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. 2020 Jul 16;216(1):135–144. doi: 10.1534/genetics.120.303475

A New Role for Neuropeptide F Signaling in Controlling Developmental Timing and Body Size in Drosophila melanogaster

Jade R Kannangara *, Michelle A Henstridge *, Linda M Parsons †, Shu Kondo ‡, Christen K Mirth *,1,✉, Coral G Warr †,1,2
PMCID: PMC7463290  PMID: 32675276

Abstract

As juvenile animals grow, their behavior, physiology, and development need to be matched to environmental conditions to ensure they survive to adulthood. However, we know little about how behavior and physiology are integrated with development to achieve this outcome. Neuropeptides are prime candidates for achieving this due to their well-known signaling functions in controlling many aspects of behavior, physiology, and development in response to environmental cues. In the growing Drosophila larva, while several neuropeptides have been shown to regulate feeding behavior, and a handful to regulate growth, it is unclear if any of these play a global role in coordinating feeding behavior with developmental programs. Here, we demonstrate that Neuropeptide F Receptor (NPFR), best studied as a conserved regulator of feeding behavior from insects to mammals, also regulates development in Drosophila. Knocking down NPFR in the prothoracic gland, which produces the steroid hormone ecdysone, generates developmental delay and an extended feeding period, resulting in increased body size. We show that these effects are due to decreased ecdysone production, as these animals have reduced expression of ecdysone biosynthesis genes and lower ecdysone titers. Moreover, these phenotypes can be rescued by feeding larvae food supplemented with ecdysone. Further, we show that NPFR negatively regulates the insulin signaling pathway in the prothoracic gland to achieve these effects. Taken together, our data demonstrate that NPFR signaling plays a key role in regulating animal development, and may, thus, play a global role in integrating feeding behavior and development in Drosophila.

Keywords: neuropeptide F receptor, neuropeptide F, insulin signaling, ecdysone, developmental timing, Drosophila melanogaster

WHEN faced with variation in the quantity and quality of the diet, young animals must adjust their feeding behavior, metabolism, growth, and developmental time. Failure to do so has profound consequences on their ability to survive to adulthood and to resist future stress. Extensive research into the regulation of food intake has uncovered a handful of neuropeptides that mediate changes in feeding behavior in response to diet, including the highly conserved Neuropeptide F signaling pathway (Beck 2001; Williams et al. 2001; Garczynski et al. 2002; Wu et al. 2003). Developmental processes, such as growth and developmental time to adulthood, are controlled through the action of the conserved insulin and steroid hormone signaling pathways (Caldwell et al. 2005; Colombani et al. 2005; Mirth et al. 2005). However, we know little about the extent to which feeding behavior and developmental processes are coordinated, and the molecular mechanisms necessary for this coordination.

The fruit fly, Drosophila melanogaster, provides an excellent model in which to study the molecular mechanisms that integrate feeding behavior with developmental processes. Drosophila development proceeds through three larval stages (instars), after which the animal initiates pupariation and metamorphosis to become an adult. The timing of the transitions between these developmental stages is regulated by a series of precisely timed pulses of the steroid hormone, ecdysone, produced and secreted by the prothoracic gland (PG; Nijhout et al. 2014). Because these insects grow primarily during the larval stages, ecdysone dictates the length of the growth period, ceasing growth once metamorphosis begins (Caldwell et al. 2005; Mirth et al. 2005; Nijhout et al. 2014). In this way, ecdysone determines final adult size.

