Energy Homeostasis Control in Drosophila Adipokinetic Hormone Mutants

Abstract

Maintenance of biological functions under negative energy balance depends on mobilization of storage lipids and carbohydrates in animals. In mammals, glucagon and glucocorticoid signaling mobilizes energy reserves, whereas adipokinetic hormones (AKHs) play a homologous role in insects. Numerous studies based on AKH injections and correlative studies in a broad range of insect species established the view that AKH acts as master regulator of energy mobilization during development, reproduction, and stress. In contrast to AKH, the second peptide, which is processed from the Akh encoded prohormone [termed “adipokinetic hormone precursor-related peptide” (APRP)] is functionally orphan. APRP is discussed as ecdysiotropic hormone or as scaffold peptide during AKH prohormone processing. However, as in the case of AKH, final evidence for APRP functions requires genetic mutant analysis. Here we employed CRISPR/Cas9-mediated genome engineering to create AKH and AKH plus APRP-specific mutants in the model insect Drosophila melanogaster. Lack of APRP did not affect any of the tested steroid-dependent processes. Similarly, Drosophila AKH signaling is dispensable for ontogenesis, locomotion, oogenesis, and homeostasis of lipid or carbohydrate storage until up to the end of metamorphosis. During adulthood, however, AKH regulates body fat content and the hemolymph sugar level as well as nutritional and oxidative stress responses. Finally, we provide evidence for a negative autoregulatory loop in Akh gene regulation.

ENERGY homeostasis requires continuous compensation for fluctuations in the energy expenditure and availability of food resources. Organisms thus build up reserves under positive energy balance and catabolize them when the balance turns negative to retain stable levels of circulating energy fuel. Insulin signaling induces the uptake of excessive circulating sugars, thus promoting reserve accumulation (reviewed, e.g., in Saltiel and Kahn 2001; Cohen 2006), whereas energy mobilization is under the control of glucagon and glucocorticoid signaling in mammals (reviewed, e.g., in Rui 2014; Charron and Vuguin 2015) and adipokinetic hormone (AKH) signaling in insects (reviewed, e.g., in Van der Horst 2003; Lorenz and Gäde 2009; Bednářová et al. 2013a). Consistent with their fundamental physiological function in energy mobilization, AKHs are found not only in insects, but are common in Protostomia, where they have been identified both in Ecdyszoa (in Arthropoda, Tardigrada, and Priapulida) and Lophotrochozoa (in Mollusca, Rotifera, and Annelida) (Gäde 2009; Hauser and Grimmelikhuijzen 2014). Nevertheless, physiological functions of AKHs have been studied mainly in Arthropoda. Similar to mammals, also insects store lipids in the form of triacylglycerides (TGs) and as carbohydrates in the form of glycogen. The main storage organ for lipid and glycogen in insects is the fat body, which can thus be considered as the functional equivalent of mammalian liver and white adipose tissue (Azeez et al. 2014). Under energy-demanding conditions, AKHs get released from the organ of their synthesis and storage, the corpora cardiaca (CC), into the hemolymph to reach their cognate G protein-coupled receptors (GPCRs) (called AKH receptors, AKHRs) expressed on the fat body cells. This induces TG and glycogen breakdown, which leads to the production and release of circulating carbohydrates (trehalose and glucose), lipids [diacylglycerides (DGs)], proline, or a combination of these fuel molecules, depending on the species` preference. Despite the divergence in the preferred type of circulating fuel, the role of AKH in the mobilization of energy stores is evolutionarily conserved among insects (reviewed in Gäde and Auerswald 2003; Lorenz and Gäde 2009). Consistently, AKH was identified as the hyperglycemic hormone in cockroaches (Steele 1961), as the hyperlipaemic hormone in Locusta (Beenakkers 1969; Mayer and Candy 1969) and as the hyperprolinaemic hormone in the fruit beetle Pachnoda sinuata (Auerswald and Gäde 1999). Next to this primary, catabolic role, numerous studies have implicated AKH in a puzzling diversity of additional physiological functions ranging from behavior (Kaun et al. 2008), locomotion (Kodrík et al. 2000; Socha et al. 2008), and reproduction (Lorenz 2003; Attardo et al. 2012) to digestion (Vinokurov et al. 2014), heart beat control (Noyes et al. 1995), sleep (Metaxakis et al. 2014), immunity (Adamo et al. 2008), oxidative stress resistance (Bednářová et al. 2013a,b; Plavšin et al. 2015), aging (Katewa et al. 2012; Waterson et al. 2014), and muscle contraction (Stoffolano et al. 2014). It is not clear so far whether these pleiotropic effects of AKH result from changes in energy homeostasis, or rather reveal the existence of distinct AKH-regulated signaling pathways, which implement independent functions of AKH. Individual studies used typically different species of various ontogenetic stages, and thus it remains elusive, whether the described roles reflect the general functions of AKH, which might be developmentally conserved across ontogenetic stages and evolutionarily conserved across insect species. The majority of AKH studies have been done in the popular insect endocrinology models like Locusta, Manduca, Gryllus, or Pyrrhocoris, where AKH roles have been addressed mostly as physiological changes induced by injections of synthetic AKH, or as correlations between the hormone titer and varying environmental or physiological conditions. A more comprehensive understanding of the AKH functions requires genetic loss-of-function analyses.

Recent advances in available technologies, like mass spectrometry (MS), and the emergence of the CRISPR/Cas9 tool for genomic engineering added to the advantages of the model insect species Drosophila melanogaster, strengthening its potentials in the field of insect endocrinology. Drosophila already proved to be an excellent model system to investigate several other conserved hormonal pathways, including, e.g., insulin/insulin-like signaling (Kannan and Fridell 2013; Nässel et al. 2015). However, in contrast to the numerous scientific reports dealing with AKH roles in nondrosophilid species, only a limited number of studies have focused on AKH signaling in Drosophila so far. In the absence of a specific mutant, tools for AKH studies have remained limited to the ablations of CC cells (Kim and Rulifson 2004; Lee and Park 2004; Isabel et al. 2005), stimulation of secretory activity of CC cells by their depolarization (Kim and Rulifson 2004), mutations of the receptor (Grönke et al. 2007; Bharucha et al. 2008; Waterson et al. 2014), overexpression of Akh complementary DNA (cDNA) (Kim and Rulifson 2004; Lee and Park 2004; Grönke et al. 2007; Katewa et al. 2012; Baumbach et al. 2014b), and Akh RNA interference (RNAi) (Braco et al. 2012; Baumbach et al. 2014b). Ablation of CC cells, or their depolarization, are not AKH-specific manipulations, as these endocrine cells also produce other hormones like limostatin, which affects metabolism by regulation of insulin signaling (Alfa et al. 2015). In addition, it cannot be excluded that a limited amount of hormone is produced prior to the ablation and interferes with the investigation of the early developmental functions of AKH. Overexpression of AKH cDNA or RNAi are also not explicit methods to address AKH functions. Next to the typically incomplete down-regulation by RNAi, another level of complication comes from the structure of the Akh gene product. Processing of the 79-amino-acid-long AKH prohormone results in two peptides: AKH and adipokinetic hormone precursor-related peptide (APRP) (Figure 1A). Hence, overexpression of Akh RNAi decreases both AKH and APRP and overexpression of Akh cDNA increases both of these peptides. Differentiation between the effects of AKH and APRP loss is especially important in the light of the potential hormonal functions of APRP. Even though no role of APRP was described so far (Hatle and Spring 1999), its evolutionary conservation, common ancestry with mammalian growth hormone–releasing factors (Clynen et al. 2004), and release upon stimulation with melatonin (Huybrechts et al. 2005) all argue in favor of an endocrine function of this peptide. Thus, unequivocal study of AKH and APRP functions requires creation of specific mutants, which we describe in this study.

Figure 1.

Genomic organization of the Akh gene locus and molecular identity of the AKH single mutant Akh^A and the AKH plus APRP double mutants Akh^AP and Akh^SAP. (A) Genomic organization of the Akh gene flanked by the Ras64B and CG32260 genes on the left arm of the D. melanogaster third chromosome (open boxes represent transcribed regions, filled boxes open reading frames). Wild-type Akh (Akh⁺) encodes a prohormone consisting of the signal peptide (yellow), the AKH octapeptide (red), a protease cleavage site (light gray), and the APRP (blue). Note that the RNAi target sequence spans most of the Akh open reading frame. Scale bars represent DNA sequences in base pairs (bp). Schematic drawing (A) and sequence representation (B) of the molecular lesions of Akh mutants compared to the prohormone (pAKH⁺) coding sequence. The AKH-specific mutant Akh^A lacks the sequences coding for the two amino acids (DW) at the C-terminal positions 7 and 8 of the AKH octapeptide. In AKH plus APRP double mutants Akh^AP and Akh^SAP, AKH coding sequences are deleted and APRP expression is compromised due to frame shift (Akh^AP) or due to deletion of the signal peptide coding sequence including the translation start codon (Akh^SAP). The underlined sequence in B corresponds to the Akh gRNA and the black triangle indicates the Cas9 cleavage site used for Akh mutant generation by CRISPR/Cas9-mediated Akh genome engineering. For details see text.

Biochemical studies identified a single Drosophila AKHR receptor (Park et al. 2002; Staubli et al. 2002), which has been functionally analyzed in vivo (Grönke et al. 2007; Bharucha et al. 2008). However, these studies do not exclude AKH signal transmission via other receptor pathways. Thus, throughout our study, we compared the newly created AKH mutants with mutants lacking the AKHR.

