RESEARCH ARTICLE

The role of glycogen in development and adult fitnessin DrosophilaTakayuki Yamada1, Okiko Habara1, Yuka Yoshii1,2, Ryota Matsushita1,2, Hitomi Kubo1, Yosui Nojima1,* andTakashi Nishimura1,2,‡

ABSTRACTThe polysaccharide glycogen is an evolutionarily conserved storageform of glucose. However, the physiological significance of glycogenmetabolism on homeostatic control throughout the animal life cycleremains incomplete. Here, we describe Drosophila mutants that havedefective glycogen metabolism. Null mutants of glycogen synthase(GlyS) and glycogen phosphorylase (GlyP) displayed growth defectsand larval lethality, indicating that glycogen plays a crucial role in larvaldevelopment. Unexpectedly, however, a certain population oflarvae developed into adults with normal morphology. Semi-lethalityin glycogen mutants during the larval period can be attributed tothe presence of circulating sugar trehalose. Homozygous glycogenmutants produced offspring, indicating that glycogen stored in oocytesis dispensable for embryogenesis. GlyS and GlyP mutants showeddistinct metabolic defects in the levels of circulating sugars andtriglycerides in a life stage-specific manner. In adults, glycogen as anenergy reserve is not crucial for physical fitness and lifespan undernourished conditions, but glycogen becomes important under energystress conditions. This study provides a fundamental understanding ofthe stage-specific requirements for glycogenmetabolism in the fruit fly.

KEY WORDS: Drosophila, Glycogen, Trehalose, Sugar metabolism,Physical fitness

INTRODUCTIONGlucose serves as a major energy source and also donates its carbonto most other synthesized molecules, such as the amino acids,nucleotides and fatty acids. Surplus glucose derived from dietarycarbohydrates is stored as branched polysaccharide glycogen ortriglycerides (TAGs) in the body for future energy needs (Saltiel andKahn, 2001; Chng et al., 2017; Mattila and Hietakangas, 2017).Proper regulation of anabolism and catabolism of stored energyreserves is crucial to sustain metabolic homeostasis throughout theanimal life cycle.The fruit fly Drosophila has two-programmed starvation periods,

embryo and pupa, during its life cycle. Glycogen is stored at thelate stage of oogenesis by metabolic remodeling in mitochondria(Sieber et al., 2016). Metabolic analysis has revealed that glycogen

stored in oocytes is consumed during embryogenesis (Tennessenet al., 2014; Matsuda et al., 2015). Consistently, the onset of aerobicglycolysis occurs in late-stage embryos through transcriptionalinduction (Tennessen et al., 2011). Likewise, glycogen stored infeeding larvae gradually decreases during metamorphosis (Gálikováet al., 2015; Matsuda et al., 2015). These observations suggest thatdeveloping embryos and pupae are using glycogen to produce ATPenergy as well as to generate biomolecules needed for cellularproliferation and differentiation to sustain embryonic developmentandmetamorphosis. However, the vital role of glycogen during thesedevelopmental periods has not been directly tested.

Glycogen metabolism is governed by evolutionarily conservedglycogen synthase and phosphorylase by the concerted action ofbranching and de-branching enzymes (Roach et al., 2012).In mammals, glycogen is primarily stored in the cells of the liverand muscles. Liver glycogen plays an important role in the glucosecycle tomaintain circulating sugar levels, whereasmuscle glycogen isdirectly utilized through glycolysis to maintain muscle activities(Roach et al., 2012; Petersen et al., 2017). In Drosophila larvae, wehave shown previously that the tissue-specific regulation of glycogenmetabolism in the fat body, an organ equivalent to the mammalianliver, plays a crucial role in the maintenance of circulating sugarsunder fasting conditions (Yamada et al., 2018). In adults, glycogen isstored in flight muscles and consumed during flight (Wigglesworth,1949). Interestingly, emerging evidence suggest that geneticmanipulation of glycogen metabolism extends lifespan in severalspecies, includingCaenorhabditis elegans andDrosophila (Bai et al.,2013; Gusarov et al., 2017; Post et al., 2018). On the other hand, theprogressive accumulation of glycogen in neurons leads to neuronalcell death, locomotion deficits and reduced lifespan, therebycontributing to physiological aging (Duran et al., 2012). Althoughthe requirements for glycogen metabolism likely change overdevelopmental time in a tissue-specific manner, it remains unclearhow animals respond to loss of a major energy reserve and maintainbody homeostasis through compensatory metabolic mechanisms.