The PG produces and secretes ecdysone in response to various environmental cues, such as the day–night cycle, nutrition, and tissue damage (Mirth et al. 2005; McBrayer et al. 2007; Koyama et al. 2014; Jaszczak et al. 2016). These external cues are communicated to the PG via the action of a number of secreted peptides. Nutritional signals are particularly important, and are communicated throughout the body via the insulin signaling pathway. When larvae are well fed, they secrete insulin-like peptides (Dilps) into the bloodstream (Brogiolo et al. 2001). In the Drosophila PG the insulin receptor (InR) is activated by the Dilps, which, in turn, leads to the activation of ecdysone biosynthesis genes and therefore the production of ecdysone (Caldwell et al. 2005; Colombani et al. 2005; Mirth et al. 2005). Starvation early in the third larval instar delays the onset of metamorphosis by delaying the timing of an early ecdysone pulse (Caldwell et al. 2005; Colombani et al. 2005; Mirth et al. 2005; Shingleton et al. 2005). Later in the third larval instar, starvation accelerates developmental timing (Mirth et al. 2005; Stieper et al. 2008), presumably by accelerating the production of at least one of the later ecdysone pulses. This highlights how the effects of nutrition change growth outcomes over developmental time. As well as regulating development, the quantity and quality of nutrients in the diet also cause larvae to alter both the amount and the quality of foods they consume (Rodrigues et al. 2015; Almeida de Carvalho and Mirth 2017). Several peptide hormones and neuropeptides have been shown to regulate different aspects of feeding behavior in the fly (Wang and Wang 2019). Among these, Neuropeptide F (NPF) signaling increases feeding rates and affects food choice in response to poor food quality (Wu et al. 2003; Wu et al. 2005). The mammalian homolog of NPF, Neuropeptide Y, also regulates feeding behavior in response to food quality (Brown et al. 1999; Mercer et al. 2011). To guarantee that the animal survives, these changes in feeding behavior must be appropriate to the changes needed in the different stages of animal development. We therefore wondered if NPF signaling may act as a global coordinator of development and feeding behavior in response to nutritional signals.

Here, we show that the NPF receptor (NPFR) regulates development in Drosophila by regulating the production of ecdysone in the PG. We further show that NPFR signaling exerts its effects on developmental timing and body size by interacting with the insulin signaling pathway in the PG, revealing that it acts as a previously undescribed regulator of insulin signaling in this gland. Our data demonstrate that NPF signaling, well known for regulating feeding behavior across species, also plays a key role in regulating animal development by affecting the production of developmental hormones.

Materials and Methods

Drosophila stocks

The following stocks were used: w1118 (BL5905), NPF-Gal4 (BL25682), from the Bloomington Drosophila Stock Center, UAS-NPFR RNAi (v9605), UAS-NPFR RNAi (v107663), UAS-NPF RNAi (108772) from the Vienna Drosophila Resource Center, NPFRSK8 mutant (Kondo and Ueda 2013; Ameku et al. 2018), phm-Gal4-22, UAS-mCD8::GFP and UAS-dicerII; phm-Gal4-22, gifts from Michael O’Connor, University of Minnesota, Minneapolis (Ono et al. 2006). All flies were maintained at 25° on fly media containing, per liter: 7.14 g potassium tartrate, 0.45 g calcium chloride, 4.76 g agar, 10.71 g yeast, 47.62 g dextrose, 23.81 g raw sugar, 59.52 g semolina, 7.14 ml Nipagen (10% in ethanol), and 3.57 ml propionic acid.

Developmental timing assays and body size analysis

Parental flies were allowed to lay eggs on 25 mm apple juice agar plates for 3–4 hr; 24 hr later, 15 L1 larvae were picked into standard food vials. A total of 10 replicates were collected from each cross. Time to pupariation of the F1 offspring were scored every 8 hr. Larvae for all experiments were raised inside an insulated, moist chamber at 25° in the dark. As a proxy for body size, following their eclosion photos of the pupal cases from the developmental timing assays were taken using a light compound microscope at 2.5× magnification. Pupal case length was measured using Fiji.

Immunocytochemistry

For PG size studies, wandering larvae from each genotype were collected, and anterior halves of the larvae were dissected and fixed for 30 min in 4% formaldehyde in PBTx [0.01% Triton-X in phosphate-buffered saline (PBS)]. Samples were washed four times over 1 hr in PBTx, and then incubated in 50 μl RNAase for 20 min. Samples were incubated in 4′,6-diamidino-2-phenylindole (DAPI; 1:400 in PBTx) for 2 min, and washed in PBTx. Samples were stored in Vectashield (Vector Laboratories), and PGs were dissected under a light compound microscope in PBS. Dissected PGs were mounted onto a slide and were visualized using confocal microscopy (Olympus CV1000). Measurements of PG area were quantified using Fiji. For anti-FoxO staining, larvae were staged at L3 and at the appropriate time points, dissected and fixed in 4% formaldehyde in PBS for 45 min at room temperature. Samples were then washed in PBTx and blocked for 30 min in 5% goat serum in PBTx and rabbit anti-FoxO (1:500; a gift from Pierre Leopold,) was added to 5% goat serum in PBTx. Samples were allowed to incubate at 4° overnight. We then washed the samples in PBTx, and anti-rabbit Alexa 488 (1:500; Invitrogen) in 5% goat serum in PBTx was added in the dark and allowed to incubate for 1.5 hr at room temperature. Samples were then washed and incubated in DAPI (1:400; PBTx) for 2 min. After washing in PBTx again, we added Phalloidin (1:1000; PBTx) to the samples and allowed them to incubate at room temperature for 20 min. Samples were washed and stored in Vectashield (Vector Laboratories) before further dissection onto poly-l-lysine-coated coverslips and analyzed using confocal microscopy (CV1000; Olympus).