Here, we present the first functional analysis of AKH single mutants and AKH plus APRP double mutants in essential biological processes ranging from embryogenesis, metamorphosis, reproduction, lipid and carbohydrate storage, and mobilization to stress resistance. Our work shows that developmental mobilization of energy stores, oogenesis, and locomotion are under the control of AKH-independent pathways in Drosophila. However, Drosophila AKH is involved in response to stress, including nutritional and oxidative challenge. We also show that the metabolic roles of AKH in stored energy sources in Drosophila are developmental-stage specific; whereas AKH signaling is dispensable for accumulation and mobilization of storage lipids and glycogen during larval and pupal periods, it acquires important roles in homeostasis of storage lipids, but not storage carbohydrates during adulthood.

Materials and Methods

Fly stocks and fly husbandry

If not stated otherwise, D. melanogaster were reared at 25° in a 12-hr light/12-hr dark cycle, on standard Drosophila medium (5.43 g agar, 15.65 g yeast, 8.7 g soy flour, 69.57 g maize flour, 19.13 g beet syrup, 69.57 g malt, 5.43 ml propionic acid, and 1.3 g methyl 4-hydroxybenzoate per 1 liter of medium; supplier information available on request). Experiments were started with ∼150 eggs per 68-ml vial. Freshly eclosed adults were collected and kept at density of ∼50 females + 50 males per 68-ml vial and transferred to fresh media every second day. Pupae were staged according to Bainbridge and Bownes (2008).

Stocks used to create Akh mutant lines:

The stocks used to create Akh mutant lines are as follows: y¹ sc* v¹; P{v*; BFv-U6.2_Akh_gRNA}attP40 (this study); w*; Kr^If-1/CyO; D¹/TM3, Ser¹ (Bloomington Drosophila Stock Center, BDSC 7198); P{ry^+t7.2 = hsFLP}1, y¹ w¹¹¹⁸; P{y^+t7.7 w^+mC = UAS-Cas9.P}attP2, P{w^+mC = GAL4::VP16-nos.UTR}CG6325^MVD1 (BDSC 54593, Port et al. 2014); and w*; TM3, Sb¹, P{2xTb1-RFP}TM3/ln(3L)D D¹ (BDSC 36338).

Mutant stocks:

The mutant stocks are as follows: w*; Akh^A (this study); w*; Akh^AP (this study); w*; Akh^SAP (this study); and y*w*; AkhR¹ (Grönke et al. 2007).

Mutant and balancer lines established after backcrossing into w¹¹¹⁸ for nine generations:

The mutant and balancer lines established after backcrossing into w¹¹¹⁸ for nine generations are as follows: w¹¹¹⁸; Akh^A/ TM3, Ser¹ floating (this study); w¹¹¹⁸; Akh^AP/TM3, Ser¹ floating (this study); w¹¹¹⁸; AkhR¹ (this study), and w¹¹¹⁸; +/CyO; +/TM3, Ser¹ (transient line).

Additional stocks:

Additional stocks are as follows: w*; akhp-GAL4, UAS-mCD8 GFP; akhp-GAL4/SM5a-TM6 Tb (Kim and Rulifson 2004); Akh RNAi (VDRC 11352; w¹¹¹⁸; akhp-GAL4, UAS-mCD8 GFP/CyO (this study); w¹¹¹⁸; akhp-GAL4, UAS-mCD8 GFP/CyO; Akh^A/TM3 Ser¹ floating (this study); w¹¹¹⁸; akhp-GAL4, UAS-mCD8 GFP/CyO; Akh^AP/TM3 Ser¹ (this study); w¹¹¹⁸; akhp-GAL4, UAS-mCD8 GFP/CyO; Akh^SAP/TM3 Ser¹ (this study); and Canton-S; w¹¹¹⁸ (Vienna Drosophila RNAi Center, VDRC 60000).

CRISPR/Cas9-mediated mutagenesis of the Akh gene

The Akh^A (AKH⁻), Akh^AP, and Akh^SAP (AKH⁻ APRP⁻) mutants were generated by CRISPR/Cas9-mediated genomic engineering according to Kondo and Ueda (2013). For details on the generation of the mutants see Supporting Information, File S1.

Creation of Akh^A, Akh^AP, and Akh^SAP stable stocks and backcrossing of the mutant alleles to a common genetic background:

Homozygous stocks were established from the progeny of selected males (genotype: w*; Akh*/TM3, Sb¹, P{2xTb1-RFP) carrying molecular lesions in the Akh gene, which were molecularly characterized by genomic sequencing to reveal the following three Akh mutants used in this study: Akh^A, Akh^AP, and Akh^SAP. As physiological parameters are prone to confounding genetic background effects, the Akh^A, Akh^AP, and Akh^SAP alleles, together with the previously described null mutation of the AKH receptor, AkhR¹ (Grönke et al. 2007), and a CyO TM3, Ser¹ balancer line (based on BDSC 7198) were backcrossed into standard w¹¹¹⁸ genetic background (VDRC 60000) for nine generations prior to stock establishment. For primer sequences used to track the mutations during the backcrossing, see File S1.

If not stated otherwise all physiological assays were conducted on the backcrossed mutants and the genetically matched control.

Mass spectrometry

Dissection and sample preparation for mass spectrometry:

Retrocerebral complexes (RCs) of adult flies were dissected in ice-cold ammonium chloride buffer (1.404 g Na₂HPO₃ × 2 H₂O, 0.262 g NaH₂PO₄ × H₂0, 8.8 g NaCl in 1 liter aqua bidest; pH 7.1) using fine forceps and a stereo binocular. Single preparations were washed in an ice-cold drop of MS grade water (TraceSELECT Ultra, Fluka Analytical, St. Louis) and either transferred to a stainless steel plate for direct tissue profiling or collected in 20 µl ice-cold extraction buffer [50% MeOH, 0.5% formic acid (FA), 49.5% H₂O (v/v)] in a 0.5-ml protein LoBind tube (Eppendorf, Hamburg). Extracts were incubated for 30 min on ice and centrifuged at 4° and 12,000 × g for 15 min. Subsequently, extracts were incubated for 1 min in an ultrasonic bath filled with ice and centrifuged for 15 min at 4° and 12,000 × g. Ultrasonic bath incubation was repeated three times for better peptide extraction. Resulting supernatants were stored at −20° for further analysis. For direct tissue profiling (mass fingerprints), transferred tissues were left to dry and subsequently covered with 50–75 nl of 2,5-dihydroxybenzoic acid (Sigma-Aldrich, St. Louis; 10 mg/ml DHB, in 20% ACN, 1% FA, and 79% H₂O) matrix solution using a fine glass capillary. For on-plate disulfide reduction, samples were covered with 0.1 µl freshly prepared 1,5-diaminonapthalene (Sigma-Aldrich; 10 mg/ml DAN, in 50% ACN, 0.1% trifluoroacetic acid, and 49.9% H₂O) matrix solution. For control experiments and APRP identification, Canton-S (CS) wild-type flies were used.

Reduction of disulfide bonds and carbamidomethylation of cysteines:

Extracts of RCs were treated as described in Sturm and Predel (2015). Supernatants were first reduced to a volume of 5 µl by vacuum centrifugation and mixed with 20 µl of 50 mM ammonium bicarbonate (ABC) buffer. The pH value was adjusted to 8 with NaOH. Disulfide bonds were reduced by adding 1,4-dithiothreitol (200 mM; DTT) in ABC buffer to an end concentration of 10 mM DTT, for 1 hr at 37°. Subsequently, cysteines were carbamidomethylated by adding iodoacetamide (200 mM; IAA) in ABC buffer to an end concentration of 40 mM, at 37° for 1 hr in darkness. Excess IAA was precipitated by adding DTT with a final concentration of 40 mM DTT. The resulting mixture was incubated for 30 min at room temperature. Samples were acidified with 0.5% acetic acid (AA) and loaded on an activated (100% MeOH) and equilibrated (0.5% AA) homemade C18 spin column (Empore 3M extraction disc; IVA Analysentechnik, Meerbusch, Germany) as described in Rappsilber et al. (2007). The column was rinsed with 100 µl 0.5% AA. Peptides were eluted with different concentrations of MeOH (10, 20, 25, 30, 40, 50, 80, and 100% with 0.5% AA) onto a sample plate and subsequently mixed with DHB [sample matrix ration 1:1 (v/v)].

MALDI-TOF mass spectrometry:

Mass spectra were acquired on a Bruker UltrafleXtreme TOF/TOF mass spectrometer (Bruker Daltonik, Bremen, Germany) under manual control in reflectron positive ion mode. Instrument settings were optimized for mass ranges of m/z 600–4000, m/z 3000–10,000, and m/z 8000–15,000. The instrument was calibrated for each mass range, using a mixture of bovine insulin, glucagon, ubiquitin, and cytochrome C (Sigma-Aldrich) or the Peptide Calibration Standard II (no. 222570, Bruker Daltonik). MS² spectra were acquired in postsource decay (PSD) mode without a collision gas. All obtained data were processed with FlexAnalysis 3.4 software package (Bruker Daltonik).

Fecundity assay

Fecundity was measured as daily egg scores of individual females during the first 10 days of their lives. One female and two males of the same genotype that eclosed within a 24-hr period were placed on standard fly food with active dry yeast added on the top (∼5 mg of yeast per vial). Flies were transferred daily to fresh vials and the eggs were counted under a stereomicroscope. Fecundity was expressed as mean cumulative number of eggs laid by a single female during the first 10 days of life. Females that died or escaped during the experiment were excluded from the analyses. A total of 26–30 females were tested per genotype. Data were analyzed by one-way ANOVA. After 10 days, body size of females was measured as described below.

