Introduction

Neural circuitry underlying mature adult behaviours emerges from a combination of developmental gene expression programs and experience-dependent neuronal activity. Holometabolous insects such as Drosophila, reconfigure their nervous system during metamorphosis to generate circuitry capable of supporting adult behaviours (Levine, 1984). During pupal development, neurons undergo maturation of their electrical properties with a gradual increase in depolarizing responses and consequent synaptic transmission (Hardie et al., 1993; Jarvilehto and Finell, 1983). Apart from voltage-gated Ca²⁺ channels and fast-acting neurotransmitters specific to ionotropic receptors, the developing nervous system also uses neuromodulators, that target metabotropic receptors, show slower response kinetics and are capable of affecting a larger subset of neurons by means of diffusion aided volumetric transmission (Taber and Hurley, 2014). Although neuromodulators alter intrinsic neuronal properties (Marder, 2012) by changes in gene expression, the molecular mechanisms through which this is achieved and maintained over developmental timescales needs further understanding.

Ca²⁺ signals generated by neuronal activity can determine neurotransmitter specification (Spitzer, 2012), synaptic plasticity (O’Hare et al., 2022; Takechi et al., 1998), patterns of neurite growth (Gu and Spitzer, 1995) and gene expression programs (Ciceri et al., 2022; Rosenberg and Spitzer, 2011) over developmental timescales (McKinney et al., 2022) where output specificity is defined by signal dynamics. Neuromodulators generate intracellular Ca²⁺ signals by stimulation of cognate G-Protein Coupled Receptors (GPCRs) linked to the inositol 1,4,5– trisphosphate (IP₃) and Ca²⁺ signalling pathway. IP₃ transduces intracellular Ca²⁺ release through the endoplasmic reticulum (ER) localised ligand-gated ion channel, the inositol 1,4,5-trisphosphate receptor (IP₃R; Streb et al., 1984), followed by store-operated Ca²⁺ entry (SOCE) through the plasma membrane localised Orai channel (Prakriya and Lewis, 2015; Thillaiappan et al., 2019). Cellular and physiological consequences of Ca²⁺ signals generated through SOCE exhibit different time-scales and sub-cellular localisation from activity induced signals, suggesting that they alter neuronal properties through novel mechanisms.

Expression studies, genetics and physiological analysis support a role for IP₃/Ca²⁺ and SOCE in neuronal development and the regulation of adult neuronal physiology across organisms (Hasan and Sharma, 2020; Mitra and Hasan, 2022; Somasundaram et al., 2014). In Drosophila, neuromodulatory signals of neurotransmitters and neuropeptides stimulate cognate GPCRs on specific neurons to generate IP₃ followed by intracellular Ca²⁺ release and SOCE (Agrawal et al., 2013; Megha and Hasan, 2017; Shakiryanova et al., 2011). Genetic and cellular studies identified neuromodulatory inputs and SOCE as an essential component for maturation of neural circuits for flight (Pathak et al., 2015; Venkiteswaran and Hasan, 2009). A focus of this flight deficit lies in a group of central dopaminergic neurons (or DANs) that include the PPL1 and PPM3 DANs (Liu et al. 2012) marked by THD’ GAL4, alternately referred to henceforth as flight promoting DANs (fpDANs). Among the fpDANs some PPL1 DANs project to the γ2α’1 lobe of the Mushroom Body (MB; Mao and Davis 2009) a dense region of neuropil in the insect central brain, that forms a relay centre for flight (Sharma and Hasan, 2020). Axonal projections from Kenyon cells (KC) carry sensory inputs (Cervantes-Sandoval et al., 2017; Tsao et al., 2018; Yagi et al., 2016) and those from DANs carry internal state information (Mao and Davis, 2009; Riemensperger et al., 2005; Zolin et al., 2021) to functional compartments of the MB, where dopamine release modulates the KC and MB output neuron synapse (MBONs; Aso et al. 2014). The KC-DAN-MBON tripartite synapse, carries a dynamically updating representation of the motivational and behavioural state of the animal (Aso and Rubin, 2016; Berry et al., 2015; Owald et al., 2015; Waddell, 2016). Here we have investigated the molecular mechanisms that underlie the ability of neuromodulatory acetylcholine signals, acting through SOCE during circuit maturation, to determine fpDAN function required for Drosophila flight.

Results

A spatio-temporal requirement for SOCE determines flight

To understand how neuromodulator stimulated SOCE alters neuronal properties over developmental time scales we began by refining further the existing spatio-temporal coordinates of SOCE requirement for flight. Flight promoting dopaminergic neurons (DANs) with a requirement for SOCE have been identified earlier by expression of a dominant negative Orai transgene, which renders the channel Ca²⁺ impermeable, thus abrogating all SOCE (Orai^E180A; Figure 1A; Pathak et al. 2015; Yeromin et al. 2006). These include a smaller subset of THD’GAL4 marked DANs, where acetylcholine promotes dopamine release through the muscarinic acetylcholine receptor (mAchR) by stimulating IP₃/Ca²⁺ signaling (Sharma and Hasan, 2020). Orai^E180Aexpression in a subset of 21-23 dopaminergic neurons marked by THD’ GAL4 (Figure 1B) led to complete loss of flight (Figure 1C) while overexpression of the wildtype Orai transgene had a relatively minor effect (Figure S1A).

Figure 1.

THD’ marks DANs in the PPL1, PPL2, and PPM3 clusters of the adult central brain (Figure 1B) of which the PPL1 (10-12) and PPM3 (6-7) cell clusters constitute the fpDANs (Pathak et al., 2015). Among these 16-19 DANs, two pairs of PPL1 DANs projecting to the γ2α’1 lobes of the mushroom body (marked by the MB296B split GAL4 driver and schematised in Figure S1B; Aso et al. 2014; Aso and Rubin 2016) were identified as contributing to the flight deficit to a significant extent (Figures 1C; S1C). The MB296B DANs form part of a central flight relay centre identified earlier as requiring IP₃/Ca²⁺ signaling (Sharma and Hasan, 2020). That the THD’ neurons are required for flight was further confirmed by inhibiting their function. Acute activation of either a temperature sensitive Dynamin mutant (Shibire^ts; Kitamoto 2001), which blocks exocytosis at an elevated temperature, or acute expression of the tetanus toxin light chain fragment (TeTxLC; Sweeney et al. 1995) which cleaves Synaptobrevin, an essential component of the synaptic release machinery in THD’ neurons led to severe flight defects (Figure S1D).

