment (S4C and S4D Fig), and glial Sima overexpression even in the A6 segment of the trh mutant failed to increase satellite boutons significantly (S5A and S5B Fig). Thus, the upregulation of Wg by Sima may be segment-dependent, which awaits further study. Motor neurons in the ventral nerve cord project much longer axons to muscles in posterior segments compared to anterior ones. It has been shown that axonal transport to posterior segments is more vulnerable to inefficient transport conditions. For example, mutation of long-chain Acyl-CoA synthetase impairs the balance between anterograde and retrograde transport, causing distally-biased axonal aggregations and affecting the growth and functioning of synapses [54]. It is possible that glia-derived Wg signals may not be efficiently transported to posterior segments during hypoxia. This polar difference in synaptic activity and bouton morphology may contribute to the uncoordinated movements of the trh mutant larvae. Alternatively, defective locomotion in posterior segments of the trh mutant is independent of the glial modulation of bouton morphological changes. Larval forward locomotion, propelled by peristaltic contraction, is controlled by different circuits targeting anterior and posterior segments. The GABAergic SEZ-LN1 neurons specifically control posterior A6 and A7 segmental muscle contraction by inhibiting A27h premotor neurons, which promotes longitudinal muscle contraction during larval forward crawling [55]. Specific alteration of the circuit in the posterior segments may lead to the locomotion defect in the trh mutant.
All flies were reared at 25°C. w1118 was used as wild-type control and to backcross with trh1 or trh2. The sources of fly strains are as follows: trh2, elav-GAL4, MHC-GAL4, repo-GAL4, UAS-trh, UAS-sima, UAS-sima-RNAi, Nrv2-GAL4, and His2Av-mRFP were obtained from Bloomington Drosophila Stock Center (BDSC); trh1, NP6293-GAL4, and UAS-wg-RNAi from Kyoto Stock Center; and UAS-trh-RNAi, UAS-fga-RNAi, and UAS-fz2-RNAi from Vienna Drosophila Resource Center (VDRC). Also used were btl-GAL4[56], moody-GAL4 [57], alrm-GAL4[58], UAS-wg, and GFP-ODD [29]. The repo-cyto-GFP line was generated with the sequence for cytoplasmic GFP under the control of the 4. 3 kb repo promoter, which recapitulates the full repo expression pattern. Larvae in a food vial were transferred at 1 day after egg laying (AEL) to a ProOx (model 110, BioSpherix, Lacona, NY) oxygen-controlled chamber. Oxygen or nitrogen was infused into the chamber to a desired concentration (5% or 50%), which was maintained until assay. The NMJ phenotypes were analyzed as previously described [59]. For live tissue preparation to detect repo-cyto-GFP expression, non-fixed larvae were dissected and the larval fillets were incubated with anti-horseradish peroxidase (HRP, 1: 10) in phosphate buffered saline (PBS) for 10 minutes. Primary antibodies used were against Synapsin (3C11, mouse, 1: 100; Developmental Studies Hybridoma Bank, DSHB), HRP-Cy5 (rabbit, 1: 100; Jackson ImmunoResearch), Dlg (mouse, 1: 100, DSHB), GluRIIA (mouse, 1: 100, DSHB), dPAK (rabbit, 1: 1000), GluRIII (rabbit, 1: 1000), Brp (nc82, mouse, 1: 100, DHSB), Sima (guinea pig, 1: 1000) [60], Repo (mouse, 1: 1000, DSHB), Wg (4D4, mouse, 1: 10, DSHB), and Futsch (22C10, mouse, 1: 100, DSHB). Secondary antibodies used were anti-rabbit or -mouse 488, Cy3, or Cy5 (1: 1000, Jackson ImmunoResearch). For Wg immunostaining, larval preparations of different genotypes were marked and immunostaining was performed in the same test tube. NMJs in muscle 6/7 of A3 segments (or A2-A6 in Fig 7 and A6 in S4 and S5 Figs) of wandering third-instar larvae were analyzed. Confocal images were acquired via LSM510 confocal microscopy (Carl Zeiss) using 40x water, 40x water immersion (for live tissue in Fig 5E), or 100x oil objectives. All presented images are projections of confocal z-stacks. Numbers of satellite boutons, total boutons, and microtubule loops were counted manually. The percentage of satellite boutons was calculated as the number of satellite boutons divided by the total boutons (satellite + normal ones) for each NMJ, and the average percentage