Hatchability assay

Eggs laid by the females at days 6 and 10 of the fecundity assay were kept at 25°, and the percentage of hatched eggs (hatchability) was determined 24 hr later. Data from the 6th day are plotted in Figure 3A. Embryos from 26–30 females were tested per genotype; hence, hatchability of 2147–2695 eggs was tested per genotype. Data were arcsine square root transformed and subsequently analyzed by one-way ANOVA followed by Tukey´s honest significant difference (HSD) post hoc test.

Figure 3.

AKH, APRP, and AKHR are dispensable for preadult development. Plotted are means ± SEM. (A) AKH, APRP, and AKHR are not required for embryogenesis. Slightly, but significantly decreased hatchability of Akh^A as compared to controls; Akh^AP and AkhR¹ had normal hatchability (one-way ANOVA, F_3,105 = 8.65, P < 0.001; Tukey’s HSD: P < 0.05). (B) AKH, APRP, and AKHR are not necessary for larval to adult survival; no significant difference between any of the mutants when compared to control (one-way ANOVA, F_3,101 = 3.86, P < 0.05; Tukey’s HSD: P < 0.05). (C) Developmental rate (measured as time span from egg laying to adult eclosion) was not extended in AKH single mutants nor AKH plus APRP double mutants nor AkhR mutants. On the opposite, deficiency in the AKH signaling slightly increased the speed of development; in the case of Akh^AP and AkhR¹, this effect reached statistical significance (one-way ANOVA, F_3,585 = 71.44, P < 0.001; Tukey’s HSD: P < 0.05). (D) AKH, APRP, and AKHR are all dispensable for female fecundity. No difference in egg laying of the mutants compared to the genetic control (one-way ANOVA, F_3,107 = 1.8, P = 0.15). (E) AKH, APRP, and AKHR are all dispensable for regulation of body size. No difference in the thorax length between mutants and genetic control is seen (one-way ANOVA, F_3,103 = 1.94, P = 0.13). Plotted data were obtained on female flies.

Viability (larval to adult survival) assay

Larva to adult viability was expressed as percentage of flies that eclosed from the hatched eggs. The same larvae from the hatchability assay were used. Per genotype, 1239–2473 larvae were tested. Data were arcsine square root transformed and subsequently analyzed by one-way ANOVA.

Developmental rate determination

Developmental rate was measured as the time from the egg laying to adult eclosion. Four-day-old parental flies were transferred from the standard food to standard food supplemented with sprinkled active yeast and this food was replaced daily for 3 consecutive days to prevent egg retention. Then, the parental flies were allowed to lay eggs on fresh food for 2 hr. Embryos laid within this interval were collected and their density was adjusted to ∼150 embryos per 68-ml vial and kept under standard conditions afterward. The number of eclosed flies was recorded three times per day (at 8 am, 3 pm, and 8 pm), until the last fly eclosed. The experiment was repeated twice. Between 196 and 418 eclosed flies were tested per genotype per trial. Data were log transformed before performing one-way ANOVA and Tukey´s HSD post hoc test.

Body size determination

Thorax length of females used for the fecundity assay was measured using an Axiophot Zeiss microscope equipped with a digital camera AxioCam HRc and ZEN 2011 software. Thorax length was measured from the posterior tip of the scutellum to the base of the most anterior humeral bristle (French et al. 1998). Thorax length of 25–29 females was measured per genotype. Data were analyzed by one-way ANOVA.

Preparation of homogenates for metabolic measurements (glycogen, lipid, and protein determination)

Male flies were homogenized by Mixer Mill (MM400, Retsch), in 1.2-ml collection tubes (Qiagen) with 5-mm metal beads and 600 μl 0.05% Tween-20. Homogenates were heat inactivated for 5 min at 70°, centrifuged for 3 min at 3500 rpm (= 2200 rcf; Heraeus Megafuge 1.0R, swing-out rotor no. 2704), and 400 μl of the supernatant was transferred into 96 Master block well plates (Greiner Bio One) for storage.

Protein determination

Protein content was determined by Pierce BCA Protein Assay Kit according to the instructions of the manufacturer, with the following modification: samples were analyzed in a 96-well plate format, using 50 μl of the fly homogenate per 250 μl reaction volume. For each genotype, four to six replicates of five flies each were tested per developmental stage or starvation time point. Experiments were repeated at least three times.

Lipid determination by coupled colorimetric assay

Lipid measurements were done as described in Hildebrandt et al. (2011). For a more detailed description see File S1.

Glycogen determination

Glycogen measurements in the 96-well plate format were based on the conversion of glycogen to glucose by amyloglucosidase (Sigma) and on its subsequent measurement by the glucose assay (GO) kit (Sigma) as described in Tennessen et al. (2014). For a more detailed description see File S1.

Determination of circulating sugars

Hemolymph samples (three replicates of 30 flies each per genotype) were collected by centrifugation (6 min, 9000 rcf at 4°) of decapitated flies in a holder tube (0.2-ml tube with five holes of 0.6-mm diameter) placed in a 0.5-ml collection tube. A total of 1 μl of the collected hemolymph was diluted with 99 μl of 0.05% Tween-20 and immediately heat inactivated at 70° for 5 min. The homogenate was further diluted 1:6 prior to the sugar measurements. Measurements of circulating sugars were performed using a modification of the Tennessen method (Tennessen et al. 2014). For a more detailed description see File S1.

Thin layer chromatography

The thin layer chromatography (TLC) analysis was performed as described by Baumbach et al. 2014a), with minor modifications. For a more detailed description see File S1.

Confocal laser scanning microscopy of adult fat body tissue

Adult fat body tissue of 6-day-old male flies was dissected in ice-cold 1× PBS. Flies were mechanically fixed with a preparation pin through the thorax with the ventral side upwards. Then the fly was cut with a fine scissor in transversal plane directly after the abdominal tergite 6. Additional cuts were performed in the coronal plane along the tergital–sternital intersections. The sternital parts, the digestive and reproductive system, were removed to expose the cuticle-attached fat body. This carcass preparation was embedded in 1× PBS containing Bodipy493/503 (38 µM; Invitrogen, D3922) for lipid droplet staining, DAPI (3,6 µM; Invitrogen, D1306) for nuclei staining, and CellMask Deep Red (5 µg/mL; Invitrogen, C10046) for plasma membrane staining. Images were acquired in 16-bit mode with a Zeiss LSM-780 microscope and a C-Apochromat 40×/1.20 W Korr FCS M27 objective adjusted in dynamic signal range for the control genotype. For fluorescence detection, the following settings were used: DAPI (Excitation (Ex): 405 nm, Emission (Em): 410–468 nm), Bodipy493/503 (Ex: 488 nm; Em: 490–534 nm), and CellMask (Ex: 561 nm; Em: 585–747 nm). Images were analyzed with ImageJ v1.49m for lipid droplet area by first applying a “Gaussian blur” filter (2.0 pixel range) to a single optical section, in to smooth the edges of the lipid droplets. Afterward, a binary image with discrete lipid droplets was generated by thresholding (removal of “below 60%”). The “watershed” tool was applied to the image to separate the area of clustered lipid droplets. Finally, the particle analyzer was applied on the picture [size (µm²): 0.1→∞; circularity: 0.01–1.0; mark outlines] for area determination of the discretely detected particles. Lipid droplets from 29–33 cells were tested per genotype.

Statistical significance of differences between the lipid droplet area sizes of controls and Akh^A mutants were tested using the Mann–Whitney test with OriginPro 9.1.0.

Ex vivo confocal laser scanning microscopy and quantification of corpora cardiaca cells

For analysis of GFP expression, samples were handled as described by Pitman et al. (2011). Adult male and female flies were collected 6 days after eclosion and RCs were dissected in ice-cold phosphate-buffered saline (1.86 mM NaH₂PO₄, 8.41 mM Na₂HPO₄, 175 mM NaCl) containing 4% paraformaldehyde and fixed for 120 min under vacuum at room temperature. Samples were washed three times for 10 min in PBS containing 0.1% Triton X-100 and twice in PBS before mounting in glycerol with 20% PBS. Imaging was performed on a Zeiss LSM Meta 510 microscope and images were processed in Amira 5.4 (FEI, Hillsboro, OR). Resulting image stacks were contrast adjusted and the cell numbers were independently counted by two experimenters.

Starvation resistance assay, lipid and glycogen mobilization upon starvation

Seven-day-old flies (three to five replicates per genotype, 25–30 flies per replicate) were transferred to 0.6% agarose (in water) and the dead flies were scored every 7–10 hr. The statistical significance of differences between starvation survival curves was analyzed by log-rank test. The mobilization of energy stores upon starvation was analyzed by lipid and glycogen measurements on five to six replicates of five flies each per genotype and starvation time point as described above. Experiments were repeated in at least two independent trials. Effect of the genotype was analyzed by one-way ANOVA followed by Tukey´s HSD post hoc test. To test for the effects of genotype, duration of starvation exposure, and their interactions, two-way ANOVA was used.

Food intake assay by food labeling

Seven-day-old mated female flies were transferred to fly food medium containing 0.04% bromophenol blue and supplied with or without 20 mM paraquat for 4 hr. At this point in time, the number of flies with blue dye in the abdomen was scored by visual inspection. The assay was done as a blinded experiment, meaning that the genotypes were anonymized to the experimenter during scoring. The experiment was repeated at least twice, with 30–80 flies per genotype and tested time point. Data were analyzed for statistical significance by two-tailed Fischer exact test.