Here, we generated and characterized Drosophila defective forglycogen metabolism. We observed that more than half of themutants are lethal in larvae. However, to our surprise, a certainpopulation of larvae undergoes pupariation and enters adulthood.Importantly, we did not observe an essential contribution ofglycogen during embryogenesis and metamorphosis. Glycogensynthase (GlyS) mutant adults showed reduced physical fitness, butthis could conceivably be an indirect consequence of metabolicdefects, including a reduction in trehalose. Glycogen phosphorylase(GlyP) mutants displayed normal flight performance, climbingability and adult lifespan, comparable to those of control flies,suggesting that glycogen as a fuel source is largely dispensable foradult fitness. In contrast, the importance of glycogen metabolismbecomes apparent under fasting conditions. This study providesthe first direct evidence that glycogen metabolism plays anReceived 24 January 2019; Accepted 21 March 2019

1Laboratory for Growth Control Signaling, RIKEN Center for Biosystems DynamicsResearch (BDR), 2-2-3 Minatojima-Minamimachi, Chuo-ku, Kobe, Hyogo 650-0047, Japan. 2Graduate School of Biological Science, Nara Institute of Science andTechnology, 8916-5 Takayama, Ikoma, Nara 630-0101, Japan.*Present address: Laboratory of Bioinformatics, National Institutes of BiomedicalInnovation, Health and Nutrition, 7-6-8, Saito-asagi, Ibaraki, Osaka 567-0085,Japan.

‡Author for correspondence (t-nishimura@cdb.riken.jp)

T.N., 0000-0002-6773-0302

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© 2019. Published by The Company of Biologists Ltd | Development (2019) 146, dev176149. doi:10.1242/dev.176149

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important role in larval development and adult fitness but not inembryogenesis and metamorphosis in Drosophila.

RESULTSGeneration and validation of GlyS and GlyP null allelesIn Drosophila, a set of single orthologous genes are involvedin glycogen metabolism (Fig. 1A); namely, GlyS, 1,4-α-glucan

branching enzyme (AGBE), GlyP and the de-branching enzymeamylo-α-1,6-glucosidase, 4-α-glucanotransferase (AGL; FlyBaseannotation symbol CG9485). To examine the requirements ofglycogen metabolism over the life cycle, we generated null mutantsof GlyS and GlyP (Fig. 1B,C). GlyS8 mutants, carrying a smalldeletion at the N-terminal coding region, were created using theCRISPR/Cas9 system. GlyP3-13 mutants, in which mutants lack the

Fig. 1. See next page for legend.

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entire first exon, were made by imprecise excision of a P-element.Unexpectedly, we found that homozygous mutants of GlyS8

and GlyP3-13 were viable at a significantly lower ratio thanheterozygotes (see below). Homozygous adults of GlyS8 andGlyP3-13 displayed normal morphology (Fig. 1D).To confirm whether newly generatedGlyS8 andGlyP3-13mutants

were functional null alleles, we examined glycogen levels duringthe wandering stage (late third-instar larvae) and in 1-week-oldadult males and females. As expected, the amount of glycogensignificantly decreased in GlyS mutants, but increased in GlyPmutants (Fig. 1E). GlyS mRNA levels were drastically decreased inadults (Fig. 1F), most likely due to nonsense-mediated mRNAdecay that eliminates mRNAs containing a premature terminationcodon (Karousis et al., 2016).GlyPmRNAwas almost undetectableowing to the lack of a transcriptional start site.To demonstrate directly that GlyS8 and GlyP3-13 mutants

completely abolish glycogenesis and glycogenolysis, we developedenzyme activity assays using isotope-labeled glucose, followed bymass spectrometric analyses. Glycogenesis by GlyS was assessed bymeasuring the incorporation rate of U-13C-glucose into glycogenfrom uridine diphosphate (UDP)-U-13C-glucose. As expected,no incorporation was observed in tissue homogenates from GlySmutant larvae even though glycogen was exogenously added to thereaction mixture (Fig. 1G). Similarly, glycogenolysis by GlyP wasassessed by measuring the release rate of U-13C-glucose-phosphatefrom 13C-labeled glycogen that was purified from adult flies feda U-13C-glucose-containing diet (Fig. S1A,B). No release of13C-glucose-phosphate was detected in GlyP mutant homogenates.Together, these results unambiguously indicate thatGlyS8 andGlyP3-

