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. 2021 Jul 13;219(1):iyab112. doi: 10.1093/genetics/iyab112

Loss of the Dβ1 nicotinic acetylcholine receptor subunit disrupts bursicon-driven wing expansion and diminishes adult viability in Drosophila melanogaster

Danielle Christesen 1,, Ying Ting Yang 1, Wei Chen 1, Philip Batterham 1, Trent Perry 1
Editor: K M O’Connor-Giles
PMCID: PMC8633089  PMID: 34849910

Abstract

Cholinergic signaling dominates the insect central nervous system, contributing to numerous fundamental pathways and behavioral circuits. However, we are only just beginning to uncover the diverse roles different cholinergic receptors may play. Historically, insect nicotinic acetylcholine receptors have received attention due to several subunits being key insecticide targets. More recently, there has been a focus on teasing apart the roles of these receptors, and their constituent subunits, in native signaling pathways. In this study, we use CRISPR-Cas9 genome editing to generate germline and somatic deletions of the Dβ1 nicotinic acetylcholine receptor subunit and investigate the consequences of loss of function in Drosophila melanogaster. Severe impacts on movement, male courtship, longevity, and wing expansion were found. Loss of Dβ1 was also associated with a reduction in transcript levels for the wing expansion hormone bursicon. Neuron-specific somatic deletion of Dβ1 in bursicon-producing neurons (CCAP-GAL4) was sufficient to disrupt wing expansion. Furthermore, CCAP-GAL4-specific expression of Dβ1 in a germline deletion background was sufficient to rescue the wing phenotype, pinpointing CCAP neurons as the neuronal subset requiring Dβ1 for the wing expansion pathway. Dβ1 is a known target of multiple commercially important insecticides, and the fitness costs exposed here explain why field-isolated target-site resistance has only been reported for amino acid replacements and not loss of function. This work reveals the importance of Dβ1-containing nicotinic acetylcholine receptors in CCAP neurons for robust bursicon-driven wing expansion.

Keywords: nicotinic acetylcholine receptor, CCAP, bursicon, wing expansion, somatic CRISPR

Introduction

The nicotinic acetylcholine receptors (nAChRs) are the most abundant neuronal receptor in the insect brain, responding to the native neurotransmitter acetylcholine to facilitate fast excitatory synaptic neurotransmission (Florey 1963; Croset et al. 2018). Their ubiquity in the insect nervous system, combined with their pharmacological sensitivity, also makes them ideal targets for neuroactive insecticides. Interactions between nAChRs and insecticides, and the role of nAChRs in insecticide resistance, are well-covered in the literature (Lansdell and Millar 2000; Tomizawa and Casida 2005; Millar and Denholm 2007; Perry et al. 2012; Crossthwaite et al. 2017; Homem et al. 2020). Now, unraveling the endogenous signaling roles of nAChRs across many cholinergic pathways will enhance our understanding of contributions to insect life-history traits, also offering insights into potential fitness costs underlying insecticide resistance.

nAChRs are pentameric ligand-gated ion channels, formed by five subunits arranged symmetrically around a central cation-selective pore (Corringer et al. 2000; Karlin 2002). In Drosophila melanogaster, there are 10 genes encoding nAChR subunits (Sattelle et al. 2005). Seven of these encode α subunits (Dα1-Dα7) which contain a motif (YXCC) that coordinates acetylcholine binding (Kao et al. 1984; Karlin and Akabas 1995). Three genes encode β subunits (Dβ1–Dβ3). These lack the di-cysteine motif. The subunit composition defines the pentamer “subtype” and determines the receptor pharmacology (Millar 2003; Sattelle et al. 2005; Thany et al. 2007; Lansdell et al. 2012; Ihara et al. 2020). Single-cell sequencing analysis reveals different levels of expression of individual subunit genes across the brain, suggesting that the spectrum of subtypes present may vary between different neurons and synapses (Croset et al. 2018). This variability in subtype distribution and pharmacology means that a given subunit may have specialized roles in different biological processes. Six Drosophila subunits (Dα1, Dα3, Dα4, Dα5, Dα6, and Dα7) have been briefly described for their roles in behaviors such as sleep (Shi et al. 2014; Wu et al. 2014; Somers et al. 2017), courtship (Somers et al. 2017), learning and memory (Barnstedt et al. 2016), and escape (Fayyazuddin et al. 2006). Meanwhile, five subunits (Dα1, Dα2, Dα6, Dβ1, and Dβ2) are known to have insecticide interactivity, based on heterologous expression (Watson et al. 2010, Ihara et al. 2020) and strong resistance phenotypes seen in individual loss of function mutants (Perry et al. 2008, 2012; Watson et al. 2010;).

The Dβ1 subunit is well overdue for investigation. In 2011, a field population of the global agricultural pest, Myzus persicae, the peach-potato aphid, was found to harbor a nonsynonymous mutation in the β1 nAChR subunit (Mpβ1R81T) that conferred resistance to neonicotinoids (Bass et al. 2011). The potential impact on M. persicae control could have been devastating, especially given the species’ capacity to vector hundreds of plant viruses to a huge range of host plants worldwide (Blackman and Eastop 2000). Fortunately, the Mpβ1R81T population did not thrive (Bass et al. 2011), and there has only been one other report of a field-evolved resistant β1 allele (Aphis gossypii Agβ1R81T, Koo et al. 2014) despite over 20 years of ongoing neonicotinoid usage. In vivo and in vitro studies identify β1 as a neonicotinoid target in multiple insect species (Li et al. 2010; Shi et al. 2012; Homem et al. 2020), so why haven’t more insects evolved resistance via disruption of the β1 subunit?