Previous work has demonstrated that intracellular Ca²⁺ release through the IP₃R, that precedes SOCE (Figure 1A), is required during late pupal development in THD’ marked DANs for flight (Pathak et al., 2015; Sharma and Hasan, 2020). The precise temporal requirement for SOCE in THD’ neurons was investigated by inducing Orai^E180A transgene expression for specific periods of pupal development and in adults with the tubulinGAL80^ts based temperature sensitive expression system, (TARGET-Temporal And Regional Gene Expression Targeting; McGuire et al., 2004). Approximately 100 hours of pupal development were binned into 24 hour windows, wherein SOCE was abrogated by Orai^E180Aexpression, following which, normal development and growth was permitted. Flies were assayed for flight 5 days post eclosion. Abrogation of SOCE by expression of the Orai^E180A dominant negative transgene at 72 to 96 hrs after puparium formation (APF), resulted in complete loss of flight as compared to minor flight deficits observed upon expression of Orai^E180A during earlier developmental windows (Figure 1D). Abrogation of flight also occurred upon Orai^E180A expression during the first 2 days post-eclosion (see discussion) whereas abrogation of SOCE 5 days after eclosion as adults resulted in only modest flight deficits (Figure 1D), indicating that SOCE is required for maturation of THD’ marked flight dopaminergic neurons. Earlier work has shown that loss of SOCE by expression of Orai^E180A does not alter the number of dopaminergic neurons or affect neurite projection patterns of PPL1 and PPM3 neurons (Pathak et al., 2015). Of these two clusters, PPM3 DANs project to the ellipsoid body in the central complex (Kong et al., 2010), and PPL1 DANs project to the mushroom body and the lateral horn (Mao and Davis, 2009) and aid in the maintenance of extended flight bouts (Sharma and Hasan, 2020).

Loss of SOCE in late pupae leads to a re-organisation of neuronal gene expression

The molecular consequences of loss of SOCE in THD’ neurons were investigated next by undertaking a comprehensive transcriptomic analysis from fluorescence activated and sorted (FACS) THD’ neurons with or without Orai^E180A expression and marked with eGFP (Figure 1E; S1E). THD’ neurons were obtained from pupae at 72 ± 6h APF (essential SOCE requirement for flight; Figure 1E). Expression analysis of RNAseq libraries generated from THD’ neurons revealed a reorganisation of the transcriptome upon loss of SOCE (Figure 1F). Expression of 822 genes was downregulated (with a fold change cut-off of < –1) as assessed using 3 different methods of Differentially Expressed Genes (DEG) analysis (Figure 1G) whereas 137 genes were upregulated (Figure S1F). To understand if loss of SOCE affects genes expressed in neurons from the pupal stage, we used the modEncode dataset (Celniker et al., 2009; modENCODE Consortium et al., 2010) to reconstruct developmental trajectories of all genes expressed in THD’ neurons (Figure S1G). Using an unsupervised clustering algorithm, we classified genes expressed in THD’ neurons into 3 clusters where ‘Cluster 0’ is low expression throughout development; ‘Cluster 1’ exhibits a larval peak in expression and ‘Cluster 2’ exhibits a pupal peak in expression (Figure 1H). The majority of expressed genes (>95%) classify as low expression throughout (Cluster 0) and were removed from further analysis. Genes belonging to either the downregulated gene set or the upregulated gene set in SOCE-deficient THD’ neurons and a random set of genes were analysed further. A comparison of the proportion of DEGs classified in the larval and pupal clusters revealed that >75% of downregulated genes belonged to the pupal peak (Cluster 2) whereas <20% of either the upregulated genes or a random set of genes classified as belonging to the pupal peak (Figure 1I). Moreover, downregulated genes exhibit higher baseline expression in wildtype pupal THD’ neurons (Figures S1H, I). These analyses suggest that SOCE induces expression of a set of genes in THD’ neurons during the late pupal phase which are subsequently required for flight. These are henceforth referred to as SOCE-responsive genes.

SOCE regulates gene expression through a balance of Histone 3 Lysine 36 trimethylation and Histone 3 Lysine 27 trimethylation

Gene ontology (GO) analysis of SOCE-responsive genes revealed ‘Transcription’, ‘Ion transporters’, ‘Ca²⁺ dependent exocytosis’, ‘GPCRs’, ‘Synaptic components’, and ‘Kinases’ as top GO categories (Figure 2A). Among these, ‘Transcription’ and ‘Ion transporters’ represented categories with the highest enrichment. Further analysis of genes that classified under ‘Transcription’ revealed SET domain containing genes that encode histone lysine methyltransferases implicated in chromatin regulation and gene expression (Dillon et al., 2005) and among the SET domain containing genes, Set1 and Set2 showed distinct downregulation (Figure 2B). One of these genes, the H3K36me3 methyltransferase Set2 (Figure 2C), was identified in a previous RNAseq from larval neurons with loss of IP₃R-mediated Ca²⁺ signaling (Mitra et al., 2020). RT-PCRs from sorted THD’ neurons confirmed that Set2 levels are indeed downregulated (by 76%) in THD’ neurons expressing Orai^E180Aand to the same extent as seen upon expression of Set2^RNAi (79% downregulation; Figure 2C). Set1 and Set2 perform methyltransferase activity at H3K4 and H3K36 histone residues respectively (Ardehali et al., 2011; Stabell et al., 2007). Both these markers are epigenetic signatures for transcriptional activation, where H3K4me3 is enriched at the 5’ end of the gene body and H3K36me3 is enriched in the gene bodies of actively transcribed genes; schematised in Figure S2A). In the adult CNS, Set2, is expressed at 4-fold higher levels compared to Set1 (Figure S2B; modENCODE Consortium et al., 2010). Set2 also shows a steep 5.5 fold increase in expression levels during the pupal to adult transition, compared to a 2 fold increase in Set1 (Figure S2B). These observations coupled with the previous report of Set2 function downstream of IP₃R/Ca²⁺ signaling in larval glutamatergic neurons(Mitra et al., 2020), led us to pursue the role of Set2 in THD’ neurons.

Figure 2.

Set2 encodes the only Drosophila methyltransferase which is specific for H3K36 trimethylation (Stabell et al., 2007). To understand the functional significance, if any, of down-regulation of Set2 in THD’ neurons upon loss of SOCE, four independent Set2 RNAi constructs were obtained. Knockdown of Set2 in THD’ neurons with all four RNAi lines tested resulted in significant flight defects (Figure 2D; S2C). Set2 function for flight was additionally verified in the PPL1 DANs projecting to the γ2α’1 lobe of the mushroom body (MB296BGAL4; Figure S2D). To test the hypothesis that Set2 expression and function for flight is regulated by SOCE, we overexpressed Set2 in the Orai^E180A background, using GAL4-UAS driven heterologous Set2^WTexpression or CRISPR-dCas9::VPR driven overexpression (Gilbert et al., 2013) from the endogenous locus, and measured flight (Figure 2D data in purple and brown). Flies expressing Orai^E180Ain the THD’ neurons, which are flightless, exhibit significant rescue in flight bout durations upon Set2 overexpression, through either method (Figure 2D, compare red with purple and brown data), but not upon expression of a control transgene, GCamP6m (Figure S2E, compare data in green and red). Additionally, overexpression of a key SOCE component, the ER Ca²⁺ sensor STIM in a Set2^RNAi background led to a rescue (Figure S2E, compare pink and grey data). Excess STIM presumably drives higher SOCE sufficient to rescue flight bout durations caused by deficient Set2 levels. These two sets of data taken together with downregulation of Set2 upon expression of Orai^E180A (Figure 2C) support the hypothesis that SOCE driven expression of Set2 in fpDANs is required for flight. That Set2 function downstream of SOCE, occurs specifically through its methyltransferase activity was supported through experiments where knockdown of the demethylase Kdm4B, the primary histone demethylase expressed in these neurons (Figure S2F), rescued flight in THD’>Orai^E180A flies (Figure 2D; data in pink). Knockdown of the lower expressed Drosophila H3K36 demethylase isoform, Kdm4A, did not show a similar rescue (Figure S2G; data in pink). Moreover, both Set2 depletion and loss of SOCE (THD’>Orai^E180A) lead to a decrease in the normalised H3K36me3 signal in THD’ neurons (Figure 2E).