Growth rate

Parental flies were allowed to lay on 25 mm apple juice agar plates for 3–4 hr. Parent flies were removed and the eggs were allowed to develop for a further 24 hr. Around 15–20 L1 larvae were picked into standard food vials and were allowed to develop for a further 72 hr; 6–8 replicates were picked for each genotype. Individual larvae were then floated in 20% sucrose to retrieve them from the vials, and one replicate was weighed using a microbalance (Mettler Toledo) each morning and evening until the larvae started to pupariate. Weight over time was recorded and analyzed using Prism 7.

Ecdysone feeding

To make 20-hydroxyecdysone (20E) food, a stock solution of 10 mg/ml of 20E (Cayman Chemical) was dissolved in 96% EtOH. To reach a final concentration of 0.15 mg/ml, 15 μl of the stock solution was added per 1 g blended fly medium. For the control food, 96% EtOH was used without 20E addition. A total of 10 young L3 larvae were picked into the vials and allowed to feed ad libitum. Time to pupariation was measured every 8 hr; 10 replicates were used per genotype.

Quantitative PCR

Total RNA was extracted from the anterior halves of 10–15 larvae using TRIsure (Bioline). After DNase treatment, total RNA concentration was quantified and ≤5 μg of total RNA was converted to cDNA using a 1:1 mix of oligo DT and random hexamer primers, and reverse transcriptase (Bioline). Quantitative PCR (qPCR) was performed using SYBR Green PCR MasterMix (Bioline). Primer sequences for phm, dib, and rpl23 were borrowed from McBrayer et al. (2007). Sequences for e74b are as follows: e74B (F- 5′ CGGAACATATGGAATCGCAGTG, R- 5′ CATTGATTGTGGTTCCTCGCTG 3′).

Ecdysone titer quantification

Larvae were synchronized by collecting newly ecdysed L3 larvae every 2 hr. A sample of 8–10 larvae was weighed on a microbalance (Mettler Toledo), and then preserved in methanol. Prior to assaying, the samples were homogenized and centrifuged, and the resulting methanol supernatant was dried. Samples were resuspended in 50 μl of enzyme immunoassay (EIA) buffer (0.4 M NaCl, 1 mM EDTA, 0.1% bovine serum albumin, and 0.01% sodium azide in 0.1 M phosphate buffer). 20E EIA antiserum and 20E acetylcholinesterase tracer were purchased from Cayman Chemicals.

Immunoblotting

Five L3 larvae were homogenized in 80 μl of lysis buffer [50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 2.5 mM EDTA, 0.2% Triton X, 5% glycerol, complete EDTA-free protease inhibitor cocktail (Roche)], and spun at 500 × g for 5 min at 4°. Reducing buffer was added to all samples before boiling and separation by SDS-PAGE (any kDa TGX, Bio-Rad) followed by transfer onto an Immobilon-P membrane (Millipore). Membranes were probed with either 1:1000 antiphosphorylated Drosophila Akt (4054S; Cell Signaling), or 1:1,000,000 anti-α-tubulin (Sigma, B-5-1-2), washed, and incubated with HRP-conjugated secondary antibody (1:10,000, Southern Biotech). Immunoblots were developed using ECL prime (GE healthcare), and imaged using a chemiluminescence detector (Vilber Lourmat). pAkt blot images were quantified using Fiji and differences between genotypes determined by unpaired t-tests from six biological replicates.

Nutritional plasticity

Food of varied caloric concentrations was made by diluting our standard food (SF) as described above with 0.5% agar (Gelita). The food concentrations used were 0.1× (10% SF, 90% agar), 0.25× (25% SF, 75% agar), 0.5× (50% SF, 50% agar), and 1× (100% SF). Eggs were picked onto these diluted foods and pupal weight was measured using a microbalance (Mettler Toldeo). For each genotype, at least 10 replicates of 15 larvae were raised on each food concentration. Differences in genotypes was determined by linear regression analysis using GraphPad Prism.