The simplest explanation for the absence of naturally occurring loss of function alleles is that nAChRs containing β1 subunits may have an essential role(s) in the brain. β1 is the most highly conserved nAChR subunit among insects (Jones and Sattelle 2010). It is also the most highly and broadly expressed of all nAChR subunits in the D. melanogaster nervous system (Graveley et al. 2011; Croset et al. 2018; Allen et al. 2020), suggesting loss of β1 subunits could be quite detrimental to viability. Indeed, Homem et al. (2020) recently generated a D. melanogaster Dβ1R81T strain and observed significant reductions in fertility, male longevity, larval crawling, and adult climbing performance in individuals homozygous for the R81T allele. The R81T amino acid replacement only slightly alters acetylcholine binding affinity (Shimomura et al. 2006; Ihara et al. 2020), so it is probable that complete loss of the Dβ1 subunit would have more severe consequences for viability.

Here, we have used the CRISPR-Cas9 system to investigate the complete loss of the Dβ1 subunit in D. melanogaster. We describe the pleiotropic consequences of germline genomic deletion, which include a prominent disruption in wing expansion and severely reduced adult viability. By generating GAL4-driven somatic deletions, we have also pinpointed a neuronal subset, the CCAP neurons, that require Dβ1 to execute timely release of the wing expansion hormone, bursicon. This represents the first association of a specific nAChR subunit with hormone-driven developmental cues and suggests many roles in a variety of life-history traits are yet to be uncovered for this important gene family.

Methods

Fly strains and maintenance

All fly strains used and generated are described in Supplementary Tables S1 and S2. The wAC9 control is a strain we previously generated for a collection of nAChR CRISPR mutants (Perry et al. 2021). It contains the autosomes of the actin-Cas9 strain, with a substituted w- X chromosome (and therefore lacks an actin-Cas9 element).

All flies were maintained on standard semolina fly media at 25°C under a 12:12 hours light: dark cycle unless otherwise stated. Due to the low viability of Dβ1Δex5, Dβ1Δex3, and Dβ1-TG4, any strains containing these alleles were maintained over the TM3.GFP fluorescent balancer chromosome. In preparation for all adult experiments, embryos were collected for 24 hours on apple juice agar plates and first instar larvae lacking the fluorescent balancer chromosome were transferred to standard semolina fly media where they were reared until adulthood. For larval experiments, embryos were collected for 3 hours on apple juice agar plates and first instar larvae lacking the fluorescent balancer chromosome were synchronized within 3 hours of hatching then reared on apple juice agar plates supplemented with yeast paste.

Generating Dβ1 genomic deletions using CRISPR-Cas9

Generation of Dβ1Δex5 is described elsewhere (Perry et al. 2021), and we have used the same methods to generate the smaller deletion, Dβ1Δex3. Briefly, sgRNAs targeting the Dβ1 5’ region (77 bp upstream of the 5’UTR; Dβ1_5’UTR_sgRNA), exon 3 (Dβ1_sgRNA1), and exon 5 (Dβ1_sgRNA2) were designed using the crisprflydesign.org online tool and integrated in pairs into the pCFD4 plasmid (Addgene: 49411) using the recommended protocol (Port et al. 2014) (sgRNA positions in Figure 1, primer sequences in Supplementary Table S3). Plasmids were microinjected into attP40 (25709 v-) embryos then transgenics were identified both phenotypically (v+ eye color rescue) and molecularly (Vermillion_F and 25709_External_R1 primers). Homozygotes were recovered by self-crossing the heterozygous offspring of injectees (crossing scheme in Supplementary Figure S1). Homozygotes were then crossed to nanos-Cas9 flies, producing F1 offspring with all components required for CRISPR (crossing scheme in Supplementary Figure S2). Genomic cutting was detected molecularly using Dβ1_5’UTR_sgRNA_HMA_F/Dβ1_sgRNA1_HMA_R, or Dβ1_5’UTR_sgRNA_HMA_F/Dβ1_sgRNA2_HMA_R, and sequenced using the Dβ1_5’UTR_sgRNA_HMA_F primer. Successful genomic deletions were crossed to incorporate the X chromosome and chromosome 2 from the wAC9 strain, and the TM3.GFP balancer on chromosome 3 (crossing scheme in Supplementary Figure S2). sgRNA homozygotes containing the Dβ1_5’UTR_sgRNA/Dβ1_sgRNA2 pair were also crossed into GAL4 backgrounds to enable tissue-specific somatic CRISPR crosses with UAS-Cas9 (final crosses outlined in Supplementary Figure S3) (Meltzer et al. 2019). Genomic cutting was detected in these somatic CRISPR mosaics using Dβ1_5’UTR_sgRNA_HMA_F/Dβ1_sgRNA2_HMA_R.

Figure 1.