To understand if deficient H3K36me3 is found in the SOCE-responsive genes, we quantified the enrichment of H3K36me3 at SOCE-responsive loci from an H3K36me3 ChIP-Seq dataset (modENCODE Consortium et al., 2010) from wild-type adult heads (Figure 2F). We observed an enrichment of the H3K36me3 signal over the gene bodies of the SOCE-responsive genes (Figure 2F). Additionally, genes that were more affected by loss of SOCE (greater extent of downregulation), had a correspondingly higher H3K36me3 signal (Figure 2G) as compared with a random set of genes (Figure 2G; data in grey), further indicating that one of the ways SOCE regulates gene expression is through Set2 mediated deposition of H3K36me3.

The gene E(z) that encodes a component of the PRC2 complex (EZH2), which deposits H3K27me3 is up-regulated upon loss of SOCE (Figure 2B) suggesting that SOCE could affect additional histone modifications. While H3K36me3 is a marker for transcriptional activation (Krogan et al., 2003), H3K27me3 is a repressive signature which silences transcription (Cai et al., 2021). These two marks are antagonistic as deduced from studies which have shown that deposition of H3K36me3 allosterically inhibits the Polycomb Repressive Complex (PRC2), thereby preventing it from depositing the H3K27me3 signature at the same genomic loci (Finogenova et al., 2020). A comparison of H3K36me3 and H3K27me3 peaks over the 822 SOCE-responsive genes revealed a higher enrichment of H3K36me3 versus H3K27me3 in wild type fly heads (Figure 2H; modENCODE Consortium et al., 2010), suggesting that robust expression of these genes in adult neurons occurs through a SOCE-dependent mechanism initiated in late pupae (schematised in Figure 2I). To test this hypothesis, we fed SOCE-deficient flies (THD’GAL4>Orai^E180A) GSK343, a pharmacological inhibitor (Verma et al., 2012) of the EZH2 component of PRC2 that is known to reduce H3K27me3 chromatin marks and performed flight assays. Feeding of GSK343 lead to partial rescue of flight in THD’> Orai^E180A flies in a dose dependent manner with maximal flight bout durations of up to 400 sec (Figure 2J; S2H).

Normal cellular responses of PPL1 DANs require Orai mediated SOCE and H3K36 trimethylation

Next we investigated Ca²⁺ responses in SOCE and H3K36 trimethylation deficient THD’ PPL1 DANs upon stimulation with a neuromodulatory signal for the muscarinic acetylcholine receptor (mAChR). It is known that THD’ marked PPL1 DANs receive cholinergic inputs that stimulate store Ca²⁺ release through the IP₃R, leading to SOCE through Orai (Figure 3A; Ebihara et al., 2006; Sharma and Hasan, 2020). In support of this idea, either expression of Orai^E180A or knockdown of the IP₃R attenuated the response to carbachol (CCh), a mAChR agonist, in PPL1 DANs (Figure 3B-D; Figure S3A-C). Importantly, knockdown of Set2 also abrogated the Ca²⁺ response to CCh (Figure 3B-D), whereas overexpression of a transgene encoding Set2 in THD’ neurons either with loss of SOCE (Orai^E180A) or with knockdown of the IP₃R (itpr^RNAi), lead to significant rescue of the Ca²⁺ response, indicating that Set2 is required downstream of SOCE (Figure 3B-D) and IP₃/Ca²⁺ signalling (Figure S3A-C).

Figure 3.

An understanding of how Set2 might rescue cellular IP₃/Ca²⁺ signaling was obtained in an earlier study where it was demonstrated that Set2 participates in a transcriptional feedback loop to control the expression of key upstream components such as the mAChR and the IP₃R in a set of larval glutamatergic neurons (Mitra et al., 2020). To test whether Set2 acts through a similar transcriptional feedback loop in the current context, THD’ DANs were isolated using FACS from appropriate genetic backgrounds and tested for expression of key genes required for IP₃/Ca²⁺ signaling (mAChR and itpr) and SOCE (Stim and Orai). Loss of SOCE upon expression of Orai^E180A lead to a significant decrease in the expression of itpr (72% downregulation; Figure 3E), mAChR (56% downregulation; Figure 3E), Stim (63% downregulation; Figure 3E) in THD’ DANs. As expected Orai expression (Figure 3E) increased 2-fold presumably due to overexpression of the Orai^E180A transgene. Importantly, knockdown of Set2 in THD’ neurons also led to downregulation of itpr (43%), mAChR (78%) and Stim (61%) (Figure 3E). Overexpression of Set2 in the background of Orai^E180Aled to a rescue in the levels of itpr (69% increase), mAChR (43% increase), and Stim (40% increase) (Figure 3E). Overexpression of Set2 in wildtype THD’ neurons resulted in upregulation of mAChR and Orai (Figure 3E). These findings indicate that while SOCE regulates Set2 expression (Figure 2C), Set2 in turn functions in a feedback loop to regulate expression of key components of IP₃/Ca²⁺ and SOCE such as mAchR, IP₃R, STIM and Orai (Figure 3E). Thus, ectopic Set2 overexpression in SOCE deficient THD’ neurons, leads to increase in expression of key genes that facilitate intracellular Ca²⁺ signalling and SOCE (schematised in Figure 3F).

Trithorax-like or Trl is an SOCE responsive transcription factor in THD’ DANs

The data so far identify SOCE as a key developmental regulator of the neuronal transcriptome in THD’ marked DANs, where it upregulates Set2 expression and thus enhances the activation mark of H3K36 trimethylation on specific chromatin regions. However, the mechanism by which SOCE regulates expression of Set2, and other relevant effector genes remained unresolved (Figure 3F). In mammalian T cells, SOCE leads to de-phosphorylation of the NFAT (Nuclear Factor of Activated T-cells) family of transcription factors by Ca²⁺/Calmodulin sensitive phosphatase Calcineurin, followed by their nuclear translocation and ultimately transcription of relevant target genes (Hogan et al., 2003). Unlike mammals, the Drosophila genome encodes a single member of the NFAT gene family, which does not possess calcineurin binding sites, and is therefore insensitive to intracellular Ca²⁺ and SOCE (Keyser et al., 2007).

In order to identify transcription factors (TFs) that regulate SOCE-mediated gene expression in Drosophila neurons, we examined the upstream regions (up to 10 kb) of SOCE-regulated genes using motif enrichment analysis (Zambelli et al., 2009) for TF binding sites (Figure 4A). This analysis helped identify several putative TFs with enriched binding sites in the regulatory regions of SOCE-regulated genes (Figure 4B, S4A) which were then analyzed for developmental expression trajectories (Hu et al., 2017) in the Drosophila CNS (Figure 4B; lower panel). Among the top 10 identified TFs, the homeobox transcription factor Trl/GAF had the highest enrichment value, lowest p-value (Figure 4B) and was consistently expressed in the CNS through all developmental stages, with a distinct peak during pupal development (Figure 4B; lower panel). To test the functional significance of Trl/GAF, we tested flight in flies with THD’ specific knockdown of Trl using two independent RNAi lines (Figure 4C) as well as in existing Trl mutant combinations that were viable as adults (Figure S4B). In all cases significant flight deficits were observed (Figure 4C, S4B). Moreover, the flight deficits could be rescued by raising SOCE either directly through overexpression of STIM or indirectly by Set2 overexpression (Figure 4C; data in brown and pink respectively).