Paraquat resistance assay: application of paraquat on the nerve cord

The assay was done according to Cassar et al. (2015) with minor modifications. For a more detailed description see File S1.

Locomotor activity assay

Spontaneous activity of ad libitum–fed flies and starvation-induced hyperactivity of starved flies was tested using the Drosophila Activity Monitor 2 system (TriKinetics). Flies were briefly anesthetized with CO₂ and loaded individually into the monitoring tubes containing standard medium or 0.6% agarose (in water). Spontaneous locomotion was recorded over the first week of life. Test of the starvation-induced hyperactivity was started on 7-day-old flies; in parallel, siblings of these flies were monitored under ad libitum feeding conditions. For each genotype and feeding condition, 32 male flies were analyzed. Experiments were repeated at least twice. Spontaneous locomotor activity was measured as total number of midline crossings over the first week of life. Starvation-induced hyperactivity was analyzed both by visual inspection of the locomotion patterns of individual flies and by counting the total number of midline crossings over the complete starvation period and during the last 12 hr pre mortem. Quantitative data were analyzed by one-way ANOVA followed by Tukey´s HSD post hoc test.

Startle-induced vertical climbing

The climbing assay is based on the “countercurrent distribution” method described by Benzer (1967) with modifications. For a more detailed description see File S1.

Flight performance assay

Flight performance analysis was based on the assay developed by Benzer (1973) and modified by Babcock and Ganetzky (2014). For a more detailed description see File S1.

Data availability

All fly strains generated in this work (see Supporting Information for details) are available on request.

Results

Generation of AKH single mutant and AKH plus APRP double mutant flies by CRISPR/Cas9-mediated genome engineering

The Akh gene encodes a prohormone that gives rise to the mature AKH and APRP peptides after signal peptide removal and proteolytical processing (Figure 1A). To study the developmental and metabolic functions of AKH and APRP, we employed CRISPR/Cas9-assisted mutagenesis to create an AKH-specific mutant (Akh^A) and AKH plus APRP double mutant flies (Akh^AP and Akh^SAP) (Figure 1, A and B). Mutations were obtained according to Kondo and Ueda (2013) by mutagenesis in the male germline expressing Akh-targeting gRNA, UAS-Cas9, and nanos-GAL4 (Port et al. 2014). The Akh^A allele represents an AKH-specific in-frame deletion of the two C-terminal amino acids of the AKH octapeptide, which leaves the APRP sequence unaffected (Figure 1, A and B). MS analysis on RC peptides confirmed the presence of the predicted AKH hexapeptide and its processing intermediates in Akh^A mutant flies (Figure 2). Moreover, Akh^A mutants express APRP peptide dimers indistinguishable from Akh⁺ control flies (Figure 2, Figure S1, and Figure S2). In contrast, MS analysis detected no peptides encoded by the Akh gene in the Akh^AP and Akh^SAP mutants or in flies subject to an Akh RNAi knockdown, while the profile of other RC peptides was unchanged (Figure 2). This is consistent with the molecular identity of the Akh^AP and Akh^SAP mutants. The Akh^AP allele carries a 19-bp deletion in the AKH region, which causes a frameshift upstream of the APRP coding sequence (Figure 1, A and B). Similarly, the 206-bp deletion in the Akh^SAP allele also removes the AKH coding sequence along with the signal peptide sequence and the translation initiation side of the prohormone (Figure 1, A and B). Collectively, the sequencing and peptide MS data identify Akh^AP and Akh^SAP as specific AKH plus APRP double loss-of-function mutants. We also propose Akh^A to be a AKH-specific null allele of Akh as the AKH^A hexapeptide lacks the tryptophan at position 8, which was shown to be essential for Drosophila AKH receptor activation by structure–activity studies (Caers et al. 2012).

Figure 2.

Characterization of AKH single and AKH plus APRP double mutants by mass spectrometry. Comparison of recorded MALDI-TOF mass spectra of single retrocerebral complex preparations from Akh⁺ (black) control, Akh mutants (Akh^A in blue, Akh^AP in red, Akh^SAP in green) and Akh RNAi knockdown (gray) flies in the mass ranges m/z 700–1340 (A) and m/z 10,220–10,320 (B). In the control flies, the full set of Akh gene products (black labels) was detected (see panel C and Figure 1). Deletion of the codons for the two C-terminal AKH amino acids DW in Akh^A mutants resulted in truncated AKH products (blue labels; pQLTFSPa, 696.3 [M+Na]⁺; pQLTFSPGK-OH, 860.4 [M+H]⁺, 882.3 [M+Na]⁺; pQLTFSPGKR-OH, 1016.4 [M+H]⁺), leaving the APRP sequence unaffected. Akh^AP flies, which carry a deletion causing a frame shift mutation, and Akh^SAP flies, which miss the signal peptide coding for the sequence including the Akh translation start codon, showed no ion signals for putative Akh translation products in all analyzed ranges. Also Akh RNAi flies lack the AKH and the peptides. Ion signals from neuropeptides of other genes (pink peak labels) were unaffected in all genotypes analyzed and served as marker (Dm-sNPF-1^4–11/sNPF-2^12-19, 974.6 [M+H]⁺; Dm-sNPF-3, 982.6 [M+H]⁺; Dm-sNPF-4, 985.6 [M+H]⁺; Dm-MS 1247.7, [M+H]⁺; Dm-sNPF-1, 1329.8 [M+H]⁺). All spectra were recorded in reflectron positive mode and all ion signals are labeled with monoisotopic masses. Signal intensities were scaled (100%) to control fly pQLTFSPDWGK-OH [M+H]⁺ signal (1161.6 m/z) in A and to the dimer [M+H]⁺ signal (10292.0 m/z) in B. Note that peak labels annotated in A and B are bold in C.

Thus, the new Akh mutants allow addressing AKH-specific functions (revealed by the Akh^A) and APRP-specific functions (revealed by the comparison of Akh^AP and Akh^A phenotypes). Moreover, comparison of Akh^A mutant phenotypes with phenotypes of the AKH receptor deletion mutant AkhR¹ (Grönke et al. 2007) can support or challenge the view of a single ligand/receptor pair in Drosophila AKH signaling.

AKH signaling and APRP are dispensable for developmental and fitness-related functions in Drosophila

AKH signaling is regarded as the central regulator in systemic energy mobilization control acting antagonistically to insulin signaling. Accordingly, we first tested the Akh dependency of nonfeeding ontogenetic stages (i.e., embryogenesis and metamorphosis) and of biosynthetically demanding processes such as oogenesis. As developmental and physiological traits are sensitive to confounding genetic background effects, all AKH pathway and APRP mutants were crossed into a common w¹¹¹⁸ background for nine generations prior to the phenotypic analyses.

Comparative analysis of Akh^A, Akh^AP, AkhR¹, and the genetically matched control flies revealed no gross abnormalities in hatchability (i.e., egg to L1 larval survival; Figure 3A), viability (i.e., L1 larval to adult survival; Figure 3B), developmental time (egg to adult; Figure 3C), or female fecundity (Figure 3D) in flies lacking AKH signaling. Moreover, the body size of AKH signaling mutants was unaffected (Figure 3E, Figure S3). We noticed slightly, but significantly reduced hatchability in Akh^A. However, this effect was not caused by the deficiency in the AKH signaling itself, as the AkhR mutant and AKH plus APRP double mutants had normal hatchability and larval to adult survival (Figure 3, A and B).

Taken together, in contrast to the dramatic effects of insulin signaling deficiency (Grönke et al. 2010), absence of AKH does not cause gross abnormalities in development or reproduction.

AKH signaling and APRP are dispensable for mobilization of lipids and glycogen during Drosophila development

Development encompasses nonfeeding periods like embryogenesis, moltings, and metamorphosis, when the organism relies exclusively on stored energy reserves. To address the fuel utilization during Drosophila metamorphosis, and to test a possible involvement of AKH in this process, we followed the glycogen and lipid changes in Drosophila from the wandering stage of the third instar larvae (3L, end of the feeding) through pupation (P0), termination of the pupal development (P13–P15), until the posteclosion values in the immature adults (within 10 hr after eclosion) (Figure 4, A and B). As expected, the developmental stage had a highly significant effect on the amount of lipid reserves (see statistical analysis for Figure 4A). Lipid stores increased shortly before initiation of metamorphosis, reaching their highest values at early pupation and gradually decreased afterward (Figure 4A). In contrast to lipids, the glycogen content already reached its maximum at the 3L larval stage, decreased toward the P0 stage, fell to very low levels at P13–P15, and increased within the first day after eclosion (Figure 4B).

Figure 4.

Developmental changes of carbohydrate and lipid stores in AKH single, AKH plus APRP double, AkhR mutants, and their genetic control. Plotted are means ± SEM. (A) Lipid content (expressed as glycerides). No difference is seen in the lipid content among the tested genotypes at the stage of wandering third instar larvae (3L) (one-way ANOVA, F_3,20 = 1.38, P = 0.28), or at the beginning of metamorphosis at P0 (one-way ANOVA, F_3,14 = 3.1, P = 0.06), or at the end of pupal development at P13–P15 (one-way ANOVA, F_3,16 = 2.56, P = 0.43). Note the nonsignificant trend toward increased lipid content in the Akh and AkhR mutants at the first day after adult eclosion (one-way ANOVA, F_3,16 = 3.28, P = 0.05). Strong lipid mobilization is observed in all genotypes during metamorphosis; no significant interaction is observed between the genotype and the effect of the developmental stages (two-way ANOVA, genotype and developmental stage as fixed effects, genotype: F_3,66 = 2.94, P = 0.04, developmental stage: F_3,66 = 119.9, P < 0.001, genotype × developmental stage: F_9,66 = 1.86, P = 0.07). (B) Glycogen content. No significant difference is observed in the glycogen content in the mutants at the stage of wandering third instar larvae (3L) (one-way ANOVA, F_3,20 = 3.67, P = 0.03), or at the beginning of metamorphosis at P0 (one-way ANOVA, F_3,20 = 2.94, P = 0.06), or at the end of pupal development at P13–P15 (one-way ANOVA, F_3,16 = 2.57, P = 0.09). Note the statistically nonsignificant trend toward lower glycogen levels in all AKH signaling mutants compared to the controls. Strong glycogen mobilization during metamorphosis is observed; no interaction between the genotype and tested developmental stages is observed (two-way ANOVA, genotype and developmental stage as fixed effects, genotype: F_3,75 = 3.87, P = 0.012, developmental stage: F_3,75 = 51.92, P < 0.001, genotype × developmental stage: F_9,75 = 0.76, P = 0.65).