Figure 1

Genomic deletion of the Dβ1 nAChR subunit gene. (A) Two deletions were used in this study: Dβ1Δex5 and Dβ1Δex3. The deleted regions are shown flanked by synthetic guide RNAs (sgRNA, red). The positions of the Dβ1 ligand-binding domain (LBD, green), transmembrane domains (TM1-4, green), exons (e1-6, purple), and qRT-PCR primers (orange) are shown above. The position of the T2A-GAL4 insertion in intron 1 is also indicated (Dβ1-TG4, blue). (B) qRT-PCR indicates loss of Dβ1 transcripts in both deletion strains. Error bars indicate SEM (n = 4 biological replicates, ***P < 0.001, Student’s t-test).

Generating Dβ1-TG4 using CRIMIC

An sgRNA targeting Dβ1 intron one was designed using the crisprflydesign.org online tool and integrated into the pU6-BbsI-chiRNA plasmid (Addgene: 45946) using the recommended protocol (Gratz et al. 2013). Homology arms of approximately 1 kb adjacent to the sgRNA were amplified using Q5 PCR (NEB) from genomic DNA of the actin-Cas9 strain and ligated into the pT-GEM(attP2-SA(1)-T2A-GAL4-Hsp70-lox2-3XP3-RFP) plasmid (Addgene: 62893) using the recommended protocol (Diao et al. 2015). The Dβ1-TG4 strain was made by microinjection of pU6-BbsI-chiRNA and pT-GEM plasmid DNA into actin-Cas9 embryos. Transgenic adults were selected by the eye-specific RFP marker and crossed to incorporate the X chromosome and chromosome 2 from the wAC9 strain, and the TM3.GFP balancer on chromosome 3 (crossing scheme in Supplementary Figure S4).

Generating cDNA rescue constructs

Total RNA was extracted from the wild-type line Armenia14. The Dβ1 open reading frame was then PCR amplified using Dβ1_clone_F and Dβ1_clone_R primers (Supplementary Table S3) and cloned into pGEM-T Easy (Promega). Clones were sequenced, sub-cloned into the NotI site of the pUAST-attB vector (Bischof et al. 2007), and microinjected into attP40 (25709 w-) embryos. Successful transgenics were screened phenotypically for w+ eye color rescue, and molecularly using SV40 and 25709_External_R1 primers. Homozygotes were recovered by self-crossing the heterozygous offspring of injectees. Homozygotes were then crossed to incorporate the Dβ1Δex5 deletion and the TM3.GFP balancer on chromosome 3 for use in rescue experiments (rescue crosses outlined in Supplementary Figure S3). For the chromosome 3 driver CCAP-GAL4, a recombined CCAP-GAL4, Dβ1Δex5 line was generated.

Developmental mortality assays

First instar larvae were transferred from apple juice agar plates to standard semolina fly food, reared for 2 weeks, then scored for stage of mortality (50 per vial, 5 independent vials, n=250). Individuals that died during pupal stages P1-P11 or P12-P15 (Bainbridge and Bownes 1981) were scored as “pupal” and “eclosing,” respectively.

Wing morphology scoring

Adults were scored for wing morphology (n=250) using nomenclature established by Luan et al. (2006) as a guide. Unexpanded (UE) wings had an angle ≤90° between the distal and proximal portions of the folded wing, and fully expanded (FE) wings were completely straight or only slightly curled, resembling wild-type morphology. Partially expanded (PE) wings had intermediate phenotypes, often with unfolded costal elbows.

Longevity and fertility assays

Virgin males and females were collected over 48 hours then placed on standard semolina-based fly media (10 males and 10 females per vial, 4 vials per genotype, n=80). Vials were maintained horizontally at 25°C under a 12:12 hours light: dark cycle. Flies were transferred to new vials and the number of dead flies recorded every 2–3 days (or every 4–7 days late in the experiment for long-lived genotypes) until all were deceased. A Kaplan-Meier Survival Curve was constructed and significance was calculated using a Cox-Mantel test (P<0.05). To estimate fertility, these vials were then maintained for 2 weeks at 25°C and scored for the number of eclosing adult offspring relative to the number of surviving parental females present at the time of egg-laying (n=40). Significance was calculated for each timepoint using Tukey’s Honestly Significant Difference (HSD) pair-wise test (P<0.05).

Mating assays

Virgin males and females were collected and stored individually for 3 days. For single pair crosses, an individual male and female were placed in a standard semolina fly media vial with the stopper pushed down to ∼20mm from the surface of the food. This was repeated for all 4 possible combinations of wAC9 and Dβ1Δex5 flies (n=20 per combination). Pairs were allowed to mate for 3 days, then removed from the vial, and the presence of F1 offspring recorded 2 weeks later. For filmed assays, an individual male was gently aspirated without anesthesia into a mating chamber (12 × 4 mm) and allowed to acclimate for 3 minutes. A single female was then introduced and video recording initiated. Mating pairs were recorded using a Panasonic 3CCD Ultra-Compact Digital Palmcorder for 30 minutes or until copulation commenced. This was repeated with wAC9 males and Dβ1Δex5 males, each of which were only paired with a wAC9 female (n=20 per combination). Videos were inspected manually for courtship behavior and copulation. The Courtship Index was measured using only the first 10 minutes of the recording (or only the time prior to copulation for pairs that commenced copulation within 10 minutes) and was defined as the proportion of time males spent exhibiting the following courtship behaviors: wing extension, following, tapping, and attempted copulation. Significance was calculated using Student’s t-test (P<0.05).