Figure 4.

To further test Trl function, as an effector of intracellular Ca²⁺ signaling and SOCE, we investigated genetic interactions of Trl^13C, a hypomorphic mutant Trl allele, with an existing deficiency for the ER-Ca²⁺ sensor STIM (STIM^KO), an essential activator of the Orai channel (Wang et al., 2010) as well as existing mutants for an intracellular ER-Ca²⁺ release channel, the IP₃R (Joshi et al., 2004). While STIM^KO/+ flies demonstrate normal flight bout durations, adding a single copy of the Trl^13C allele to the STIM^KO heterozygotes (i.e., STIM^KO/Trl^13C trans-heterozygotes) resulted in significant flight deficits, which could be rescued by overexpression of STIM or Set2 (Figure 4D). A single copy of Trl^13C placed in combination with a single copy of various itpr mutant alleles, also showed a significant reduction in the duration of flight bouts (Figure S4C). Together these genetic data provide strong support for Trl as a SOCE-responsive TF. Due to the strong flight deficit observed by expression of Orai^E180Awe were unable to test genetic interactions of Trl with Orai directly. Orai RNAi strains exhibit off-target effects and Orai hypomorphs are unavailable.

The requirement of Trl for SOCE-dependent gene expression was tested directly by measuring Set2 transcripts in THD’ neurons with knockdown of Trl (THD’>Trl^RNAi). Set2 has 20 Trl binding sites in a 2 Kb region upstream of the transcription start site (Figure S4D) and knockdown of Trl resulted in downregulation of Set2 (32%) and the Set2 regulated genes itpr (56%) and mAchR (73%; Figure 4E). Moreover, a trans-heterozygotic combination of hypomorphic Trl alleles (Trl^13C/62) had markedly reduced levels of brain H3K36me3 (Figure 4F).

To test if Trl drives cellular function in the fpDANs, we stimulated the mAChR with CCh and measured Ca²⁺ responses in fpDANs with knockdown of Trl. Compared to WT fpDANs, which respond robustly to cholinergic stimulation, Trl^RNAi expressing DANs, exhibit strongly attenuated responses (Figure 4G-I). Overexpression of Set2 in the Trl^RNAi background rescued the cholinergic response (Figure 4G-I). The rescue of both flight (Figure 4C, D) and the cholinergic response (Figure 4G-I) by overexpression of Set2 in flies with knockdown of Trl in THD’ neurons (THD’>Trl^RNAi), confirms that Trl acts upstream of Set2 to ensure optimal neuronal function and flight.

Trl function downstream of SOCE targets neuronal excitability through changes in ion channel gene expression including VGCCs

The results obtained so far indicate that Trl could function as an intermediary transcription factor between SOCE and its downstream effector gene Set2. Overexpression of Trl, however, is unable to rescue the loss of flight caused by loss of SOCE (THD’>Orai^E180A; Figure 5A). Moreover, Trl transcript levels are unchanged in the Orai^E180Acondition (Figure 5B) suggesting that Trl requires Ca²⁺ influx through SOCE for its function to go from an ‘inactive’ form to an ‘active’ form (schematized in Figure 5C). Although Trl does not possess a defined Ca²⁺ binding domain, it has been shown to interact with a diverse set of proteins (Lomaev et al., 2017) including Ca²⁺ sensitive kinases, and is known to undergo phosphorylation (Zhai et al., 2008). These data support a model (schematized in Figure 5C) wherein Ca²⁺ influx through SOCE sustains Trl activation, leading to expression of Set2.

Figure 5.

Next we investigated the extent to which the identified SOCE-Trl-Set2 mechanism impacts fpDAN function. ‘Ion transport’ is among the top GO terms identified in SOCE-responsive genes (Figure 2A). Indeed, multiple classes of ion channel genes including several Na^+, K⁺ and Cl^- channels are downregulated in fpDANs upon expression of Orai^E180A(Fig 5D, E). Regulation of ion channel gene expression by Trl was indicated from the enrichment of Trl binding sites in regulatory regions of some ion channel loci (Figure 5F). We purified fpDANs with knock down of Trl (THD’>Trl^RNAi-1) and measured expression of a few key voltage-gated ion channel genes (Fig 5F). NaCP60E, para (Na⁺ channels), cac, ca-α1D (VGCC subunits), hk, and eag (outward rectifying K⁺ channels) are all downregulated between 0.6-0.9 fold upon Trl knockdown (Fig 5G) as well as upon loss of SOCE (Fig 5B). Taken together, these results indicate that the expression of key voltage-gated ion channel genes which are required for the depolarization-mediated response, maintenance of electrical excitability and neurotransmitter release is a focus of the transcriptional program set in place by SOCE-Trl-Set2.

To test the functional consequences of altered ion channel gene expression downstream of Trl, neuronal activity of the fpDANs was tested by means of KCl-evoked depolarisation. PPL1 DANs exhibited a robust Ca²⁺ response that was lost upon knock down of Trl (Figure 5H-J; data in orange). In consonance with the earlier behavioural experiments (Figure 5A), overexpression of Trl⁺ in THD’ DANs did not rescue the KCl response in PPL1 DANs with loss of SOCE (OraiE^180A; Trl^OE, Figure 5I, J), further reinforcing the hypothesis that, Trl activity in fpDANs is dependent of Ca²⁺ entry through Orai.

Response to KCl is also lost in PPL1 neurons by knockdown of Set2 and restored to a significant extent by overexpression of Set2 in THD’ DANs lacking SOCE (Orai^E180A;Set2^OE; Figure 6A-C). We confirmed that the KCl response of PPL1 neurons requires VGCC function, either by treatment with Nimodopine (Xu and Lipscombe, 2001), an L-type VGCC inhibitor or by knockdown of a conserved VGCC subunit cac (cac^RNAi). Both forms of perturbation abrogated the depolarisation response to KCl in PPL1 neurons (Figures 6D-F; data in purple or orange respectively). Moreover, the Ca²⁺ entry upon KCl-mediated depolarization in PPL1 neurons is a cell-autonomous property as evident by measuring the response after treatment with the Na⁺ channel inhibitor, Tetrodotoxin (TTX; Figures 6D-F; data in magenta).

Figure 6.

Having confirmed that the Ca²⁺ response towards KCl depolarisation requires VGCCs, we tested the expression level of key components of Drosophila VGCCs (Figure 6G) in THD’ DANs, including cacophony (cac), ca-α1D, ca-α1T, and ca-ý. All four subunits of Drosophila VGCC are downregulated upon loss of SOCE (Orai^E180A; Figure 6H) in THD’ DANs whereas overexpression of Set2 in the background of Orai^E180Arestored expression of the four VGCC subunits tested (Figure 6H), thus explaining recovery of the KCl response upon Set2 overexpression in PPL1 neurons lacking SOCE (Orai^E180A; Figures 6A-C).