In analogy to AKH functions in nondrosophilid insects, Drosophila AKH can be anticipated to regulate synthesis of fat stores before metamorphosis, and their utilization during this process. However, Akh^A, Akh^AP, or AkhR¹ accumulated comparable amount of lipids at both 3L larval stage and P0 pupal stage. Similarly, lipid mobilization proceeded normally in all tested genotypes, and lipid stores were comparable between all mutants and control at the end of metamorphosis (stage P13–P15) (Figure 4A). There was a nonsignificant trend toward increased lipid levels in freshly eclosed flies (Figure 4A); however, this phenomenon is likely not connected with the metamorphosis itself, but rather foretells the adult-specific role of AKH signaling, as described in Deficiency in AKH-signaling results in adult-onset obesity and hypoglycemia. Altogether, deficiencies of AKH signaling or APRP had no effect on the lipid content at any tested developmental stage or on the storage lipid mobilization during metamorphosis (see statistical analyzes in Figure 4A).

Adipokinetic hormone was described to act hypertrehalosaemic in several insect species including Drosophila larvae (Kim and Rulifson 2004; Lee and Park 2004). Thus, we hypothesized that the loss of AKH function might result in impaired glycogen mobilization, and consequently in higher body glycogen levels. However, similarly to lipid reserves, glycogen profiles of Akh^A, Akh^AP, and AkhR¹ did not differ from the profiles of controls at any of the tested developmental stages (Figure 4B). We observed a nonsignificant trend toward lower glycogen storage in Akh^A, Akh^AP, and AkhR¹ mutants at the 3L and P0 stages (Figure 4B). Lowered glycogen storage reached statistical significance if data from different developmental time points were analyzed together, with the genotype and developmental stage as fixed effects (see statistical analyzes in Figure 4B). However, despite the trend toward lower glycogen storage at the onset of metamorphosis, the following mobilization of glycogen was not affected, and there was no interaction between the effect of genotype and developmental stage (see statistical analyzes in Figure 4B).

Thus, the above-presented experiments show that in Drosophila, AKH pathway and APRP are not required for the dynamic changes in storage lipids and carbohydrates, which are characteristic for metamorphosis.

Deficiency in AKH signaling results in adult-onset obesity and hypoglycemia

After the analyses of the roles of AKH signaling and APRP during metamorphosis, we focused on their potential metabolic functions at the adult stage of Drosophila. Interestingly, Akh^A, Akh^AP, and AkhR¹ mutants already developed adult-specific obesity within the first week after eclosion (Figure 5A). Thin layer chromatography confirmed that among the neutral lipids, storage TGs were particularly increased in the obese AKH signaling mutants (Figure 5B). When examining the age-dependent changes in the fat content during the first week of adult life (the first day after eclosion vs. 1 week later), we noticed that the lipid content of controls dramatically decreased (Figure 4A and Figure 5A). This drop in storage lipids corresponds to the histolysis of the larval/pupal fat body cells during and shortly after the metamorphosis and their functional replacement by the adult fat body cells (Nelliot et al. 2006). In contrast to the control flies, lipid content of mature Akh^A, Akh^AP, and AkhR¹ mutants remained at the posteclosion levels (Figure 4A and Figure 5A). This observation raised the question of whether the obesity of the AKH signaling mutants is based on defective clearance of larval/pupal fat body cells or on excessive lipid loading of adult fat body cells. These types of fat body are morphologically distinguishable, and thus we examined the fat body composition of 1-week-old mutants. However, dissection, staining, and confocal imaging of fat body suggests that the obesity resulted from increased cellular lipid loading of adult fat body cells (Figure 5, C and D). Altogether our data showed that AKH deficiency results in adult-onset obesity coupled with adult fat body cell hypertrophy.

Figure 5.

Adult-onset obesity and hypoglycemia in AKH signaling mutants. Shown are carbohydrate and lipid levels in 7-day-old AKH single, AKH plus APRP double, and AKHR mutants. Plotted are means ± SEM. (A) Deficiency in AKH signaling resulted in adult-onset obesity; magnitude of the phenotype was the same for Akh^A, Akh^AP, and AkhR¹ (one-way ANOVA, F_3,20 = 23.84, P < 0.001; Tukey’s HSD: P < 0.05). (B) TLC analysis illustrated that the Akh^A, Akh^AP, and AkhR¹ mutants predominately accumulate triglycerides (TGs) (FA, fatty acids; DG, diacylglyceride; MG, monoacylglyceride; S, standard. Note: As indicated by the dashed line between the control and AkhR¹ lanes, an unrelated sample has been removed from the TLC plate image). (C) Fat body cell hypertrophy in AKH signaling mutants as illustrated by confocal imaging of increased cellular lipid loading. Lipid droplets are shown in green (Bodipy493/503), cell membranes in red (CellMask Deep Red), and nuclei in blue (DAPI). Bar, 25 μm. (D) Quantification of the lipid droplet (LD) area per fat body cell. Mann–Whitney test, P < 0.001, n = 33 cells (Akh^A) and n = 29 cells (control). (E) Nonsignificant decrease in glycogen levels in Akh^A, Akh^AP, and AkhR¹ mutants compared to controls (one-way ANOVA, F_3,20 = 2.49, P = 0.091). (F) Hypoglycemia of Akh^A, Akh^AP, and AkhR¹ mutants as revealed by circulating sugar quantification (one-way ANOVA, F_3,8 = 9.97; P < 0.01; Tukey’s HSD: P < 0.05).

Next, we tested whether the stored carbohydrates, i.e., glycogen, also increased in response to the loss of AKH signaling. However, this was not the case. On the contrary, we observed a nonsignificant trend toward decreased glycogen values in all Akh^A, Akh^AP, and AkhR¹ mutants (Figure 5D). Next to the stored carbohydrates, we tested also the free circulating sugars (trehalose and glucose). When analyzing whole body samples, we were not able to detect any significant difference between the mutants and controls (data not shown), whereas hemolymph samples revealed a significant hypoglycemia in all Akh^A, Akh^AP, and AkhR¹ mutants (Figure 5E).

Altogether, these data show that AKH signaling fulfills important functions in the homeostasis of stored lipids and of circulating, but not stored carbohydrates in mature adult Drosophila flies.

AKH signaling and APRP are dispensable for spontaneous locomotor activity, startle-induced climbing, and flight performance

Mobilization of energy reserves to sustain locomotion is one of the main general functions of AKHs. However, when testing spontaneous locomotion in the absence of AKH, APRP, and AKHR over a 1-week period of time, we did not find any significant reduction of the locomotion in the obese and hypoglycemic Akh^A, Akh^AP, and AkhR¹ mutants (Figure 6A). Next, we addressed a potential role of AKH signaling or APRP in forced locomotion using a startle-induced climbing paradigm. Climbing of Akh^AP and AkhR¹ mutants was indistinguishable from controls, while the Akh^A flies showed reduced climbing performance (Figure 6B). However, this effect was unlikely caused by the absence of AKH signaling, as the receptor mutant and AKH plus APRP double mutants climbed normally. Loss of AKH signaling and APRP also did not affect flight performance (Figure 6C).

Figure 6.

No requirement of AKH signaling and APRP for spontaneous locomotor activity, or for startle-induced vertical climbing, or for flight performance. (A) No spontaneous locomotion defects of Akh^A, Akh^AP, or AkhR¹ compared to controls as revealed by cumulative activity monitoring for 1 week using the DAM2 system (one-way ANOVA, F_3,29 = 10.11, P = 0.04); Tukey’s HSD: P < 0.05). (B) Reduced startle-induced climbing ability of Akh^A but not Akh^AP or AkhR¹ compared to controls (one-way ANOVA, F_3,29 = 10.11, P < 0.001; Tukey’s HSD: P < 0.05). (C) No defects in flight performance of Akh^A, Akh^AP, or AkhR¹ (one-way ANOVA, F_3,123 = 1.28, P = 0.29).

AKH signaling contributes to the mobilization of lipids under starvation

Mobilization of energy reserves in periods of negative energy balance is the main function of AKH hormones. We used extended food deprivation to address the energy mobilization capacity of flies lacking AKH, APRP, or AKHR. Consistent with their higher body fat content, Akh^A, Akh^AP, and AkhR¹ mutants were all more starvation resistant than controls (Figure 7A). Monitoring of lipid and carbohydrate stores during starvation revealed that all three mutants mobilized both storage energy sources (Figure 7, B and C). However, analysis of the interactions between the genotype and the starvation-dependent lipid changes revealed a significant interaction between these two factors (Figure 7B), suggesting that the lack of AKH function modulates the lipid mobilization profile. In contrast to the controls, Akh^A, Akh^AP, and AkhR¹ mutants were not able to mobilize their lipid reserves completely, and thus died with higher residual lipid content (Figure 7B).