Quantification of food intake

Newly molted third-instar larvae were transferred to freshly dyed food (4.5% blue food dye) and allowed to feed for 1 hr. After feeding, larvae were removed from food using 20% sucrose solution, washed in distilled water and dried. Replicates of 10 larvae were homogenized in 80 μl of cold methanol and centrifuged for 10 min at 4°. A 60 μl aliquot of supernatant from each sample was analyzed in a spectrophotometer at 600 nm. As standards, a twofold dilution series of food dye, starting at a concentration of 4 μl dye/ml methanol was used; 5–6 biological replicates were analyzed per genotype.

Data availability

All strains are available upon request. We affirm that all data necessary for confirming conclusions of this article are present within the article and figures. Supplemental material available at figshare: https://doi.org/10.25386/genetics.12661226.

Results and Discussion

NPFR signaling regulates developmental timing

Our aim was to determine whether the NPF signaling pathway, known for controlling feeding behavior, also acts to alter development in response to nutrition. Because the known effects of nutrition on developmental time are controlled by ecdysone production in the PG, we hypothesized that NPF would act on its receptor, NFPR, on the PG cells. We therefore knocked down NPFR specifically in the PG to determine whether it affects development time. To do this, we used the PG-specific driver phantom (phm)-Gal4 to drive expression of RNAi constructs for NPFR, as well as the expression of dicer II (dcrII) (phm > NPFRi; dcrII) to enhance the RNAi knockdown (Tomari and Zamore 2005). We found that, when we knocked down NPFR in the PG with the v9605 RNAi line, we observed a significant delay to pupariation of about 35 hr (Figure 1A, P < 0.0001). With a second NPFR RNAi line (v107663), we observed a significant delay only when compared to the UAS-NPFR RNAi parental control, and not the phm-Gal4 parental control (Figure 1B, P = 0.0044). Given that the NPFR RNAi v9605 line generated the more severe phenotype, this RNAi line was used for all future tissue-specific experiments. To further verify that NPFR regulates developmental timing, we tested an NPFR loss of function mutant strain (NPFRSK8; Ameku et al. 2018). The NPFRSK8 mutant larvae also displayed a significant delay in time to pupariation compared to a heterozygous control (∼15 hr; Figure 1C, P = 0.0029).

Figure 1.

Figure 1

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NPFR regulates developmental timing. (A) Knockdown of NFPR (using UAS-NPFR RNAi v9605) specifically in the PG (using phm-Gal4>dcrII) results in a significant delay in time to pupariation compared to controls (P < 0.0001). (B) Knockdown of NPFR specifically in the PG using a second, independent RNAi line (UAS-NPFR RNAi v107663) also results in a significant delay in time to pupariation (P = 0.0044). (C) NPFR null mutants (NPFRSK8) have a significant delay in time to pupariation (P = 0.0029). (D) Knockdown of NPF in NPF-expressing neurons (using NPF-Gal4) results in a significant developmental delay (P < 0.0001). hAEL =hr after egg lay. Error bars represent ±1 SEM for all graphs. In each experiment, genotypes indicated with the same letter are not significantly different from one another, while genotypes with different letters are significantly different, as measured by ANOVA (A, B and D) or a pairwise t-test (B). Each point represents a biological replicate of 15–20 animals.

NPFR is a G-protein coupled receptor that is activated by the neuropeptide NPF (Garczynski et al. 2002). To further test the role of NPFR signaling in developmental timing, we knocked down NPF specifically in the NPF-producing neurons using NPF-Gal4. We found that these larvae exhibited a 10-hr delay in time to pupariation when compared to controls (Figure 1D, P < 0.0001). Together, these data suggest that NPF acts on NPFR on the PG cells to regulate developmental timing.

NPFR regulates the production of ecdysone in the prothoracic gland

NPFR could alter developmental timing by regulating either PG function or PG development. To test the latter possibility, we dissected PGs from phm > NPFRi; dcrII wandering larvae and measured their size. PG size was not significantly different between phm > NPFRi; dcrII and control larvae (Figure S1, P = 0.528). This suggests that NPFR signaling is more likely to affect the function of the PG, although these data do not completely rule out effects on development.