The functional significance of downregulation of VGCC subunits by the SOCE-Trl-Set2 pathway was tested directly by knockdown of the 4 VGCC subunits independently in PPL1 neurons followed by measurement of flight bout durations. Significant loss of flight was observed with knockdown of each subunit (Figure 6I) though in no case was the phenotype as strong as what is observed with loss of SOCE (Orai^E180A; Figure 1D). These data suggest that in addition to VGCC subunit expression, appropriate expression of other ion channels in the fpDANs is of functional significance. This idea is further supported by the observation that loss of flight upon loss of SOCE cannot be restored by overexpression of cac alone (Figure 6J) indicating that optimal neuronal activity requires an ensemble of genes, including ion channels, whose expression is regulated by SOCE-Trl-Set2.

THD’ DANs require SOCE for developmental maturation of the neuronal excitability threshold

The functional relevance of changes in neuronal excitability during pupal maturation of the PPL1-dependent flight circuit was tested next. Inhibition of THD’ DANs from 72 to 96 hours APF using acute induction of the inward rectifying K⁺ channel Kir2.1(Johns et al., 1999) or optogenetic inhibition through the hyperpolarizing Cl^- channel GtACR2 (Govorunova et al., 2015) resulted in significant flight deficits (Figure 7A; data in dark or light green). Similar flight defects were recapitulated upon hyperactivation of THD’ neurons through either optogenetic (CsChrimson; Klapoetke et al., 2014) or thermogenetic (TrpA1; Viswanath et al., 2003) stimulation (Figure 7A; data in orange or red), indicating that these neurons require balanced neuronal activity during a critical window in late pupal development, failing which the flight circuit malfunctions. To test if restoring excitability in fpDANs lacking SOCE (THD’>Orai^E180A) is sufficient to restore flight, we induced hyperexcitability by overexpressing NachBac (Nitabach, 2006) a bacterial Na⁺ channel. NachBac expression rescued both flight (THD’>Orai^E180A; Figure 7B) and excitability (Figures 7C-E). A partial rescue of flight was also obtained by optogenetic stimulation of neuronal activity through activation of THD’>Orai^E180A DANs using CsChrimson either 72-79 hrs APF or 0-2 days post eclosion (Figure 7E). In this case, flight rescues were accompanied by a corresponding rescue of CsChrimson activation induced Ca²⁺ entry (Figures 7F, G). Moreover inducing neuronal hyperactivation with indirect methods such as excess K⁺ supplementation (Figure S7A) or impeding glial K⁺ uptake, by genetic depletion of the glial K⁺ channel sandman (Weiss et al., 2019), partially rescued flight in animals lacking SOCE in the THD’ DANs (Figure S7B). Together, these results indicate that SOCE, through Trl and Set2 activity, determines the threshold of fpDAN activation during circuit maturation by regulating expression of voltage-gated ion channel genes, like cac.

Figure 7.

fpDANs also require a balance between H3K36me3/H3K27me3 mediated epigenetic regulation (Figure 2K). To understand if this epigenetic balance affects flight through modulation of neuronal excitability, we measured KCl-induced depolarizing responses of PPL1 DANs in Orai deficient animals (THD’>Orai^E180A) fed on the H3K27me3 antagonist GSK343 (0.5 mM). GSK343 fed animals were sorted into 2 groups (Fliers/Non-Fliers) on the basis of flight bout durations of greater or lesser than 30 s (Figure S7C), and tested for responses to KCl (Figures S7D, E). THD’>Orai^E180A flies fed on GSK343 showed a rescue in KCl responses, with a clearly enhanced response in the fliers compared to the non-fliers (Figures S7D, E), thereby reiterating the hypothesis that SOCE-mediated regulation of activity in these neurons acts through epigenetic regulation between opposing histone modifications.

THD’ DANs require SOCE-mediated gene expression for neuromodulatory dopamine release in the mushroom body γ lobe

Dopaminergic neurotransmission requires presynaptic voltage gated Ca²⁺ channels (VGCCs; Brimblecombe et al., 2015), and has been implicated in the maturation of intrinsic neuronal excitability during circuit formation (Lieberman et al., 2018). A number of fpDANs send projections to the γ2α’1 lobe of the MB (Figure 8A). We examined the consequences of loss of SOCE on CCh-evoked DA release at the γ2α’1 region of the mushroom body (Figure 8B; red arrow). For this purpose we used the dopamine sensor GRAB-DA (Sun et al., 2018). Loss of SOCE (THD’>Orai^E180A) attenuated DA release (Figures 8C, D), which could be partially rescued by overexpression of Set2. Taken together these data identify an SOCE-dependent gene regulation mechanism acting through the transcription factor Trl, and the histone modifier Set2 for timely expression of genes that impact diverse aspects of neuronal function including excitability and sustained neurotransmitter release (schematised in Figure 8E).

Figure 8.

Discussion

Over the course of development, neurons define an excitability set point within a dynamic range (Truszkowski and Aizenman, 2015), which stabilizes existing connections (Mayseless et al., 2023) and enables circuit maturation (Johnson-Venkatesh et al., 2015). In this study, we identify an essential role for the store-operated Ca²⁺ channel Orai in setting the excitability threshold of a subset of dopaminergic neurons central to a MB circuit for Drosophila flight. Orai-mediated Ca²⁺ entry, initiated by neuromodulatory acetylcholine signals, is required for amplification of a signalling cascade, that begins by activation of the homeo-box transcription factor Trl, followed by upregulation of several genes, including Set2, a gene that encodes an enzyme for an activating epigenetic mark, H3K36me3. Set2 in turn establishes a transcriptional feedback loop to drive expression of key neuronal signaling genes, including a repertoire of voltage-gated ion channel genes required for neuronal activity in mature PPL1 dopaminergic neurons.

SOCE supports gene expression transitions during critical developmental windows

Studies in the Drosophila mushroom body have identified spontaneous bouts of voltage-gated Ca²⁺ channel mediated neuronal activity, through as yet unknown mechanisms, which occur during early adulthood driving refinement and maturation of behaviours such as associative learning (Leinwand and Scott, 2021). Our studies here, on a subset of 21-23 central dopaminergic neurons (Figures 1B, C) of which some send projections to the mushroom body and regulate flight behaviour, provide a mechanistic explanation for neuromodulation-dependent intracellular signaling culminating in transcriptional maturation of a circuit supporting adult flight. In the critical developmental window of 72-96 hrs APF, a cohort of SOCE-responsive genes, that include a range of ion channels, undergo a wave of induction. Loss of SOCE at this stage thus renders these neurons functionally incompetent (Figures 3B-D, 6G-I, 7D-F) and abrogates flight (Figure 1D). Developmentally assembled neuronal circuits require experience/activity-dependent maturation (Akin and Zipursky, 2020). The requirement of SOCE for flight circuit function during the early post eclosion phase (0-2 days) suggests that maturation of the flight circuit extends into early adulthood, where presumably feedback from circuit activity may further refine synaptic strengths (Sugie et al., 2018). Expression of Orai^E180A in 5 day old adults had no effect on flight (Figure 1E), indicating that either SOCE is not required for maintaining gene expression in adults or that wildtype Orai channel proteins from late pupal and early adults perdure for long periods. The absence of any visible changes to neurite patterning suggest that SOCE in the MB-DANs works primarily to refine synaptic function. In summary, our findings here identify Orai-driven Ca²⁺ entry as a key signaling step for driving coordinated gene expression enabling the maturation of nascent neuronal circuits and sustenance of adult behaviour.