Adult climbing assays

Three-day-old flies were transferred without anesthesia into clean empty vials, allowed to acclimate for 5 minutes, then tapped down 3 times on a fly pad to initiate the negative geotaxis response (Gargano et al. 2005) (10 males and 10 females per vial, 4 vials per genotype, n=80). The vials were recorded for 10 seconds from a distance of 30 cm on an iPhone 8 (Apple Inc.). QuickTime Player (Apple Inc.) was used to capture the frame occurring 6 seconds (180 frames) post-third tap down. JPEGs were then imported into Image J (Fiji, version 2.0.0, Schindelin et al. 2012) and marked with a horizontal threshold 35 mm from the bottom of the vials. The number of flies above this threshold was scored manually. Significance was calculated using Tukey’s HSD pair-wise test (P<0.05).

Larval movement assays

Basal larval movement was measured using a modified Wiggle Index (WI) Assay (Denecke et al. 2015). Briefly, second or third instar larvae were transferred into a 24-well plate (25 larvae per well, 4 wells per genotype, n=100) containing 200 µl 5% sucrose in distilled water. Thirty-second videos were taken at 0, 15, 30, 60, 90, and 120 minutes after the addition of larvae using a Panasonic 3CCD Ultra-Compact Digital Palmcorder. Raw videos were split into JPEG image sequences using the DVDVideoSoft Converter, and further processed as previously described (Denecke et al. 2015), generating heat maps and a numerical estimate of total motility (WI). To calculate relative movement, WI values were normalized to the wAC9 WI mean. Significance was calculated using Student’s t-test (P<0.05).

qRT-PCR

Quantitative real-time PCR (qRT-PCR) was used to measure transcript levels of Dβ1 and peptide hormones in Dβ1Δex5 and Dβ1Δex3 relative to the wAC9 control. For each of 4 biological samples, 10 pharate pupae or 10 1-day-old adults were collected in liquid nitrogen. RNA was isolated using TRIsure Reagent (Bioline) and the concentration measured using the Qubit Fluorometer. cDNA was synthesized from 1µg RNA using the GoScript Reverse Transcription Kit (Promega). qRT-PCR reactions for each biological sample were carried out in triplicate using a Quanti-fast SYBR Green PCR kit (Qiagen) on the CFX384 Touch Real-Time PCR Detection System (Bio-Rad). The amount of target RNA was normalized to the geometric mean of two endogenous controls: Rpl32 and CG13220 (Van Hiel et al. 2009). mRNA levels were compared between samples using the ΔΔCT method (Bustin et al. 2009) and significance were calculated using Student’s t-test (P<0.05). All primer sequences used are provided in Supplementary Table S3 and the Minimum Information for the publication of Quantitative Real-time PCR Experiments (MiQE) checklist is also provided (Supplementary Table S4).

Results

Dβ1 genomic deletions and Dβ1-TG4

Two Dβ1 genomic deletions were used in this study (Figure 1A). The larger deletion, Dβex5, removes 5484 bp spanning the region from the 5’UTR to exon 5 (Perry et al. 2021). Most of the receptor, up to and including transmembrane domain 3, is deleted. The smaller deletion, Dβ1Δex3, removes 4148 bp, removing the region from the 5’UTR to exon 3. The beginning of the ligand-binding domain (LBD), up to and including loop D is deleted. For both deletions, loss of Dβ1 transcripts in homozygotes was confirmed with qRT-PCR (Figure 1B). Unless otherwise specified, the larger deletion, Dβ1Δex5, was used in all subsequent experiments. A Dβ1 T2A-GAL4 strain was also generated in this study (Dβ1-TG4). This strain contains a Trojan GAL4 exon in intron 1, resulting in truncation of the Dβ1 gene product and expression of GAL4 in Dβ1’s native pattern (expression pattern provided in Supplementary Figure S5). It was thus used as both a null allele and a Dβ1-specific GAL4 driver (Diao and White, 2012).

Loss of Dβ1 decreases fitness and disrupts wing expansion

The deletion of Dβ1 incurs severe fitness costs. When reared in non-competitive conditions (i.e., in the absence of heterozygous siblings), Dβ1Δex5 homozygotes exhibited reduced survival to adulthood (83%) compared to wAC9 controls (98%) (Figure 2A). One distinctive stage of mortality was mid-eclosion (Figure 2B). For those flies that developed to adulthood, median longevity was shortened significantly from 40 days (wAC9) to 6 days (Dβex5) (P<0.001, Cox-Mantel test) (Figure 2C) and these adults failed to produce any viable offspring (P<0.001, Tukey’s HSD pair-wise test, Supplementary Table S5) (Figure 2D). Adults exhibited varying disruptions in wing expansion. Only 26% attained the FE morphology, whilst 19% presented with the intermediate PE wings and 55% had the severe UE (unexpanded) morphology (Figure 2, E and F).

Figure 2.