We next tested whether NPFR signaling regulates the primary function of the PG—to produce ecdysone. We reasoned that, if NPFR acts in the PG to regulate ecdysone production, then feeding phm > NPFRi; dcrII larvae with ecdysone should rescue the developmental delay. Consistent with this prediction, supplying 20E—the active form of ecdysone—to phm > NPFRi; dcrII larvae rescued the developmental delay (Figure 2A, P < 0.0001), inducing even faster time to pupariation than one of the two controls. When we quantified ecdysone titers, we found that phm > NPFRi; dcrII animals produced significantly less ecdysone later in the third instar, between 32 and 56 hr after the third-instar molt, when compared to controls (Figure 2B).

Figure 2.

Figure 2

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NPFR regulates the production of ecdysone in the prothoracic gland. (A) Time to pupariation was measured for phm > NPFRi; dcrII larvae fed on either food supplemented with 96% EtOH (gray) or 20-hydroxyecdysone (20E) (black). Supplying phm > NPFRi; dcrII larvae with 20E is able to completely rescue the developmental delay seen when phm > NPFRi; dcrII animals are fed on control food (P < 0.0001). Each point represents a biological replicate of 15–20 animals. Analyses were performed both between genotypes and across treatments. (B) phm > NPFRi; dcrII animals have an overall reduction in ecdysone titer compared to parental controls during the late third instar. hAL3E = hr after L3 ecdysis. Five biologically independent replicates of 8–10 larvae were measured for each time point. (C–E) Relative expression of two different ecdysone biosynthetic genes, (C) phm, and (D) dib, and of the ecdysone response gene (E) e74B, is reduced in phm > NPFRi; dcrII animals as determined by quantitative PCR. In panels C–E, values were normalized using an internal control, Rpl23. hAL3E = hr after L3 ecdysis. Expression level of each gene was standardized by fixing the values at 32 hr in NPFRi/+ as one in all panels. Approximately 8–15 larvae were used for each sample, and five biologically independent samples for each time point. Error bars represent ±1 SEM for all experiments. In each experiment, genotypes sharing the same letter indicate that they are not significantly different from one another, while genotypes with contrasting letters indicate that they are statistically different (measured by an ANOVA).

This reduction in total ecdysone concentration could be due to a defect in either its biosynthesis or in its secretion. To distinguish between these two possibilities, we quantified the expression levels of two CYP450 ecdysone biosynthetic genes, phm and disembodied (dib). The mRNA expression levels of these two enzymes are well-established as reliable proxies for ecdysone biosynthesis (Colombani et al. 2005; McBrayer et al. 2007; Koyama et al. 2014). Additionally, the expression of an ecdysone response gene, ecdysone-induced protein 74EF (e74B), was quantified as a readout of ecdysone signaling activity. When NPFR was knocked down in the PG, there was an overall reduction in phm and dib between 32 and 56 hr after the third-instar molt compared to controls (Figure 2, C and D). Further, e74b expression was reduced compared to controls (Figure 2E), demonstrating lower levels of ecdysone signaling activity in larvae where NPFR was knocked down in the PG. These data suggest that NPFR signaling is involved in the regulation of ecdysone biosynthesis in the PG, although it should be noted that it could play an additional role in regulating ecdysone secretion.

Loss of NPFR signaling phenocopies loss of insulin signaling

Changes in insulin signaling also regulate development time by regulating the rate of ecdysone synthesis (Caldwell et al. 2005; Colombani et al. 2005; Mirth et al. 2005; Koyama et al. 2014). This has been demonstrated in animals which are hypomorphic for loss of insulin signaling (complete loss causes early lethality; Fernandez et al. 1995), such as flies that bear a heteroallelic combination of mutations in the insulin receptor (InR), or are homozygous for a loss of function mutation of the adaptor protein, chico (Shingleton et al. 2005). These animals take longer to reach metamorphosis and have decreased adult body sizes (Stocker and Hafen 2000; Shingleton et al. 2005). As NPFR mutants are homozygous viable, like chico mutants, we hypothesized that they may be hypomorphic for loss of insulin signaling. In support of this, we found that in addition to being developmentally delayed, NPFRSK8 mutants have smaller body sizes compared to controls (Figure 3A, P < 0.0001). Similarly, NPF >NPFi animals are smaller than controls (Figure 3B, P < 0.001). To check that the body size defect was not a result of decreased food intake due to altered feeding behavior, we quantified food intake in NPFRSK8 mutants on our standard fly food. This showed that, under well fed conditions, there were no significant differences in the amount of food consumed compared to controls (Figure S2A, P = 0.588).