SOCE regulates a balance between competing epigenetic signatures

Chromatin structure and function is dynamic over the course of brain development (Kishi and Gotoh, 2018). One way neurons achieve dynamic spatio-temporal control over developmental gene expression is via epigenetic mechanisms such as post translational histone modifications (Geng et al., 2021). Cells possess an extensive toolkit of histone modifiers, with characterised effects on transcriptional output by either activating or repressing gene expression (Bannister and Kouzarides, 2011). However, relatively little, is known about how modifiers of histone marks are in turn regulated to bring about the requisite changes in neuronal gene expression over developmental timescales.

The SET-domain containing family of histone modifiers (Figure 2B), including H3K36me3 methyltransferase Set2 (Figure 2C), identified here appear to function as key transcriptional effectors induced downstream of neuromodulatory inputs that stimulate Store-operated Ca²⁺ entry through Orai during pupal and adult maturation of flight promoting dopaminergic neurons. SOCE-driven Set2-mediated H3K36me3 enhances the expression of key GPCRS, components of intracellular Ca²⁺ signaling (Figure 2A) and ion channels (Figures 6A, B) for optimal dopaminergic neuron function (Figure 7D-F) and flight (Figure 1C). Rescue of flight by genetic depletion of an epigenetic ‘eraser’ Kdm4B, an H3K36me3 demethylase (Figure 2D) suggests that a balance between perdurance of H3K36me3 and its removal is actively maintained on the expressed genes. Interestingly, another member of the SET domain family, E(z) is up-regulated upon loss of SOCE (Figure 2B). E(z) is a component of the Drosophila PRC2 complex (Polycomb repressive complex), which represses gene expression through H2K27me3 (Margueron and Reinberg, 2011). Structural studies on Drosophila nucleosomes have elucidated that the H3K36me3 modification allosterically inhibits the PRC2 complex activity (Finogenova et al., 2020). Our findings support a model wherein at key developmental stages, SOCE determines the level of these competing epigenetic signatures thus allowing gene expression by enhancing Set2 mediated H3K36me3 (Fig 2H). Loss of SOCE leads to deficient Set2-mediated H3K36me3 (Figure 2E), a likely shifting of the balance in favour of PRC2 mediated H3K27me3 and overall repression in gene expression (as observed in Fig S1H, I). Feeding SOCE-deficient animals a pharmacological inhibitor of PRC2 (GSK434) led to a significant rescue in behaviour (Figure 2J) presumably by disinhibition of H3K27me3 mediated gene repression. Though flight initiation is rescued in a dose dependent manner, the duration of flight bouts is not sustained (Figure S2H) indicating that suppression of H3K27me3 marks in the SOCE-deficient flies is partial. Our findings indicate additional roles for other histone modifications such as Set1 mediated H3K4me3, which may work in concert to regulate developmental gene expression programs stimulated by SOCE. Future studies directed at a comprehensive cataloguing of multiple epigenetic signatures in defined neuronal subsets using cell-specific techniques should help elucidate these mechanisms.

Trl as an SOCE responsive transcription factor

SOCE-mediated regulation of gene expression has been reported in mammalian neural progenitor cells (Somasundaram et al., 2014) and Drosophila pupal neurons (Pathak et al., 2015; Richhariya et al., 2017), through as yet unknown mechanisms. A combination of motif enrichment analysis over regulatory regions of SOCE-responsive genes and expression enrichment analysis in cells of interest, helped generate a list of putative SOCE-responsive transcription factors (Figure 4A, B). Among these we experimentally validated Trl/Trithorax-like/GAGA factor as a transcription factor required for maturation of excitability in flight promoting dopaminergic neurons (Figure 4). Although Trl function has been reported in the context of early embryonic development, where it has been implicated in zygotic genome activation (Gaskill et al., 2021) (ZGA), expression of Hox genes (Shimojima, 2003), dosage compensation (Greenberg et al., 2004), and expression of a Drosophila voltage-gated calcium channel subunit in germ cell development (Dorogova et al., 2014), his is the first ever report of its role in regulating neuronal gene expression in the context of SOCE and circuit maturation.

In vitro studies on Drosophila Trl reveal that it utilizes a Q-rich intrinsically disordered domain (IDD) to self-multimerize or interact with a wide range of accessory proteins (Wilkins and Lis, 1999). Multimeric complexes of Trl reside on chromatin with an exceptionally long residence time and maintain chromatin in an ‘open’ state (Tang et al., 2022). Trl has also been reported to directly interact with components of the Transcription initiation and elongation complex (Chopra et al., 2008; Li et al., 2013), support long distance promoter-enhancer interactions (Mahmoudi, 2003), mediate active ATP-dependent chromatin remodeling by maintaining nucleosome free stretches (Tsukiyama et al., 1994), and regulating global gene expression by controlling transcriptional stalling and pausing (Tsai et al., 2016). SOCE-dependent gene regulation may rely upon multiple transcription factors acting in concert in a cell type and developmental stage specific contexts, of which Trl may be just one. Future studies directed at other possible transcription factors downstream of SOCE would be of interest.

SOCE and the control of neuronal excitability and cholinergic neuromodulation of a tripartite synapse in the mushroom body

Neurons undergo maturation of their electrical properties with a gradual increase in depolarizing responses and synaptic transmission over the course of pupal development (Jarvilehto and Finell, 1983). Here we show that a subset of dopaminergic neurons require a balance of ion channels for optimal excitation and inhibition essential for adult function. Neuromodulatory signals, such as acetylcholine are essential for acquiring this balance and act via an SOCE-Trl-Set2 mechanism during late pupal and early adult stages. The ability of Set2 overexpression to rescue SOCE-deficient phenotypes (Figures 6A-C) indicates that anomalous gene expression is the underlying basis of this phenotype. Indeed, SOCE deficient dopaminergic neurons show reduced expression of an ensemble of voltage-gated ion channel genes (Figures 5E, F). including genes encoding the outward rectifying K⁺ channel shaw, slow-inactivating voltage gated K⁺ channel shaker, a Ca²⁺ gated K⁺ channel slowpoke and cacophony a subunit of the voltage gated Ca²⁺ channel (Figure 6G). Ion channels like Shaker and slowpoke play an important role in establishing resting membrane potential (Singh and Wu, 1990), neuronal repolarisation (Lichtinghagen et al., 1990) and mediating afterhyperpolarisation (AHP) after repeated bouts of activity (Ping et al., 2011). Though dopaminergic neurons require Set2 mediated expression of cacophony for excitability, cacophony is in itself insufficient to rescue the loss in excitability and flight deficits upon loss of SOCE (Figure 6J). Both flight and KCl-depolarisation in SOCE-deficient neurons could, however, be rescued by expression of a heterologous depolarisation activated Na⁺ channel NachBac (Nitabach, 2006) (Figures 7B-D), which increases neuronal excitability by stimulating a low threshold positive feedback loop and to a lesser extent by CsChrimson-mediated optogenetic induction of neuronal activity (Figure 7E). The poorer rescue by optogenetic activation may stem from the fact that sustained bursts of activity are non-physiological and can deplete the readily releasable pool of neurotransmitters (Arrigoni and Saper, 2014; Kaeser and Regehr, 2017). Other indirect forms of inducing neuronal hyperactivation (Figures S7A, B) in SOCE-deficient neurons also achieved minor rescues in flight. These results identify SOCE as a key driver downstream of developmentally salient neuromodulatory signals for expression of an ion channels suite that enables generation of intrinsic electrical properties and functional maturation of the PPL1 DANs, and the flight circuit.