13 mutants are functional null alleles that completely eliminate thecorresponding enzymatic steps in glycogen metabolism.We previously reported that periodic acid-Schiff (PAS) staining

reliably visualizes stored glycogen and also detects mobilizationupon brief starvation in the fat body (Yamada et al., 2018).Consistent with our previous research (Yamada et al., 2018), GlySmutants showed strong reductions in the PAS signal in tissues suchas the CNS, the fat body and body wall muscles. This reduction wasfully rescued by the ubiquitous expression of C-terminal Flag-tagged GlyS by Tub-Gal4 (Fig. 1H). GlyP mutants failed tomobilize fat body glycogen upon starvation, which was completelyrescued by the expression of C-terminal Flag-tagged GlyP. The

overexpression of GlyP had no effect on PAS signals under fedconditions, as described below. Moreover, these transgenes restorednormal glycogen levels in whole animals (Fig. 1I). Thus, the UASconstructs we generated produced a functional protein.

GlyS and GlyP mutants display semi-lethality during thelarval periodBecause glycogen mutants showed a significant level of mortalityprior to the adult emergence, we next examined the lethal stage andfound that more than half of GlyS8 and GlyP3-13 mutants exhibitedlarval lethality (Fig. 2A). Consistent with results from homozygotesin each mutant allele, the transheterozygotes over a deficiency lineshowed similar lethality during the larval period. Importantly, themutant lethality was fully rescued by ubiquitous expression of GlySor GlyP (Fig. 2B), indicating that the observed larval lethality isspecific to the loss of function inGlyS and GlyP genes. By contrast,after pupariation of surviving larvae, GlyS and GlyP mutantslethality was not observed during metamorphosis (Fig. 2C). Wefurther examined the phenotypes of available transposon insertionmutants (Fig. 1B). Transheterozygotes of GlySMI01490 andGlyPMI00957 over a deficiency line exhibited semi-lethality inlarvae, but not in pupae, resulting in the emergence of escapingadults (Fig. 2A,C). Based on the degrees of larval lethality, theirinsertion sites and the mRNA level (Fig. 1F), we can conclude thatGlyPMI00957 is a null allele that is comparable to GlyP3-13, whereasGlySMI01490 is a strong hypomorphic allele, as previously reported(Yamada et al., 2018). The backcrossed GlyPk07918 (renamedGlyPlacW) is a weak hypomorphic allele. Taken together, theseresults indicate that GlyS and GlyP are required for normaldevelopment in larvae, but not in pupae.

To identify the timing of lethality, we counted the number ofsurviving larvae and found that the survival rate of both GlyS8 andGlyP3-13 mutants decreased progressively during the third instar(Fig. 2D). Molting was not related to the lethality of these mutants asevidenced by the completion of molting from the late second to thirdinstar [100% of theGlyS8mutants (n=63) and 100% of theGlyP3-13

mutants (n=71) from more than five independent experiments].Body size of these mutants was normal at 24 h after larval hatching(ALH) (i.e. late first instar) and 48 h ALH (i.e. late second instar)(Fig. 2E). However, at 72 h ALH (i.e. mid-third instar), both GlySand GlyP mutants were ∼20-34% smaller on average than agenetically matched control (w−), indicating that glycogenmetabolism is required for normal body growth during the thirdinstar. However, GlySmutants showed normal body size at the timeof pupariation in both males and females. In contrast, GlyP mutantsstill showed smaller pupae, but the size differences averaged only∼5-7% (Fig. 2F). Developmental timing until pupariation was notaffected in GlyS and GlyP mutants that survived (Fig. 2G); thus,developmental delay cannot account for the restoration of body sizein these mutants. It is of note that both GlyS and GlyP mutantsconsumed similar amounts of food compared with control at theearly third instar stage (Fig. 2H), suggesting that feeding behavior isnot affected in these mutants. Adult body weight in GlyS mutantswas almost indistinguishable from that of control flies (Fig. 2I). Incontrast, GlyP mutant males and females were smaller or leaner by∼7-10% on average. The GlyP mutant-specific decreases in bodyweight might be partly attributed to the reduction in stored lipids, asdescribed below.