Figure 3.

Figure 3

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NPFRSK8 mutants phenocopy loss of insulin signaling. (A) NPFRSK8 mutants and (B) NPF > NPFi animals have a smaller body size compared to controls, as measured by pupal length (P < 0.0001 and P < 0.001, ANOVA and pair-wise t-test, respectively). Each point represents an individual pupa, and ≥40 individuals were tested per genotype. (C) NPFRSK8 mutants have a reduced rate of growth compared to controls (P < 0.01, linear regression analysis). hAEL = hr after egg lay; 10–15 animals were tested per time point for each genotype. (D) NPFRSK8 mutants have reduced levels of phosphorylated Akt (pAkt). (E) Quantification of pAkt/Tubulin densities were standardized by fixing the values of NPFRSK8/+ to 1 (P = 0.0031, pair-wise t-test). Six biological replicates of 10 animals were used per genotype. (F) NPFRSK8 mutants fed on diets of decreasing caloric density do not adjust their body size differently to controls (P > 0.05, linear regression analysis); 10 biological replicates of 10–15 animals were used per diet. For all graphs, genotypes sharing the same letter indicate that they are not significantly different from one another while genotypes with contrasting letters indicate that they are statistically different. Error bars represent ±1 SEM for all graphs.

Whole-animal mutants of the insulin signaling pathway have smaller body sizes due to reduced growth rates (Böhni et al. 1999). We therefore measured the growth rate of NPFRSK8 mutants. This showed that NPFRSK8 mutants have a significantly reduced growth rate compared to controls (Figure 3C). Together, these data suggest that animal-wide loss of NPFR phenocopies animal-wide reduction in insulin signaling.

Given these similarities in phenotype, we hypothesized that NPFR interacts with the insulin signaling pathway to regulate ecdysone production. To test this idea, we quantified insulin signaling levels in NPFR mutants by measuring levels of phosphorylated protein kinase B (pAKT), a downstream component of the insulin signaling pathway, relative to tubulin as a loading control. A significant reduction in pAKT level was observed in NPFRSK8 mutants compared to heterozygous controls (Figure 3, D and E, P = 0.0031). This demonstrates that whole animal loss of NPFR leads to an overall reduction in insulin signaling.

To further explore this, we looked at another well-known phenotype of loss of insulin signaling; response to nutrition. Wild-type animals that are fed on less-nutritious foods have a reduction in final body size (Robertson 1963), presumably because of the resulting reduction in insulin signaling under poor nutritional conditions. When insulin signaling is suppressed in an organ, the organ loses its ability to adjust its size in response to nutrition (Tang et al. 2011). If NPFR mutants have an overall reduction in insulin signaling, then they should also have decreased body size plasticity in response to less nutritious foods. We therefore fed NPFRSK8 mutants diets of varied caloric concentration, and measured pupal weight as an indication of body size. This showed that these animals had the same sensitivity to nutrition as controls, with indistinguishable slopes between body size and the caloric concentration of the food between genotypes (Figure 3F). Taken together, these data suggest that loss of NPFR reduces, but does not fully ablate, overall insulin signaling. The reduction in insulin signaling is sufficient to cause reduced body size and growth rate, but not enough to interfere with plasticity in body size in response to poor nutrition.

NPFR negatively regulates insulin signaling in the PG

Given that NPFRSK8 mutants have reduced insulin signaling, and that knocking down NPFR in the PG reduces ecdysone synthesis, we next wanted to determine if NPFR modifies insulin signaling specifically in this organ. Changes in insulin signaling in the PG could cause a developmental delay under one of two different scenarios: (1) if insulin signaling is increased in the PG early in third-instar larvae, or (2) if insulin signaling is reduced in the PG in the mid- to late-third-instar. This is because insulin signaling has different roles before and after an important developmental checkpoint in third instar larvae, known as “critical weight” (Nijhout 2003; Mirth et al. 2005; Shingleton et al. 2005). Prior to critical weight, either starving animals or reducing insulin signaling results in a developmental delay (Beadle et al. 1938; Mirth et al. 2005; Shingleton et al. 2005). This occurs because low levels of insulin signaling delay the timing of the ecdysone pulse that is necessary to trigger the critical weight checkpoint (Mirth et al. 2005). After critical weight has been achieved, starving animals has the opposite effect and accelerates developmental timing (Mirth et al. 2005; Stieper et al. 2008; Koyama et al. 2014). This has previously been described as the “bail out response,” referring to the fact that under starvation conditions (when insulin signaling is low), the developmental program encourages the animal to pupariate (Nijhout et al. 2014; Hatem et al. 2015). Thus, an increase in insulin signaling in late-third-instar animals should cause a developmental delay.