Methods

Fly maintenance

Drosophila strains were grown on standard cornmeal medium consisting of 80 g corn flour, 20 g glucose, 40 g sugar, 15 g yeast extract, 4 ml propionic acid, 5 ml p-hydroxybenzoic acid methyl ester in ethanol and 5 ml ortho-butyric acid in a total volume of 1 l (ND) at 25°C under a light-dark cycle of 12 h and 12 h. Canton S was used as wild type throughout. Mixed sex populations were used for all experiments. Several fly stocks used in this study were sourced using FlyBase (https://flybase.org), which is supported by a grant from the National Human Genome Research Institute at the US National Institutes of Health (#U41 HG000739), the British Medical Research Council (#MR/N030117/1) and FlyBase users from across the world. The stocks were obtained from the Bloomington Drosophila Stock Centre (BDSC) supported by NIH P40OD018537. All fly stocks used and their sources are listed in Supplementary Table 1.

Flight Assays

Flight assays were performed as previously described (Manjila and Hasan, 2018). Briefly, flies aged 3–5 days of either sex were tested in batches of 8–10 flies, and a minimum of 30 flies were tested for each genotype. Adult flies were anaesthetized on ice for 2–3 min and then tethered between at the head-thorax junction using a thin metal wire and nail polish. Post recovery at room temperature for 2-3 min, an air puff was provided as stimulus to initiate flight. Flight duration was recorded for each fly for 15 min. For all control genotypes, GAL4 or UAS strains were crossed to the Wild Type strain, Canton S. Flight assays are represented in the form of a swarm plot, wherein each dot represents a flight bout duration for a single fly. The colours indicate different genotypes. The ý Flight parameter refers to the mean difference for comparisons against the shared CS control which is shown as a Cumming estimation plot(Ho et al., 2018). On the lower axes, mean differences are plotted as bootstrap sampling distributions. Each mean difference is depicted as a dot. Each 95% confidence interval is indicated by the ends of the vertical error bars. Letters beneath each distribution refer to statistically indistinguishable groups after performing a Kruskall-Wallis test followed by a post hoc Mann-Whitney U-test (p<0.05).

FACS

Fluorescence-activated cell sorting (FACS) was used to enrich eGFP labelled DANs from pupal/adult. The following genotypes were used for sorting: wild type (THD’GAL4> UAS-eGFP), Orai^E180A (THD’GAL4> Orai^E180A), Set2^IR-1 (THD’GAL4> Set2^IR-1), Orai^E180A; Set2^OE (THD’GAL4> Orai^E180A; Set2^OE), Set2^OE (THD’GAL4> Set2^OE), Trl^IR-1 (THD’GAL4> Trl^IR-1). Approximately 100 pupae per sample were washed in 1×PBS and 70% ethanol. Pupal/adult CNSs were dissected in Schneider’s medium (ThermoFisher Scientific) supplemented with 10% fetal bovine serum, 2% PenStrep, 0.02 mM insulin, 20 mM glutamine and 0.04 mg/ml glutathione. Post dissection, the CNSs were treated with an enzyme solution [0.75 g/l Collagenase and 0.4 g/l Dispase in Rinaldini’s solution (8 mg/ml NaCl, 0.2 mg/ml KCl, 0.05 mg/ml NaH2PO4, 1 mg/ml NaHCO3 and 0.1 mg/ml glucose)] at room temperature for 30 min. They were then washed and resuspended in ice-cold Schneider’s medium and gently triturated several times using a pipette tip to obtain a single-cell suspension. This suspension was then passed through a 40 mm mesh filter to remove clumps and kept on ice until sorting (less than 1 h). Flow cytometry was performed on a FACS Aria Fusion cell sorter (BD Biosciences) with a 100 mm nozzle at 60 psi. The threshold for GFP-positive cells was set using dissociated neurons from a non GFP-expressing wild-type strain (Canton S). The same gating parameters were used to sort other genotypes in the experiment. GFP-positive cells were collected directly in TRIzol and then frozen immediately in dry ice until further processing. Details for all reagents used are listed in Supplementary Table 2.

RNA-Seq

RNA from at least 600 sorted THD’ DANs from the relevant genotypes, expressing UAS-eGFP was subjected to 14 cycles of PCR amplification (SMARTer Seq V4 Ultra Low Input RNA Kit; Takara Bio). 1 ng of amplified RNA was used to prepare cDNA libraries (Nextera XT DNA library preparation kit; Illumina). cDNA libraries for 4 biological replicates for both control (THD’GAL4/+) and experimental (THD’GAL4>UAS-Orai^E180A) genotypes were run on a Hiseq2500 platform. 50-70 million unpaired sequencing reads per sample were aligned to the dm6 release of the Drosophila genome using HISAT2(Kim et al., 2015, 2019) and an overall alignment rate of 95.2-96.8% was obtained for all samples. Featurecounts(Liao et al., 2014) was used to assign the mapped sequence reads to the genome and obtain read counts. Differential expression analysis was performed using three independent methods: DESeq2(Love et al., 2014), limma-voom(Ritchie et al., 2015), and edgeR(Robinson et al., 2009). A fold change cutoff of a minimum twofold change was used. Significance cutoff was set at an FDR-corrected P value of 0.05 for DESeq2 and edgeR. Volcanoplots were generated using VolcaNoseR (https://huygens.science.uva.nl/). Comparison of gene lists and generation of Venn diagrams was performed using Whitehead BaRC public tools (http://jura.wi.mit.edu/bioc/tools/). Gene ontology analysis for molecular function was performed using DAVID(Huang et al., 2008, 2009). Developmental gene expression levels were measured for downregulated genes using FlyBase(Larkin et al., 2020) and DGET(Hu et al., 2017) (www.flyrnai.org/tools/dget/web/), and were plotted as a heatmap using ClustVis (Metsalu and Vilo, 2015) (https://biit.cs.ut.ee/clustvis/).

qRT-PCRs

Central nervous systems (CNSs) of the appropriate genotype and age were dissected in 1× phosphate-buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄ and 1.8 mM KH₂PO₄) prepared in double distilled water treated with DEPC. Each sample consisted of five CNS homogenized in 500 µl of TRIzol (Ambion, ThermoFisher Scientific) per sample. At least three biological replicate samples were made for each genotype. After homogenization the sample was kept on ice and either processed further within 30 min or stored at −80°C for up to 4 weeks before processing. RNA was isolated following the manufacturer’s protocol. Purity of the isolated RNA was estimated by NanoDrop spectrophotometer (ThermoFisher Scientific) and integrity was checked by running it on a 1% Tris-EDTA agarose gel. Approximately 100 ng of total RNA was used per sample for cDNA synthesis. DNAse treatment and first-strand synthesis were performed as described previously(Pathak et al., 2015). Quantitative real time PCRs (qPCRs) were performed in a total volume of 10 µl with Kapa SYBR Fast qPCR kit (KAPA Biosystems) on an ABI 7500 fast machine operated with ABI 7500 software (Applied Biosystems). Technical duplicates were performed for each qPCR reaction. The fold change of gene expression in any experimental condition relative to wild type was calculated as 2−ΔΔCt, where ΔΔC_t=[C_t (target gene) –C_t (rp49)] Expt. – [C_t (target gene) – C_t (rp49)]. Primers specific for rp49 and ac5c were used as internal controls. Sequences of all primers used are provided in Supplementary Table 3.