is calculated from about 6–13 NMJs for each genotype. The immunofluorescence intensities of Wg and HRP were analyzed by ImageJ. HRP-positive regions were chosen to measure mean intensities of Wg and HRP. After subtracting the intensity with the background one, the ratio of Wg levels to HRP levels was presented as the normalized Wg intensity. The overlapping area of GFP and HRP projections was chosen by the “AND” operator in ImageJ, which was divided by HRP area for the percentage. Each dot in the bar graph represents the data from a single NMJ of a larva, and 6–13 NMJs from 2–5 independent experiments were pooled for quantification. Embryos were acquired by means of LSM510 confocal microscopy (Carl Zeiss) using a 20x objective, and were analyzed as previously described [29]. For Fig 2B, each dot in the bar graph represents data from a single embryo in which fluorescence was measured in at least 35 cells. Basal transmission properties were analyzed at NMJs of muscle 6/7 in specified segments of wandering third-instar larvae as previously described [61], with some modifications. The larval body wall was dissected in cold calcium-free HL3 solution and recorded in HL3 solution containing 0. 4 mM CaCl2 at room temperature. Mid third instar larvae (feeding stage) were placed on black agar plates (2% agar with black food coloring in 25 × 20 cm2 dishes) at room temperature for filming. Video recording by a Sony Xperia Z1 camera started after 1 min habituation and lasted for 5 min, and it was analyzed using Ctrax software [62]. The (x, y) positions were used to calculate the crawling distance between two successive frames, and crawling speed was derived by dividing total distance travelled by time. The change in angle of larvae between two frames was divided by time to represent rotational angles. The forward crawling assay was a modification of a previous study [39]. Larvae were transferred into a tunnel (~1 mm width) made in 2% black agar. Specimens were video-recorded for 3–10 minutes using a Leica S8 APO microscope. Kymographs were constructed using the MultipleKymograph plug-in for ImageJ (NIH). Only forward crawling was counted, and 7–10 steps for each of ten larvae were analyzed for each genotype. Statistic data were analyzed by Mann-Whitney test or one-way ANOVA with Tukey' s Multiple Comparison post-test, and shown by scatter plots with bar using GraphPad Prism.
Oxygen is essential for animals to maintain their life such as growth, metabolism, responsiveness, and movement. It is therefore important to understand how animal cells trigger hypoxia response and adapt to hypoxia thereafter. Both mammalian vascular and insect tracheal branches are induced to enhance the oxygen delivery. However, the study of hypoxia response in the nervous system remains limited. In this study, we assess the morphology of Drosophila neuromuscular junctions (NMJs), a model system to study development and function of synapses, in two hypoxia conditions, one with raising wild-type larvae in hypoxia, and the other in the trachealess (trh) mutant in which the trachea is defective, causing insufficient oxygen supply. Interestingly, the glial processes, normally wrapping around the axons, invade into synaptic boutons of NMJs under hypoxia. Also, the hypoxia-induced factor Sima activates the Wg signal in glia and the secreted Wg signal reorganizes the synaptic boutons as a response to hypoxia. This synaptic bouton reorganization might maintain normal synaptic transmission and locomotion of larvae.
invertebrates
medicine and health sciences
oxygen
neuroscience
biological locomotion
pulmonology
animals
animal models
organisms
developmental biology
drosophila melanogaster
model organisms
hypoxia
experimental organism systems
drosophila
research and analysis methods
medical hypoxia
animal studies
animal cells
life cycles
chemistry
insects
arthropoda
cellular neuroscience
eukaryota
cell biology
neurotransmission
physiology
neurons
biology and life sciences
crawling
physical sciences
cellular types
larvae
chemical elements
Glial response to hypoxia in mutants of NPAS1/3 homolog Trachealess through Wg signaling to modulate synaptic bouton organization