We therefore set out to determine if loss of NPFR in the PG reduces or increases insulin signaling in this tissue, and if this changes over the third larval instar period. To do this we examined the localization of the transcription factor, Forkhead Box class O (FoxO), a commonly used readout of insulin signaling pathway activity (Jünger et al. 2003), in PG cells. When insulin signaling is high FoxO is cytoplasmic, and when insulin signaling is low FoxO is localized to the nucleus (Jünger et al. 2003). We knocked down NPFR in the PG and determined FoxO localization in both early and mid- to late-third-instar animals. In both cases, PG cells in phm > NPFRi; dcrII animals had significantly more cytoplasmic FoxO than controls (Figure 4, A and B; P = 0.01, P = 0.0011, respectively). This suggests that insulin signaling activity is increased in the PG in both early and late-third-instar larvae when NPFR is knocked down specifically in this tissue, and thus that NPFR normally functions to repress insulin signaling in the PG in third instar larvae. The developmental delay that was observed by knocking down NPFR throughout the third instar could then be explained if the increase in insulin signaling precritical weight is not sufficient to accelerate developmental timing, but the increase in insulin signaling during the late third instar is sufficient to cause a developmental delay.

Figure 4.

Figure 4

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NPFR negatively regulates insulin signaling in the prothoracic gland. (A) Knocking down NPFR specifically in the PG results in increased cytoplasmic FoxO accumulation in precritical weight larvae (0hAL3E) compared to controls (P = 0.01, ANOVA). (B) Knocking down NPFR specifically in the PG results in primarily cytoplasmic FoxO in postcritical weight larvae (48hAL3E) compared to controls (P = 0.0011, ANOVA). hAL3E = hr after L3 ecdysis; 8–10 PGs were analyzed per genotype. Scale bar is 10 μm. (C) Knocking NPFR down specifically in the PG results in an increase in final body size as measured by pupal length (P = 0.0041, ANOVA). Each point represents an individual pupa and ≥40 individuals were tested per genotype. For (A–C), error bars represent ±1 SEM. Genotypes sharing the same letter are not significantly different from one another, while genotypes with different letters are statistically different. (D) phm > NPFRi; dcrII animals display significantly different changes in final body size compared to controls when fed on diets of decreasing caloric density (P < 0.01, linear regression analysis); 10 biological replicates of 10–15 animals were assessed per diet.

We next measured body size when NPFR is knocked down in the PG, using pupal length as an indication of final body size. This was of interest as increasing insulin signaling before critical weight would be expected to cause a decrease in body size, whereas increasing insulin signaling after critical weight would be expected to cause an increase in body size. We found that phm> NPFRi; dcrII animals are larger than controls (Figure 4C, P = 0.0041). To ensure the body size alteration was not a result of increased food intake, we quantified food intake in the phm > NPFRi; dcrII animals. This showed that there were no significant differences in the amount of food consumed in these animals compared to controls (Figure S2B, P = 0.414). Taken together, while the FoxO localization data suggests NPFR is able to repress insulin signaling throughout L3, the combination of a developmental delay and increased body size observed when we knock down NPFR in the PG suggests that NPFR function in the PG is most important postcritical weight.

Lastly, we looked at the response to nutrition. If knocking down NPFR in the PG causes increased insulin signaling in the gland, this could result in animals being more sensitive to nutrition (Tang et al. 2011). To test this, we knocked down NPFR in the PG, and animals were fed on one of four concentrations of food. We then measured pupal weight as an indication of final body size (Figure 4D). On a diet of standard caloric concentration (1×), phm> NPFRi; dcrII animals were significantly larger than controls. As the calorie concentration in the food decreases, we observed a much steeper decrease in body size for phm > NPFRi; dcrII animals compared to controls. This demonstrates that body size is indeed more sensitive to nutrition in these animals, further supporting a negative regulatory role in insulin signaling.