Figure 2

Dβ1 deletion reduces adult fitness. Dβ1Δex5 homozygotes have low fitness levels for several traits. This reduced fitness is replicated by neuron-specific somatic deletion of Dβ1 (elav-GAL4 CRISPR) and it is rescued by expression of Dβ1 cDNA in neurons (elav-GAL4 rescue) or in Dβ1-expressing cells (Dβ1-TG4 rescue). Results for Dβ1Δex5 homozygotes are shown alongside their background control, wAC9 (w; +; +). For elav-GAL4 CRISPR, elav-GAL4 rescue, and TG4 rescue, results for experimental animals are followed by results for parental controls. (A) Deaths during early adulthood (<20 days) and the “Eclosing” phase were most common for Dβ1Δex5 and other strains lacking Dβ1 (n = 250). (B) Depicts a fly arrested during the “Eclosing” phase. (C) Kaplan-Meier survival plots with black dashed lines marking median longevity. Median longevity was shortened significantly to 6 days in Dβ1Δex5. Shading represents 95% confidence intervals (n = 80, Cox-Mantel test). (D) Number of viable eggs laid per female per day, throughout lifespan. Error bars indicate SEM and P-values for each time point are provided in Supplementary Table S5 (n = 40, Student’s t-test). (E) Dβ1Δex5 and other strains lacking Dβ1 are afflicted with UE and PE wings, with fewer achieving FE wings (n = 250). (F) Images of representative individuals from each wing phenotype category.

We were able to replicate these phenotypes using pan-neuronal somatic CRISPR. In elav-GAL4 CRISPR mosaics (elav-GAL4/w; sgRNAs/+; UAS-Cas9/+), genomic deletion of Dβ1 is restricted to neurons by expressing UAS-Cas9 under the control of elav-GAL4 in the presence of two ubiquitously expressed Dβ1-targeting sgRNAs. Like Dβ1Δex5 flies, elav-GAL4 CRISPR mosaics exhibited 79% survival to adulthood (Figure 2A). Longevity and fertility were both affected, with median adult survival shortened significantly to 16 days (P<0.001, Cox-Mantel test) (Figure 2C) and no viable offspring produced in all but one vial (Figure 2D). Wing expansion was disrupted, although in slightly less severe proportions (50% FE, 13% PE, and 37% UE) (Figure 2, E and F). The general concordance of elav-GAL4 CRISPR and Dβ1Δex5 results indicate that loss of Dβ1 function in the neurons is responsible for the fitness and wing phenotypes observed.

We further verified that loss of Dβ1 function was responsible for these phenotypes using two approaches. First, expression of Dβ1 cDNA in a Dβ1 null background was sufficient to rescue all elements of the phenotype. Full phenotypic rescues were achieved when using the Dβ1-TG4 driver (“TG4 rescue”) (Figure 2, A, C, D, and E). When using the pan-neuronal elav-GAL4 driver (“elav-GAL4 rescue”), full rescues were achieved for longevity (Figure 2C) and wing expansion (Figure 2E), while partial rescues were observed for developmental mortality (Figure 2A) and production of viable offspring (Figure 2D). No overexpression phenotypes were observed. Second, Dβ1Δex5 failed to complement two independent Dβ1 mutant alleles: the Dβ1Δex3 genomic deletion and the Dβ1-TG4 truncation allele (Supplementary Figure S6).

Loss of Dβ1 affects locomotor performance

Having casually observed reduced movement in Dβ1Δex5 larvae, relative larval movement was quantified in second and third instar larvae using a modified WI Assay (Denecke et al. 2015). This plate-based assay involves filming larvae crawling in a shallow pool of sucrose solution. Thirty-second videos were converted to heat maps and numerical estimates of total motility (WI). Compared to the wAC9 control, Dβ1Δex5 homozygotes displayed 40 and 85% relative basal movement in second and third instar larvae, respectively. Both differences were highly significant (P<0.001, Student’s t-test) (Figure 3A).

Figure 3.

Figure 3

Dβ1 deletion affects larval crawling and adult climbing performance. Results for Dβ1Δex5 homozygotes are shown alongside their background control, wAC9 (w; +; +). For elav-GAL4 CRISPR, elav-GAL4 rescue, and TG4 rescue, results for experimental animals are followed by results for parental controls. (A) A Wiggle Index assay measuring relative larval movement reveals significantly reduced crawling in second (L2) and third (L3) instar larvae. Error bars indicate SEM (n = 100, ***P < 0.001, Student’s t-test). (B) Adult climbing performance was significantly low in all genotypes lacking Dβ1. Error bars indicate SEM (n = 80, **P < 0.01, *** P < 0.001, Tukey’s HSD pair-wise test).

In adults, severe overall movement deficiencies were observed, however, we chose to limit our investigation to climbing assays (Figure 3B). Individuals that climbed beyond a 35 mm checkpoint within six seconds were deemed to have a successful climbing response. Almost all Dβ1Δex5 homozygotes failed (2%), whilst 93% of wAC9 controls were successful (P<0.001, Tukey’s HSD pair-wise test). elav-GAL4 CRISPR mosaics were also unable to climb (2%) (P<0.001). The climbing response was only partially rescued (20%) in the elav-GAL4 rescue, however, this was a statistically significant improvement in performance compared to non-rescued parental controls (both 0%, P<0.001). Full rescue was achieved using the Dβ1-TG4 driver (P<0.001).

Males lacking Dβ1 spend less time courting and fail to initiate copulation

Given the failure to produce offspring, and the reduced locomotor performance observed in previous assays, we were curious if Dβ1Δex5 adults exhibited normal mating behaviors. First, we crossed individual Dβ1Δex5 and wAC9 males and females in all four possible combinations and recorded the presence of F1 offspring (Figure 4A). Offspring were observed in almost all crosses between wAC9 males and Dβ1Δex5 females, indicating Dβ1Δex5 females are fertile. Meanwhile, crosses involving Dβ1Δex5 males never gave rise to offspring, indicating a male issue. Males were therefore investigated further in filmed 30-minute mating assays. All pairs consisting of a wAC9 male and wAC9 female initiated copulation within the 30-minute recording, while no copulation was observed when wAC9 females were paired with Dβ1Δex5 males (Figure 4B). The first 10 minutes of the recording was also used to calculate a Courtship Index, defined as the proportion of time males spent exhibiting any of the following behaviors: wing extension, following, tapping, and attempted copulation (Figure 4C). On average, Dβ1Δex5 males spent only 2% of their time courting, compared to 39% in wAC9 males (P<0.001, Student’s t-test). The only behaviors observed in Dβ1Δex5 males were weak attempts at wing extension and brief bouts of following.