Figure 7.

AKH signaling regulates the starvation response. (A) Increased starvation resistance of Akh^A, Akh^AP, and AkhR¹ compared to control (log-rank test, Akh^A vs. control: χ² = 137.63, P < 0.001; Akh^AP vs. control: χ² = 131.66, P < 0.001; AkhR¹ vs. control: χ² = 135.53, P < 0.001; n[Akh^A] = 82, n[Akh^AP] = 95, n[AkhR¹] = 105, n[control] = 110). (B) Functional but impaired storage lipid mobilization of obese Akh^A, Akh^AP, and AkhR¹ mutants as revealed by strong interaction between the genotype and starvation duration (two-way ANOVA, genotype and starvation time as fixed effects, genotype: F_3,100 = 237.7, P < 0.001, starvation time: F_4,100 = 389.6, P < 0.001, genotype × starvation time: F_12,100 = 14.31, P < 0.001). Note that Akh^A, Akh^AP, and AkhR¹ mutants did not mobilize lipid reserves completely in contrast to controls (total starvation, TS, one-way ANOVA, genotype as fixed effect, F_3,20 = 7.39, P < 0.05; Tukey’s HSD: P < 0.05). (C) Functional but impaired glycogen storage mobilization of Akh^A, Akh^AP, and AkhR¹ mutants. Note that glycogen content did not differ from each other nor from the controls at any individual time point tested (one-way ANOVA, genotype as fixed effect, 0 hr: F_3,20 = 2.49, P = 0.09; 8 hr: F_3,18 = 2.61, P = 0.08; 16 hr: F_3,20 = 2.45, P = 0.09; 24 hr: F_3,18 = 2.54, P = 0.09), but AKH signaling deficiency had a significant effect on the glycogen starvation response (two-way ANOVA, genotype and starvation time as fixed effects, genotype: F_3,76 = 3.09, P = 0.03, starvation time: F_3,76 = 152.8, P < 0.001, genotype × starvation time point: F_9,76 = 2.44, P < 0.05). (D) No difference in the starvation lifetime locomotor activity between all genotypes (one-way ANOVA, genotype as fixed effect, F_3,123 = 1.5, P = 0.22). Note that since all mutants are starvation resistant (A), average locomotor activity per day is reduced compared to controls. (E) Representative figure showing the locomotor activity distribution in AKH-deficient and control flies during starvation (TS) compared to ad libitum-fed siblings (AL). Note the starvation-induced hyperactivity shortly before death of control flies when compared to ad libitum fed siblings. Starvation-induced hyperactivity is absent in Akh^A, Akh^AP, and AkhR¹ mutants (last 12 hr of life are highlighted in black rectangle). Akh^A AL = mean locomotor activity of Akh^A siblings (n = 32) fed on standard medium. Akh^A TS = locomotor activity of representative individual Akh^A male on total starvation. Control AL = mean locomotor activity of control siblings (n = 31) fed ad libitum on standard medium. Control TS = locomotor activity of representative individual control male on total starvation. Bar below the graphs illustrates the 12-hr light (yellow)/12-hr dark (blue) cycle. (F) Quantitative analysis of the pre mortem locomotor activity supports the absence of starvation-induced hyperactivity in Akh^A, Akh^AP, and AkhR¹ mutants (one-way ANOVA, genotype as fixed effect, F_3,122 = 47.78, P < 0.001; Tukey’s HSD: P < 0.05).

In contrast to the storage lipids, the glycogen reserves of ad libitum fed Akh^A, Akh^AP, and AkhR¹ mutant flies were not increased. All mutants were able to mobilize glycogen stores at a rate comparable with controls (Figure 7C). This suggests that increased starvation survival of Akh^A, Akh^AP, and AkhR¹ was predominantly driven by the increased lipid reserves. Next, we tested whether potential changes in locomotion under starvation might contribute to the differential survival of the Akh^A, Akh^AP, and AkhR¹ mutants.

AKH signaling promotes starvation-induced hyperactivity

Locomotion adds to the negative energy balance during starvation and therefore reduces the survival time under food deprivation. Hence, we aimed to test whether the lack of AKH function contributes to starvation resistance of Akh^A, Akh^AP, and AkhR¹ via inducing hypoactivity under nutritional shortage. Indeed, total starvation lifetime locomotion of long-lived Akh^A, Akh^AP, and AkhR¹ mutant flies was unchanged compared to the short-lived controls (Figure 7D). Accordingly, the average locomotion per hour of starvation lifetime was reduced in the absence of AKH signaling.

As described previously, locomotion of starved control flies shortly before death typically exceeded the activity of ad libitum–fed siblings, reflecting a behavioral strategy of flies during extended food deprivation, which is interpreted as food-seeking behavior. This starvation-induced hyperactivity was abolished in CC-ablated flies (Lee and Park 2004), suggesting that AKH, APRP, or another factor produced in these cells is required for the process. By visual inspection of the locomotory patterns of individual flies, we noted that the starvation-induced hyperactivity was suppressed in all Akh^A, Akh^AP, and AkhR¹ mutants (Figure 7E). Quantification of the mean activity of individual flies during their last 12 hr of starvation survival confirmed the dramatic decrease of locomotion of Akh^A, Akh^AP, and AkhR¹ compared to controls (Figure 7F). We did not observe any significant differences between the AKH- and AKHR-deficient mutants (Figure 7F).

Taken together, these data show that AKH is necessary for the starvation-induced hyperactivity and that the AKH signal is transduced via the canonical AKHR receptor. Since hypoactivity is an energy-saving strategy, it might contribute to the starvation resistance of the Akh^A, Akh^AP, and AkhR¹ mutants.

AKH signaling confers oxidative stress resistance

Food deprivation is one form of metabolic stress commonly experienced by Drosophila in natural environments. AKH has also been implicated in coping with other forms of stress conditions such as oxidative stress (e.g., Kodrík et al. 2007; Večeřa et al. 2007). Exposure to foodborne paraquat showed apparently increased oxidative stress resistance of Akh^A, Akh^AP, and AkhR¹ mutants (Figure S4). However, a food-intake assay revealed increased paraquat-induced food aversion of Akh^A, Akh^AP, and AkhR¹ mutants (Figure 8A). As reduced paraquat intake could be causative for the observed apparent oxidative stress resistance of Akh signaling mutants, we switched to application of paraquat directly on the nerve cord. This food intake independent drug application revealed increased paraquat sensitivity of AKH-deficient flies (Figure 8B), suggesting a protective role of AKH in coping with oxidative stress.

Figure 8.

AKH signaling regulates the oxidative stress response. (A) Significantly reduced intake of paraquat-supplemented food in Akh^A, Akh^AP, and AkhR¹ mutants compared to controls challenges foodborne paraquat as a suitable measure to test oxidative stress resistance. Food intake was assayed using blue dye labeling of food supplemented with (+) or without (−) 20 mM paraquat for 4 hr prior to visual inspection of abdominal coloring. Fischer exact test, Akh^A vs. control: P < 0.001; Akh^AP vs. control: P < 0.001; AKHR¹ vs. control: P < 0.001; no significant difference among Akh^A, Akh^AP, and AkhR¹ (n[Akh^A] = 66, n[Akh^AP] = 64, n[AkhR¹] = 70, n[control] = 62). Note that differential food intake resulted from differential aversion to paraquat, as there was no difference among the genotypes when fed on regular food (Fischer exact test, for all comparisons, P > 0.05, (n[Akh^A] = 80, n[Akh^AP] = 81, n[AkhR¹] = 73, n[control] = 81). (B) Direct application of paraquat to the nerve cord revealed oxidative stress sensitivity of Akh^A, Akh^AP, and AkhR¹ mutants compared to controls. Fischer exact test, Akh^A vs. control: P < 0.001; Akh^AP vs. control: P < 0.001; AkhR¹ vs. control: P < 0.05; Akh^A vs. Akh^AP: P = 0.74; Akh^A vs. AkhR¹: P = 0.04; Akh^AP vs. AkhR¹: P = 0.12 (n[Akh^A] = 89, n[Akh^AP] = 93, n[AkhR¹ = 91, n[control] = 90).

Taken together, our data on nutritional and oxidative stress resistance suggest that AKH orchestrates metabolic changes in flies challenged with environmental stress factors.

The Akh gene is controlled by negative autoregulation

Homeostatic modulation of metabolism in response to changing environments requires prompt feedback regulation. Endocrine systems in general involve negative feedback loops controlling the hormone production. However, nothing is known about autoregulation of AKH levels. Glucagon, the mammalian homolog of AKH, negatively regulates its own production as revealed by compensatory overproliferation of glucagon-positive pancreatic alpha cells (Gelling et al. 2003). To test whether AKH acts in an analogous manner, we visualized the CC cells of Akh⁺, Akh^A, and Akh^AP flies by expressing GFP under indirect control of the AKH promoter (akhp-Gal4 > UAS-mCD8 GFP). The number of CC cells did not differ in any of the tested mutants (Figure 9A and Figure S5), suggesting that in contrast to its mammalian functional homolog, the insect hormone does not feed back to the cells of AKH origin via proliferation control. Interestingly, we observed a significant increase of GFP signal intensity in a subset of CC cells in AKH-deficient mutants (Figure 9, B and C), suggesting an increased Akh promoter activity in response to the lack of AKH. Consistently, the Akh messenger RNA (mRNA) abundance was increased in the Akh^A mutants (Figure 9D). These data provide first evidence that Akh is subjected to negative autoregulation at the mRNA level. Future studies will address how the AKH levels are sensed and which factors contribute to the proposed regulatory loop.