We hypothesized that the observed semi-lethality of glycogenmutants in the larval period might be explained by the presence ofcirculating sugar trehalose (Fig. 1A), which also plays an importantrole in maintaining glucose homeostasis (Elbein et al., 2003;

Fig. 1. Generation and validation of GlyS and GlyP null alleles.(A) Overview of glucose and storage sugar metabolism. Genes that functionin glycogen and trehalose metabolism are shown in red. (B) Schematic ofthe GlyS and GlyP loci and molecular nature of the mutants. Protein-codingregions and untranslated regions are represented by black boxes and whiteboxes, respectively. The P-element insertion sites are marked with invertedtriangles. (C) Sequences of the sgRNA target site and the GlyS8 deletionmutant. The 20-bp target sequence is indicated in blue, and the cleavage siteof Cas9 in indicated by the inverted triangle. (D) GlyS and GlyP null mutantswere viable without obvious morphological defects. (E) Total amounts ofglycogen were analyzed in the indicated genotypes and are plotted relative tothat of control. n=9-12 (late third instar: wandering larvae) or 9 (adult) batchesfrom 3-4 independent experiments. (F) Expression levels of GlyS and GlyPwere analyzed by qRT-PCR. n=4 batches. (G) GlyS and GlyP enzymeactivities were assessed in tissue homogenates at early third instar. Theaverage values from two independent experiments are shown. A.U., arbitraryunit; Glucose-P, glucose phosphate. (H) Expression of GlyS and GlyP in eachmutant background restored glycogenesis and glycogenolysis, respectively,as visualized by PAS staining. Tub-Gal4 was used for ubiquitous expression.Scale bars: 100 μm. (I) Expression of GlyS and GlyP in each mutantbackground fully restored glycogen levels in late third instar larvae. n=5-9batches from two independent experiments. One-way ANOVA with Dunnett’spost-hoc test (E,F,I); ***P<0.001.

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Chng et al., 2017;Mattila andHietakangas, 2017). In contrast toGlySandGlyPmutants, null alleles of the trehalose synthesis enzymeTps1show complete lethality during metamorphosis, but not during thelarval period when dietary sugar is available (Matsuda et al., 2015;Yasugi et al., 2017). To test phenotypic interaction between glycogenand trehalose metabolism, we analyzed the lethality in larvae lackingboth glycogen and trehalose metabolism.GlyP,Tps1 double mutantsshowed severe lethality during the second and third instar, whichresulted in almost complete lethality before pupariation (Fig. 2D).Moreover, GlyP,Tps1 double mutants displayed growth defects at

48 h ALH, but each single mutant had no effect (Fig. 2E,J). Thesegenetic interactions suggest that disrupting trehalose metabolism hasno effect on larval survival, disrupting glycogenmetabolism inducesa semi-lethal phenotype, whereas disrupting bothmetabolisms likelyresults in a dramatic homeostatic defect.

GlySandGlyPmutantsexhibit distinctmetabolic defects inalife stage-dependent mannerGlycogen metabolism is thought to play an important role inmaintaining postprandial glucose homeostasis. Thus, we next

Fig. 2. GlyS and GlyP mutants display semi-lethality during the larval period. (A) GlyS and GlyP mutants exhibited lethality during the larval period.(B) Expression of GlyS orGlyP rescued larval lethality in the respective GlyS and GlyPmutants. Percentages of surviving larvae were determined by the ratio toheterozygotes in each vial. (C) Glycogen mutants did not exhibit lethality during the pupal period. (D) Glycogen mutants died at increasing rates during the thirdinstar indicated by the gray region. (E) Larval volume in glycogen mutants was analyzed at the indicated time points. (F) Pupal volume in males and females.(G) Glycogen mutants did not exhibit developmental delay in the timing of puparium formation. (H) Glycogen mutants exhibited normal feeding behavior. Foodintake levels were evaluated by the rate of blue food ingestion by early third instar larvae. (I) Adult body weight was analyzed in glycogen mutant males andfemales. (J)GlyP and Tps1 double mutants showed growth defects at 48 h ALH. The numbers of vials (A-D,G), animals (E,F,J) and batches (H,I) from at least twoindependent experiments are indicated. One-way ANOVA (A-C,E-J) or two-way ANOVA (D) with Dunnett’s post-hoc test; *P<0.05, ***P<0.001; n.s., notsignificant.