Figure 4.

Figure 4

Males lacking Dβ1 fail to mate. (A) For crosses between single mating pairs, the presence of F1 offspring was recorded. The male genotype is indicated above and the female genotype is indicated below. Crosses involving Dβ1Δex5 males gave never gave rise to offspring, while Dβ1Δex5 females were shown to be fertile (n = 20 pairs per mating combination). (B) Percentage of pairs that initiated copulation within 30 minutes of a female being introduced. The female genotype is indicated above and the male genotype is indicated below. Dβ1Δex5 males do not initiate copulation (n = 20 pairs per mating combination). (C) Dβ1Δex5 males spend significantly less time courting females than wAC9 control (w; +; +) males. The female genotype is indicated above and the male genotype is indicated below. Error bars indicate SEM (n = 20 pairs per mating combination, ***P < 0.001, Student’s t-test).

Pinpointing a neuronal subset that requires Dβ1

Having observed disruptions in eclosion and wing expansion, we chose to investigate the role of Dβ1 in the CCAP neurons. These neurons secrete the neuropeptide hormones CCAP, required for eclosion (and other ecdyses) (Park et al. 2003), and bursicon, required for wing expansion and cuticle tanning (Luan et al. 2006).

We used somatic CRISPR to generate mosaics with a genomic deletion of Dβ1 specifically in CCAP neurons (“CCAP-GAL4 CRISPR”) of which there are approximately 27 pairs in the larval brain (Park et al. 2003; Karsai et al. 2013) (Figure 5A). This CCAP-specific deletion of Dβ1 was sufficient to cause the wing expansion phenotype only (Figure 5B). Wing morphology was severely affected, with 60%, 29%, and 12% displaying the UE, PE, and FE phenotypes, respectively. Meanwhile, developmental mortality, adult longevity, fertility, and climbing performance were comparable to the parental controls (Supplementary Figure S7). We performed the same somatic CRISPR experiment using the more restrictive bursicon-GAL4 driver. Bursicon has a later temporal expression pattern than CCAP (Kim et al. 2006; Di Cara and King-Jones 2013), and its expression is limited to a subset of the CCAP neurons (Kim et al. 2006; Luan et al. 2006). No phenotypes were observed, so we extracted genomic DNA from these bursicon-GAL4 CRISPR flies. We were not able to detect any CRISPR editing in pooled samples (Supplementary Figure S8), so we concluded that the lack of phenotype was actually due to inefficient CRISPR in these flies and they were not examined further.

Figure 5.

Figure 5

Dβ1 is required in CCAP neurons for bursicon-driven wing expansion. (A) Cartoon depicting CCAP neurons in a larval and pupal brain (modified from Veverytsa and Allan 2012; Moris-Sanz et al. 2015). (B) Somatic CRISPR deletion of Dβ1 specifically in CCAP neurons is sufficient to impair wing expansion (n = 250). Results for experimental animals are followed by results for parental controls. (C–H) Expression of Dβ1 driven by CCAP-GAL4 or bursicon-GAL4, in a Dβ1Δex5 background, can fully or partially rescue some Dβ1Δex5 phenotypes. Results for each rescue are followed by results for nonrescued parental controls. (C) CCAP-GAL4 rescue and bursicon-GAL4 rescue largely eliminate developmental mortality (n = 250). Given the high rate of larval mortality observed in the parental control w; +; CCAP-GAL4, Dβ1Δex5, this strain is absent from subsequent adult experiments. (D) Kaplan-Meier survival plot with black dashed lines marking median longevity. CCAP-GAL4 and bursicon-GAL4 rescue median longevity to 29 and 28 days, respectively. This is significant compared to parental controls, but less efficient than restoring Dβ1 expression in all Dβ1-expressing cells (Figure 2). Shading represents 95% confidence intervals (n = 80, P < 0.001, Cox-Mantel test). (E) Number of viable eggs laid per female per day, throughout lifespan. CCAP-GAL4 rescue and bursicon-GAL4 rescue do not significantly improve egg counts. Error bars indicate SEM (n = 40, Student’s t-test performed for each time point). (F) CCAP-GAL4 rescue and bursicon-GAL4 rescue almost completely restore normal wing expansion (n = 250). UE, unexpanded; PE, partially expanded; FE, fully expanded. (G) CCAP-GAL4 rescue and bursicon-GAL4 rescue partially but significantly improved adult climbing performance. Error bars indicate SEM (n = 80, ***P < 0.001, Tukey’s HSD pair-wise test). (H) qRT-PCR indicates 21-fold reduction in bursicon transcripts and 4-fold reduction in partner of bursicon transcripts in newly-eclosed Dβ1Δex5 homozygous adults. A nonsignificant reduction was observed for CCAP transcripts in pharate pupae. Error bars indicate SD (n = 4 biological replicates, *P < 0.05, **P < 0.01, Student’s t-test). (I) Male and female Dβ1Δex5 homozygotes have a slight reduction in cuticle pigmentation compared to wAC9 controls.