Functional imaging

Adult brains were dissected in adult hemolymph-like saline (AHL: 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 Tris, pH 7.5) after embedded in a drop of 0.1% low-melting agarose (Invitrogen). Embedded brains were bathed in AHL and then subjected to functional imaging. The genetically encoded calcium sensor GCaMP6m(Chen et al., 2013) was used to record changes in intracellular cytosolic Ca²⁺ in response to stimulation with carbachol or KCl. The GRAB-DA sensor(Sun et al., 2018) was used to measure evoked dopamine release. Images were taken as a time series on a xy plane at an interval of 1 s using a 20× objective with an NA of 0.7 on an Olympus FV3000 inverted confocal microscope. A 488 nm laser line was used to record GCaMP6m / GRAB-DA measurements. All live-imaging experiments were performed with at least 10 independent brain preparations. For measuring evoked responses, KCl/Carbachol, was added on top of the samples. For optogenetics experiments, flies were reared on medium supplemented with 200 μM ATR (all trans retinal), following which neuronal activation was achieved using a 633-nm laser line to stimulate CsChrimson(Klapoetke et al., 2014) while simultaneously acquiring images with the 488 nm laser line, and the images were acquired every 1 s. The raw images were extracted using FIJI(Schindelin et al., 2012) (based on ImageJ version 2.1.0/1.53c). ΔF/F was calculated from selected regions of interest (ROIs) using the formula ΔF/F=(Ft-F0)/F0, where Ft is the fluorescence at time t and F0 is baseline fluorescence corresponding to the average fluorescence over the first 40-time frames. Mean ΔF/F time-lapses were plotted using PlotTwist(Goedhart, 2020) (https://huygens.science.uva.nl/PlotTwist/). A shaded error bar around the mean indicates the 95% confidence interval for CCh (50 µM) or KCl (70 mM) responses. Peaks for individual cellular responses for each genotype was calculated from before or after the point of stimulation, using Microsoft Excel and plotted as a paired plot using SuperPlotsOfData(Goedhart, 2021) (https://huygens.science.uva.nl/SuperPlotsOfData/). Boxplots were plotted using PlotsOfData (Postma and Goedhart, 2019) (https://huygens.science.uva.nl/PlotsOfData/).

Immunohistochemistry

Immunostaining of larval Drosophila brains was performed as described previously(Daniels et al., 2008). Briefly, adult brains were dissected in 1x phosphate buffered saline (PBS) and fixed with 4% Paraformaldehyde. The dissected brains were washed 3-4 times with 0.2% phosphate buffer, pH 7.2 containing 0.2% Triton-X 100 (PTX) and blocked with 0.2% PTX containing 5% normal goat serum (NGS) for 2 hours at room temperature. Respective primary antibodies were incubated overnight (14–16 hr) at 4°C. After washing 3-4 times with 0.2% PTX at room temperature, they were incubated in the respective secondary antibodies for 2 hr at room temperature. The following primary antibodies were used: chick anti-GFP antibody (1:10,000; A6455, Life Technologies), mouse anti-nc82 (anti-brp) antibody (1:50), rabbit anti-H3K36me3, mouse anti-RFP. Secondary antibodies were used at a dilution of 1:400 as follows: anti-rabbit Alexa Fluor 488 (#A11008, Life Technologies), anti-mouse Alexa Fluor 488 (#A11001, Life Technologies). Confocal images were obtained on the Olympus Confocal FV1000 microscope (Olympus) with a 40x, 1.3 NA objective or with a 60x, 1.4 NA objective. Images were visualized using either the FV10-ASW 4.0 viewer (Olympus) or Fiji(Schindelin et al., 2012). Details for all reagents used are listed in Supplementary Table 2.

Western Blotting

Adult CNSs of appropriate genotypes were dissected in ice-cold PBS. Between 5 and 10 brains were homogenized in 50 μl of NETN buffer [100 mM NaCl, 20 mM Tris-Cl (pH 8.0), 0.5 mM EDTA, 0.5% Triton-X-100, 1× Protease inhibitor cocktail (Roche)]. The homogenate (10-15 μl) was run on an 8% SDS-polyacrylamide gel. The protein was transferred to a PVDF membrane by standard semi-dry transfer protocols (10 V for 10 min). The membrane was incubated in the primary antibody overnight at 4°C. Primary antibodies were used at the following dilutions: rabbit anti-H3K36me3 (1:5000, Abcam, ab9050) and rabbit anti-H3 (1:5000, Abcam, ab12079). Secondary antibodies conjugated with horseradish peroxidase were used at dilution of 1:3000 (anti-rabbit HRP; 32260, Thermo Scientific). Protein was detected by a chemiluminescent reaction (WesternBright ECL, Advansta K12045-20). Blots were first probed for H3K36me3, stripped with a solution of 3% glacial acetic acid for 10 min, followed by re-probing with the anti-H3 antibody.

In silico ChIP-Seq Analysis

H3K36me3 enrichment data was obtained from a ChIP-chip dataset (ID_301) generated in Drosophila ML-DmBG3-c2 cells submitted to modEncode(Celniker et al., 2009; modENCODE Consortium et al., 2010). Enrichment scores for genomic regions were calculated using ‘computematrix’ and plotted as a tag density plot using ‘plotHeatmap’ from deeptools2(Ramírez et al., 2016). All genes were scaled to 2 kb with a flanking region of 250 bp on either end. A 50 bp length of non-overlapping bins was used for averaging the score over each region length. Genes were sorted based on mean enrichment scores and displayed on the heatmap in descending order.

Statistical tests

Non-parametric tests were employed to test significance for data that did not follow a normal distribution. Significant differences between experimental genotypes and relevant controls were tested either with the Kruskal-Wallis test followed by Dunn’s multiple comparison test (for multiple comparisons) or with Mann-Whitney U tests (for pairwise comparisons). Data with normal distribution were tested by the Student’s T-test. Statistical tests and p-values are mentioned in each figure legend. For calculation and representation of effect size, estimationstats.com was used.

Acknowledgements

We acknowledge the Central Imaging and Flow Cytometry Facility (CIFF, NCBS) for maintenance of microscopes and the Drosophila facility (Flyfacility, NCBS) for Fly stock maintenance and development of transgenics.

Competing Interests

The authors declare no competing interests

Funding

This work was funded by grant No. BT/PR28450/MED/122/166/2018 from the Department of Biotechnology, Govt. of India and core support by NCBS, TIFR. RM and SR received graduate student fellowships from NCBS, TIFR.

Data availability

The RNA Sequencing data has been submitted to GEO under accession number GSE230134.