Taken together, given that NPFR regulates insulin signaling in the PG, these data suggest that under nutritional stress NPF could be both acting on NPFR neurons in the brain to regulate feeding behavior (Wu et al. 2003; Wu et al. 2005) while also signaling through NPFR in the PG to regulate developmental timing. In this way, NPF signaling could act as a nexus between feeding behavior and development.

In summary, here we have described a new role for the conserved feeding regulator, NPFR, in the regulation of developmental timing, animal growth rate, and body size. In the PG, our data supports a role for NPFR in negatively regulating the insulin signaling pathway and positively regulating ecdysone biosynthesis. By contrast to the results we obtained in the PG, we found that whole animal loss of NPFR generates phenotypes resembling those where insulin signaling is reduced throughout the whole body. The simplest explanation of these contrasting phenotypes is that NPFR has a second role in regulating developmental timing and body size elsewhere in the fly, perhaps due to a role in regulating insulin-like peptide production or secretion. These data not only highlight a previously undescribed mechanism by which insulin signaling and ecdysone production are regulated in the PG, but also demonstrate how a single neuropeptide signaling pathway can have functionally diverse roles within an organism in response to the same environmental cue.

Our findings raise the strong possibility that NPF may indeed coordinate feeding behavior and growth. How might it do so? Other peptides known to function in the regulation of ecdysone production are produced either in neurons that directly innervate the PG, such as PTTH (McBrayer et al. 2007), or in other tissues and secreted into circulation, such as the Dilps (Brogiolo et al. 2001; Ikeya et al. 2002). Either a local or systemic source of NPF is therefore possible for activating NPFR in the PG. While neuropeptides such as NPF are best described as having local modes of action, in the adult fly, NPF has recently been shown to be secreted from the midgut into the hemolymph, where it can act systemically (Ameku et al. 2018). In the larva, NPF is known to be expressed in dopaminergic neurons in the brain, as well as cells in the midgut (Brown et al. 1999). To our knowledge, it has not been shown to be expressed in neurons that innervate the larval PG. This suggests that it is more likely that systemic rather than local NPF activates NPFR in the larval PG cells. Systemic NPF produced in response to nutritional stress could thus act on both NPFR neurons in the brain to regulate feeding behavior and on NPFR in the PG to regulate developmental timing, and, in so doing, NPF could coordinate feeding behavior and development.

In conclusion, this study has provided evidence to show that NPFR signaling, best known for its regulation of feeding behavior, also functions in the Drosophila PG to control developmental timing and body size via regulation of insulin signaling and ecdysone production. To our knowledge, NPF represents the first neuropeptide described to play a role in regulating both feeding behavior and development in response to nutritional conditions, and, thus, the first candidate for coordinating these processes in response to environmental cues. Given that the mammalian homolog of NPF, NPY, also has a role in regulating feeding behavior in response to nutritional stress, it would be of great interest to explore if it too is a candidate for coordinating behavior and development.

Acknowledgments

We would like to thank Karyn Moore, Emily Kerton, and the Australian Drosophila biomedical research facility (OzDros) for technical support; Michael O’Connor for providing fly stocks; and Pierre Leopold for the FoxO antibody. M.A.H. is a National Health and Medical Research Council (NHMRC) Early Career Fellow. This work was supported by an Australian Research Council (ARC) grant to C.G.W. and C.K.M. Stocks obtained from the Bloomington Drosophila Stock Center [National Institutes of Health (NIH) P40OD018537] were used in this study. The authors declare no competing interests.

Author contributions: C.G.W. and C.K.M. conceived the experiments, interpreted the data and co-led the work. J.R.K. conceived the experiments, interpreted the data, and performed the experiments. M.A.H interpreted the data and assisted with experiments. L.M.P. interpreted the data. S.K. generated the NPFR mutant Drosophila strain. J.R.K., C.G.W. and C.K.M. wrote the manuscript, with assistance from L.M.P.

Footnotes

Supplemental material available at figshare: https://doi.org/10.25386/genetics.12661226.

Communicating editor: R. Duronio

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Associated Data

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

Data Availability Statement

All strains are available upon request. We affirm that all data necessary for confirming conclusions of this article are present within the article and figures. Supplemental material available at figshare: https://doi.org/10.25386/genetics.12661226.

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