Having evidence that Dβ1 plays a key role in CCAP neurons, we moved on to consider whether expression of Dβ1 in CCAP neurons alone is sufficient for viability. To test this, CCAP-GAL4 and bursicon-GAL4 were used to drive expression of UAS-Dβ1 cDNA in a Dβ1Δex5 homozygous background. For CCAP-GAL4, full rescue was achieved for developmental mortality (Figure 5C) and wing morphology (Figure 5F). This suggests Dβ1 is required almost exclusively in CCAP neurons to fulfill its roles in wing expansion and survival through development. Partial rescue was achieved for CCAP-GAL4 for adult longevity (28 days) (Figure 5D), and to a lesser extent, fertility (Figure 5E) and climbing performance (14%) (Figure 5G), indicating Dβ1 also plays critical roles in other neuronal subsets affecting aging, mating, and climbing ability.

bursicon-driven expression of UAS-Dβ1 was slightly less efficient in its capacity to rescue these phenotypes. For bursicon-GAL4, full rescue was achieved only for wing morphology (Figure 5F). Partial rescue was achieved for bursicon-GAL4 for developmental mortality (86%) (Figure 5C) and adult longevity (29 days) (Figure 5D) and to a lesser extent, fertility (Figure 5E) and climbing performance (4%) (Figure 5G). No phenotypes were observed from overexpression of Dβ1 with CCAP-GAL4 or bursicon-GAL4.

To test whether loss of Dβ1 disrupts peptide hormone expression in CCAP neurons, we used qRT-PCR to measure hormone transcripts in Dβ1Δex5 relative to the wAC9 control. Transcripts were measured in a life stage where the respective hormone is known to be active (Di Cara and King-Jones 2013); CCAP transcripts where measured in pharate pupae, whilst bursicon and its heterodimer partner, partner of bursicon, were measured in newly eclosed adults. A significant 21-fold reduction in bursicon transcripts was detected in Dβ1Δex5 (P<0.01), as was a significant 4-fold reduction in partner of bursicon (P<0.05) (Figure 5H). A small reduction in CCAP was also detected in Dβ1Δex5 however this was not statistically significant due to variation between biological replicates.

Finally, given that bursicon plays an important role in cuticle tanning, the cuticles of Dβ1Δex5 and wAC9 adults were also compared, however only a subtle difference in pigmentation was observed (Figure 5I).

Discussion

Understanding nAChR biology is a vital step in dissecting the cholinergic circuits of the insect brain. It also provides valuable insight into how nAChR-targeting chemicals used for pest control may affect insect development, physiology, and behavior. In the present study, we provide the first functional description of a deletion of the Dβ1 nAChR subunit, a known insecticide target, and the most highly expressed nAChR subunit in the Drosophila brain. As expected, loss of Dβ1 was highly disruptive, with individuals having a shortened lifespan, reduced mating success, limited larval and adult movement, and unexpanded wings. This reduction in viability was partially attributed to CCAP neurons requiring Dβ1 expression and linked to loss of the peptide hormone bursicon. These are by far the most severe pleiotropic consequences described for an nAChR mutant so far (Somers et al. 2017; Homem et al. 2020, Perry et al. 2021). Deletion mutants for nine of the ten Drosophila nAChR subunits have now been described, and while not all the traits examined here have been analyzed in these other mutants, no subunits aside from Dβ1 have been observed with a wing expansion phenotype (Perry et al. 2021).

Our findings follow recent work on a resistance-conferring mutation in Dβ1, the R81T amino acid replacement (Homem et al. 2020). Homem and colleagues describe similar fitness costs in their CRISPR-generated Dβ1R81T allele. Like the Dβ1Δex5 deletion, Dβ1R81T mutant flies were infertile and had poor larval crawling performance. Reduced longevity and climbing performance were also described, albeit with sex-specific differences that we did not observe in our own lines. Notably, the authors did not describe any disruptions in wing expansion in either sex, but it is important to keep in mind the nature of the alleles when comparing these sets of results. Our deletions, Dβ1Δex5 and Dβ1Δex3, result in complete loss of the Dβ1 subunit. In the Dβ1-expressing neurons, this absence could simply result in loss of cholinergic signal, but there is evidence that the absence of one subunit can also alter the levels of other nAChR subunits at the level of transcription (Perry et al. 2021). In contrast, the R81T mutation does not result in loss of the subunit (Homem et al. 2020) and there is evidence that this amino acid replacement has only a subtle effect on the binding affinity of acetylcholine (Shimomura et al. 2006; Ihara et al. 2020). Thus, subsets of neurons involved in different biological processes and behaviors may have different vulnerability to a simple reduction in neuron firing (as in Dβ1R81T) versus complete loss of a subunit with possible nAChR compensation (as in Dβ1Δex5).