Figure 9.

AKH deficiency does not affect corpora cardiaca cell number but reveals a negative autoregulatory loop on Akh transcription. Fluorescence tagging of CC cells under indirect control of the Akh promoter (akhp-Gal4 > UAS-mCD8 GFP) detected no change in CC cell number between Akh⁺ controls and in Akh^A mutants (A; two-tailed Student’s t-test: P = 0.34; Akh⁺ n = 10 flies, Akh^A n = 9 flies) but increased somatic GFP signal in a subset of CC cells (arrowheads in the maximum intensity projection in B, quantification in C; two-tailed Student’s t-test: P = 0.004 Akh⁺ n = 160 cells, Akh^A n = 140 cells). (D) Increased Akh mRNA abundance in Akh^A mutants compared to Akh⁺ controls as revealed by qPCR (two-tailed Student’s t-test: P < 0.001; n = 3 biological replicates per genotype).

Discussion

AKH signaling has been initially described as a master regulator of insect energy mobilization in a broad range of biological contexts. This view has been shaped by numerous elegant studies on the effects of ectopic application of AKH and on correlations between the hormone titer and insect physiology (metabolism, developmental stage, behavior) in diverse insect species (reviewed in Vroemen et al. 1998; Van der Horst et al. 2001; Gäde and Auerswald 2003; Van der Horst 2003; Lorenz and Gäde 2009). However, data on the consequences of selective impairment of AKH function are scarce. Here, we conducted the first comparative loss-of-function analysis of Drosophila AKH mutants and AKH plus APRP double mutants. These specific mutants, together with the AKH receptor mutant in the same genetic background, allowed testing of developmental, reproduction-, stress-, and metabolism-related functions of AKH signaling, its dependency on the AkhR, as well as addressing the putative hormonal roles of the functionally enigmatic APRP. We discuss below our main results in the context of general predictions of AKH roles in insects, which have so far been based mainly on AKH studies in nondrosophilid species.

Developmental roles of AKH signaling

Insect embryogenesis, as a nonfeeding stage, is dependent on mobilization of maternally supplied lipid reserves (Beenakkers et al. 1985; Arrese and Soulages 2010). Intriguingly, Drosophila Akh and AkhR are already expressed at embryogenesis (Kim and Rulifson 2004; Lee and Park 2004; Grönke et al. 2007), suggesting that AKH might have biological functions in energy mobilization from early developmental stages onwards. However, our data show that AKH, APRP, and AKHR are dispensable for embryonic development.

Next to embryogenesis, nonfeeding periods of insect life cycle also include moltings and metamorphosis. Metamorphosis involves rebuilding of larval structures into adult body, and this process completely depends on oxidizing energy stores accumulated during the larval period (Agrell and Lundquist 1973). A general role of AKH in this process has been predicted based on, e.g., experimental evidence that AKH injections cause differential glycogen or lipid mobilization in a developmental stage-specific manner in Manduca sexta (Gäde and Beenakkers 1977) or Zophobas atratus (Slocinska et al. 2013; Gołębiowski et al. 2014). Preference for lipids or glycogen as energy sources and dynamics of their usage during metamorphosis vary considerably among species (Nestel et al. 2003; Dutra et al. 2007). Similarly, as recently described by Matsuda et al. (2015), we also noticed that metamorphosis of Drosophila is fueled to a considerable extent by glycogen, but lipids are mobilized as well. We monitored the effect of AKH signaling deficiency upon glycogen and lipid levels at several time points before and during Drosophila metamorphosis to detect potential differences in the dynamics of lipid and glycogen mobilization in the absence of AKH signaling or APRP. Consistent with the previously described absence of any effects of CC ablation on larval lipids (Lee and Park 2004), we did not detect significant changes in the larval lipid content in any of the Akh^A, Akh^AP, and AkhR¹ mutants. Larval glycogen values also did not statistically differ from controls, meaning that AKH deficiency did not affect starting levels of energy sources at the onset of metamorphosis. Measurements of glycogen and lipids throughout metamorphosis excluded any differences in the energy mobilization of the tested mutants. Thus, Drosophila AKH signaling is dispensable for proper accumulation of energy stores at larval stage, as well as for their mobilization during metamorphosis. Consistent with the lack of metabolic phenotypes during metamorphosis, larval to adult survival of flies deficient in AKH signaling did not differ from the genetically matched controls. Thus, similarly to its mammalian functional homolog glucagon (Gelling et al. 2003), AKH is dispensable for developmental functions. Nevertheless, it is possible that other pathways compensate for absence of AKH, thus obscuring the detection of AKH-related developmental functions in the mutants. A prime candidate for such an alternative lipid mobilization pathway involves the Brummer lipase, as bmm mutants are embryonic lethal (Grönke et al. 2005), and the gene is known to work in parallel with AkhR in starvation-induced storage fat mobilization in Drosophila adults (Grönke et al. 2007). Alternatively, AKH signaling might instead have fine-tuning functions that would become obvious only under particular suboptimal conditions. Our study was conducted under a protected laboratory environment, with controlled diet, temperature, animal density, etc. However, development is highly plastic, and life history traits like developmental time, body size, fecundity, and viability are sensitive to environmental changes. AKH signaling was repeatedly connected with stress responses (Bednářová et al. 2013a), and thus, it might also play some context-dependent roles in adjusting metabolism and speed of development during metamorphosis. This hypothesis argues favorably for the recent finding that the developmental time is extended in the AkhR mutants, when raised on a low-yeast diet (Kim and Neufeld 2015).

AKH signaling controls lipid homeostasis in adult Drosophila

Metabolic pathways governing the energy balance during preadult and adult development might differ from those maintaining this balance during adulthood. Several genes such as inositol 1,4,5-tris-phosphate receptor Itp-r83A (Subramanian et al. 2013) or perilipin1 (Beller et al. 2010) act as antiobesity genes specifically at the adult stage of Drosophila. Here we show that this is also the case for AKH signaling. Obesity of Akh^A and Akh^AP mutants is in line with the previous reports on the effect of mutations in the AKH receptor (Grönke et al. 2007; Bharucha et al. 2008). Earlier data on the AKH roles in glycogen storage were rather contradictory, reporting both no changes (Grönke et al. 2007), as well as significant increase in the body glycogen content (Bharucha et al. 2008) of AKH receptor mutants. In contrast, the role of AKH in circulating sugars in adults was not addressed previously. In larva, however, CC cell ablation was shown to cause hypoglycemia (Kim and Rulifson 2004; Lee and Park 2004; Isabel et al. 2005). We were not able to detect any significant differences in the free sugars when analyzing whole body samples; however, when hemolymph samples were analyzed, we observed significant hypoglycemia in Akh^A, Akh^AP, and AkhR¹ mutants. This reduction of circulating sugars was not coupled with an increase in the stored carbohydrates. On the contrary, we observed the opposite trend toward lowered glycogen levels in Akh^A, Akh^AP, and AkhR¹ mutants. Therefore, increased uptake of circulating sugars and their subsequent use for lipogenesis is one hypothesis on the etiology of Akh^A, Akh^AP, and AkhR¹-dependent obesity to be tested in the future.

AKH signaling is not required for Drosophila reproduction

Insect reproduction is an energetically demanding process, as females deposit a considerable amount of energy reserves into the developing oocytes. Mobilization of energy reserves for oogenesis was predicted to be AKH regulated (Lorenz and Gäde 2009). Consistently, AKH is required for reproduction of tsetse fly Glossina morsitans (Attardo et al. 2012), but on the contrary, AKH prevents vitellogenesis and egg production in the locust Locusta migratoria (Glinka et al. 1995) and the cricket Gryllus bimaculatus (Lorenz 2003). Accordingly, relevance and mode of action of AKH signaling in insect oogenesis appears to vary considerably among species. This diversity of AKH functions is also reflected by the differential expression of the AKH receptor in ovaries. For example, AKHR is expressed in the ovaries of the mosquito Aedes aegypti (Kaufmann et al. 2009), but not in those of Drosophila (Grönke et al. 2007; Bharucha et al. 2008). In the current study, we did not detect any changes in fecundity in the absence of AKH signaling, suggesting that in Drosophila, oogenesis-dependent fat mobilization is under the control of an alternative lipolytic pathway. However, we cannot exclude that AKH plays a role in reproduction and vitellogenesis under natural conditions, which likely requires more metabolic adaptability to environmental changes.

Roles of AKH in locomotion

It is widely accepted that AKH has an important regulatory function in insect locomotion (Lorenz 2003; Van der Horst 2003; Lorenz and Gäde 2009). This view is supported by many studies describing correlations between the release of AKH and activities like flight and walking (Lorenz 2003), and on experimental increase of locomotion, such as walking of the firebug Pyrrhocoris apterus, by ectopic applications of AKH (Kodrík et al. 2000, 2002). Ablation of CC cells in Drosophila decreased spontaneous locomotion (Isabel et al. 2005). However, this effect was likely caused by other factors produced in CC cells, as the Akh^A, Akh^AP, and AkhR¹ mutants tested in this study had normal spontaneous locomotion. Thus, neither the AKH signaling deficiency nor the resulting obesity affected spontaneous movement. This suggests that either the locomotion-related roles of AKH signaling are not evolutionarily conserved or that the regulatory potential of AKH as demonstrated by gain-of-function studies is not exploited by Drosophila under laboratory environmental conditions.