Using the somatic CRISPR-Cas9 system (Meltzer et al. 2019) and GAL4-UAS rescue system, we identified CCAP neurons as a neuronal subset where Dβ1 expression was required for robust wing expansion. Disrupted wing expansion was observed at high frequency in our CCAP-GAL4 CRISPR mosaics, and this phenotype was fully rescued in CCAP-GAL4 and bursicon-GAL4 driven rescues. We also observed a mild reduction in tanning of the cuticle in Dβ1Δex5 mutants. Furthermore, inspection of available single-cell RNA-seq datasets reveal co-expression of Dβ1 with CCAP, bursicon and partner of bursicon in both the larval brain (Ravenscroft et al. 2020) and the adult ventral nerve cord (Allen et al. 2020) (Supplementary Figure S9). Bursicon has long been documented as the wing expansion and cuticular tanning hormone (Fraenkel and Hsiao 1965), and the molting hormone cascade that culminates in bursicon’s release from CCAP neurons is well-studied (Di Cara and King-Jones 2013). So how might Dβ1-containing nAChRs fit into this hormone cascade? The delayed release of bursicon is regulated by excitation of the CCAP neurons (Luan et al. 2006) and, to our knowledge, the neurotransmitter/s responsible for this excitation have not been identified. We propose the signal may be cholinergic, requiring Dβ1-containing nAChRs. However, this study did not examine the cholinergic signal and it will be important to test the electrophysiological response of CCAP neurons to acetylcholine in wild-type flies and Dβ1 mutants to determine what current remains in the absence of Dβ1-containing nAChRs. An alternative hypothesis is that Dβ1-containing nAChRs are required in CCAP neurons during development such that loss of Dβ1 alters the connectivity and functionality of CCAP neurons. This hypothesis is also likely to explain the transcript-level loss of CCAP, bursicon, and partner of bursicon we observed in Dβ1Δex5. It is also possible that altered expression of other nAChR subunits present in CCAP neurons may be contributing to the phenotypic effects of loss of Dβ1 (Perry et al. 2021). Based on single-cell RNA-seq data, all subunits except Dβ3 are detected in CCAP neurons, (Allen et al. 2020, Ravenscroft et al. 2020). Ultimately, our results firmly place Dβ1-containing receptors as a key component of CCAP neuron signaling, however, investigation to further tease apart when and how Dβ1 is required will be of value.

Phenotypes that showed unsuccessful rescue (fertility) or partial rescue (longevity and climbing performance) in our CCAP rescue experiments indicate important roles for Dβ1 subunits in other neurons outside the CCAP subset. The observed reduction in Dβ1Δex5 male courtship, and the subsequent failure to mate, could indicate a role for Dβ1-containing receptors in neurons regulating mating behavior. Indeed, some of these neurons have already been identified as cholinergic (Acebes et al. 2004; Schinaman et al. 2014; Somers et al. 2017; Ishimoto and Kamikouchi 2020). Alternatively, any mating issues could be a side effect of impaired movement. Aside from poor climbing performance, we observed reduced general movement, and it is possible Dβ1-containing nAChRs are required in cholinergic muscle control (Hsu and Bhandawat 2016; Malloy et al. 2019). An approach similar to that used here, to specifically delete the Dβ1 gene in neural circuits controlling male courtship, would provide insight into whether the mating issues observed are due to altered signaling in these courtship circuits, or due to a more general impact on movement.

While our experiments have yielded significant insights into roles of Dβ1, there remains considerable work to be performed on other circuits and with other nAChR subunits if we are to gain a complete picture of nAChR function in insect nervous systems. Recent developments in heterologous nAChR expression (Ihara et al. 2020) and single-cell sequencing (Croset et al. 2018; Brunet Avalos et al. 2019; Allen et al. 2020) will help accelerate this research, determining which subunits form functional subtypes, and whether those subunit combinations are natively expressed in the same neurons. Moving forward, simultaneous disruption of multiple subunits may also be required to unearth certain phenotypes. This would be useful in the identification of the nAChR α subunits co-assembling with Dβ1 to influence wing expansion. As we have seen here, this work highlights some of the reasons why nAChR-targeting insecticides became such a commercial success, yet also hints at why this insecticide class has run into issues relating to collateral damage to beneficial insect populations (Goñalons and Farina 2015; Wu-Smart and Spivak 2016; Woodcock et al. 2017; Crall et al. 2018; Sánchez-Bayo and Wyckhuys 2019; Martelli et al. 2020). On one hand, the severe fitness costs we have described may have safeguarded growers against widespread insecticide resistance evolving in pest species. On the other hand, our findings provide insight into how sublethal doses might be disrupting specific neuronal pathways in nonpest species. Ongoing efforts to understand native roles of insect nAChRs will be vital for refining pest control strategies and predicting the behavioral and physiological impacts on species living in insecticide-contaminated environments.

Acknowledgments

Fly strains were sourced from the Bloomington Drosophila Stock Centre and the Australian Drosophila Biomedical Research Support Facility provided quarantine facilities for imported fly strains.

Funding

Funding was provided through an Australian Research Council Discovery Project awarded to P.B. (DP160100332), and an Australian Postgraduate Award, and Dame Margaret Blackwood Soroptimist Scholarship awarded to D.C.

Conflicts of interests: None declared.

Data availability

The authors affirm that all data necessary for confirming the conclusions of this article are represented fully within the article and its tables and figures. All fly strains are available upon request. Supplementary files have been uploaded to figshare: https://doi.org/10.25386/genetics.14593902.

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Data Availability Statement

The authors affirm that all data necessary for confirming the conclusions of this article are represented fully within the article and its tables and figures. All fly strains are available upon request. Supplementary files have been uploaded to figshare: https://doi.org/10.25386/genetics.14593902.


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