AKH signaling, together with octopamine, was also hypothesized to act analogously to vertebrate adrenaline during the “flight or fight” reaction (Lorenz and Gäde 2009). However, we did not detect any defects in the startle-induced climbing of AKH signaling mutants. Thus, other pathways exist to ensure the energy supply for this kind of movement in Drosophila.

Roles of AKH in the starvation response

Periods of starvation are coupled with rapid mobilization of energy reserves (Arrese and Soulages 2010). Starvation-induced mobilization of lipids in Rhodnius prolixus has been recently shown to be dependent on the AKH receptor (Alves-Bezerra et al. 2015). Given the hyperglycemic and hyperlipaemic effects of AKH in adult insects subjected to negative energy balance (Gäde and Auerswald 2003; Van Der Horst 2003; Lorenz and Gäde 2009), we studied Drosophila AKH functions during starvation. Akh^A, Akh^AP, AkhR¹ mutants were considerably more resistant to starvation than their genetically matched controls. Lipids were mobilized in all Akh^A, Akh^AP, and AkhR¹ mutants, however, to a lower extent than in controls, resulting in higher residual lipids in flies starved to death. These data are consistent with the earlier finding that AKHR cooperates with a second lipolytic pathway involving the Brummer lipase to orchestrate starvation-induced storage lipid mobilization (Grönke et al. 2007). Nevertheless, in the context of starvation, this alternative lipolytic pathway compensates for AKH absence only to a limited extent, as lipid mobilization is not completed at the time of death.

In contrast to lipids, the glycogen-mobilization response of all Akh^A, Akh^AP, and AkhR¹ mutants was similar to controls during the first 24 hr of starvation, when glycogen reserves became almost completely depleted. Accordingly, AKH signaling mutants survive a considerably long period without glycogen reserves, fueling their metabolism by stored lipids.

Prolonged starvation increases foraging behavior, which can be observed as increased locomotion that exceeds the activity of ad libitum–fed flies and overwrites the circadian activity pattern. In Drosophila, CC cell ablation suppresses this hyperactivity (Lee and Park 2004), consistent with the view that this behavior requires Akh gene products or other CC cell-produced factors. On the contrary, locomotion of AkhR mutants under starvation was described as identical to controls (Bharucha et al. 2008). In our study, both visual inspections of locomotory patterns of individual flies under starvation, as well as quantification of their total activity shortly before death revealed that in contrast to control flies, starvation failed to increase locomotion in Akh^A and Akh^AP but also AkhR¹ mutants. Thus, starvation triggers increase in locomotion via AKH, and its signal is transduced via the canonical AKHR.

Altogether, starvation resistance of flies that are deficient in AKH signaling results from increased lipid reserves, but reduced energy expenditure due to absence of starvation-induced hyperactivity might contribute to the resistance as well. While the starvation resistance of obese AKH-deficient flies might be advantageous to survive periods of paucity even under natural environmental conditions, the failure to induce foraging behavior would likely have fatal consequences. Thus, under natural conditions, AKH signaling is likely required to orchestrate adaptive responses to nutritional shortage, resulting in increased foraging.

Roles of AKH signaling in oxidative stress resistance

As documented by numerous studies on nondrosophilid insects, AKH plays a complex role in adaptation to oxidative stress. Ectopic applications of AKH to the linden bug P. apterus increased sensitivity to insecticide-triggered oxidative stress, such as to endosulfan, to malathion (Velki et al. 2011), and to permethrin (Kodrík et al. 2010). However, the AKH titer positively correlated with oxidative stress induced by bacterial toxin in Leptinotarsa decemlineata (Kodrík et al. 2007), and by paraquat in P. apterus (Večeřa et al. 2007) and L. decemlineata (Kodrík et al. 2007). Moreover, co-injection of AKH alleviated the effect of paraquat by enhancing the antioxidative capacity in the firebug (Večeřa et al. 2007), suggesting a protective role of AKH during oxidative stress response.

In Drosophila, foodborne paraquat exposure is the standard assay for testing oxidative stress resistance (Rzezniczak et al. 2011; Sun et al. 2013) and many fly studies successfully used this single agent (e.g., Alic et al. 2011; Barnes et al. 2014; Lin et al. 2014). Following this protocol, we observed increased paraquat resistance in all tested mutants deficient in AKH signaling. However, when analyzing the food intake during the experiment, we observed that AKH signaling mutants were hypophagic, when compared to controls fed on the same concentration of paraquat. Accordingly, reduced paraquat intake, not the oxidative stress resistance itself, might be causative for the extended survival time of the mutants exposed to foodborne paraquat. Indeed, AKH deficiency reduced paraquat resistance when differences in paraquat intake were bypassed by direct application of the drug on the nerve cord. Consistent with the protective role of AKH in the context of paraquat resistance in firebug (Večeřa et al. 2007), and with positive regulation of antioxidant enzymes by AKH as recently described in Drosophila (Bednářová et al. 2015), our experiment argues for the conserved role of AKH signaling in antioxidant defense. The exact mode of action whereby AKH facilitates the adaptive response to oxidative challenge awaits further detailed studies.

Our experiment on paraquat feeding vs. paraquat application also showed that the very same stressor could have differential effects on the very same genotypes, depending on the mode of its application. This outlines the importance of reevaluation of oxidative stress resistance tests dependent on spontaneous feeding, in particular when animals with unequal energy reserves are tested.

The functionally orphan adipokinetic hormone precursor-related peptide

APRP is the second peptide being processed from the Akh-encoded prohormone with currently unknown biological function (Figure 1, A and B). Our data confirmed the presence of APRP dimers in the CC cells of Drosophila; however, flies lacking APRP revealed no physiological defects in any of the tested processes. In particular, Akh^AP were fully viable, had normal developmental timing, body size, and oogenesis, which argues against the discussed ecdysiotropic role of APRP (De Loof et al. 2009) in Drosophila. Moreover, APRP plus AKH double mutants were indistinguishable from AKH single and AkhR mutants in all of the tested metabolic phenotypes.

The lack of systemic functions of Drosophila APRP is in line with the missing metabolic effects in response to injections of APRPs from the lubber grasshopper Romalea microptera (Hatle and Spring 1999). Together with the recent finding of high sequence variability at certain positions of the APRP peptide in cockroaches (Sturm and Predel 2015) this supports the view that APRP might mainly or exclusively have scaffold peptide function as proposed by the model of AKH processing in L. migratoria (Baggerman et al. 2002). However, a comprehensive analysis of potential systemic APRP functions awaits the availability of an APRP-specific single mutant.

Autoregulation of the Akh gene

Glucagon, the functional homolog of AKH in mammals, autoregulates its own production via proliferation control of the pancreatic alpha cells (Gelling et al. 2003) and via regulation of their secretory activity (Ma et al. 2005; Cabrera et al. 2008). Our data show that unlike its mammalian homolog, AKH does not affect the number of CC cells. Nevertheless, CC cell numbers appear to be subject to environmental or genetic variation, as our study detected more adult CC cells (16 ± 0.15 SEM) than was described previously using the same technical approach (13 ± 0.6 SEM) (Lee and Park 2004).

Regulation of the circulating AKH titer at the level of secretory activity of CC cells under energy-demanding conditions has been linked to the AMPK pathway (Braco et al. 2012). However, a role of AKH in AMPK activation, which would establish an autoregulatory loop, has not been addressed. Similarly, up-regulation of Akh on the transcriptional level in response to loss of insulin signaling has been described in Drosophila (Buch et al. 2008). But again, it is unclear whether this antagonistic regulation involves an autoregulatory mechanism. Here, we have shown that lack of AKH function increases Akh mRNA levels. Consistently, a GFP reporter under the indirect control of the Akh promoter showed the same regulatory response as did the endogenous Akh gene. These data provide the first evidence for a negative autoregulation of AKH, which is mediated by the Akh promoter. Future work will address the regulatory factor(s) acting on the Akh promoter, and the biological relevance of the predicted negative autoregulation of the Akh gene.

Conclusions

AKH signaling is considered to be a master regulator of energy mobilization in insects in various biological contexts, including development, reproduction, locomotion, and stress response. Whereas many of these functions were derived from AKH gain-of-function and correlation studies, here we addressed for the first time AKH in vivo functions using Drosophila AKH and AKH plus APRP double mutants. We showed that AKH signaling is dispensable for energy mobilization during the preadult stages of ontogenesis, but this pathway is of importance for storage lipid homeostasis in Drosophila adults. Ad libitum–fed AKH-deficient flies are obese and hypoglycemic, suggesting that lipid accumulation might result from increased cellular uptake of circulating sugars and enhanced lipogenesis in the fat body. Under food deprivation, AKH signaling contributes to lipid mobilization and induces starvation-induced hyperactivity, which likely reflects foraging behavior. We also provide evidence that AKH signaling confers oxidative stress resistance. Our study did not find any phenotype that could be attributed to the lack of APRP, arguing against its endocrine role in all of the tested processes. Comparison between the effects of Akh^A vs. AkhR¹ showed that the metabolic phenotypes of AKH are transduced via the canonical AKHR receptor. Surprisingly, several vital energy-demanding processes, such as locomotion, reproduction, and lipid and glycogen mobilization during preadult development are independent of AKH signaling. These results could be explained by evolutionary divergence of energy mobilization pathways in insects, reflecting the variability in insect life histories connected with differential preference for fueling energy stores as lipids, glycogens, or proteins. The rapid advance of genome engineering technologies, such as the CRISPR/Cas9 system used in this study, will hopefully result in AKH-specific mutants in a wider range of insect species, thus contributing to better understanding of the physiological functions of this ancient hormone system and their diversification during insect evolution.