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analyzed the amount of glucose, trehalose and cellular lipid storageTAGs in glycogen mutants.GlyS and GlyPmutants showed distinctmetabolic defects in a stage-dependent manner (Fig. 3A). In thewandering larvae, trehalose was significantly decreased in GlyPmutants but not in GlyS mutants. By contrast, no defects wereobserved in glucose and TAGs. Unexpectedly, at the adult stage,GlySmutants showed significant reductions in steady-state levels oftrehalose and glucose, whereas GlyP mutants showed mild, if any,reductions. By contrast, male and female GlyP mutants showed adrastic reduction (∼50%) in TAGs. Collectively, defects inglycogen metabolism in whole animals do not increase thesemetabolites; rather, the defects in glycogen metabolism result in adrop in either circulating sugar or lipid storage.To further clarify potential metabolic alterations during the larval

stage, a widely targeted metabolomics analysis was conducted usingwhole animals in the wandering larvae by liquid chromatographycoupled with tandem mass spectrometry (LC-MS/MS) (Table S1).The overall results of changes in 158 water-soluble metabolites were

visualized using a principal component (PC) analysis. The score plot,in which each point represents an individual sample, revealed a clearseparation between three genotypes (control w−, GlyS mutants andGlyPmutants) (Fig. 3B).Venn diagramcomparisons further revealedspecific and common metabolic defects in GlyS and GlyP mutants(Fig. 3C). Interestingly, three of five common defects between GlySand GlyP mutants were in the amino acids threonine, histidineand glutamate. A volcano plot showed significant reduction inthreonine and an increase in histidine and glutamate (Fig. 3D,E).Because these three amino acids are glucogenic and used for energyproduction through pyruvate or 2-oxoglutarate, amino acidmetabolism is likely to be altered in mutant larvae. Thiaminepyrophosphate (TPP) is an essential co-factor for several enzymes incarbohydrate metabolism, including pyruvate dehydrogenase and2-oxoglutarate dehydrogenase, which is a rate-limiting step in thetricarboxylic acid (TCA) cycle. Reduction in TPP in both GlyS andGlyP mutants further implies that defects in glycogen metabolismpotentially cause abnormalities in primary metabolic pathways.

Fig. 3. GlyS and GlyP mutants show distinct metabolic defects in a life stage-dependent manner. (A) Relative amounts of trehalose, glucose and TAGin glycogen mutants. n=9-12 (late third instar) or 9 (adult) batches from 3-4 independent experiments. (B) PC analysis of water-soluble metabolites in thecontrol and glycogenmutants (late third instar). Ellipses of clusters show the 95% confidence regions for each sample group. (C) Numbers of significantly differentmetabolites (q<0.01 with respect to control) betweenGlyS andGlyPmutants. (D) Volcano plots showing q-values versus fold change (FC) from themetabolomicsdata. (E) Relative amounts of metabolites that significantly changed in glycogen mutants. Control (A-E): a genetically matched strain, w1118. n=6 batches (B-E);one-way ANOVA with Dunnett’s post-hoc test (A,E); *P<0.05, **P<0.01, ***P<0.001; n.s., not significant.

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In addition to changes in amino acids, both GlyS and GlyPmutants decreased acetyl-CoA and acetoacetyl-CoA (Fig. 3E).These decreases coincided with increases in acetylcarnitine andacylcarnitines. Although the total amount of TAGs was not changed(Fig. 3A), these results suggest active lipolysis or impaired lipidmetabolism at the late larval stage. Together, these results andadditional support by enrichment analysis of metabolomics data(Table S1) indicate that defects in glycogen metabolism impactamino acid and lipid metabolism.

Larval lethality in glycogen mutants is partly rescued byenriched foodsTo elucidate the cause of larval lethality, we then investigatedwhetherglycogen mutants exhibit sensitivity to malnutrition. We previouslyreported that Tps1mutants are sensitive to sugar deprivation and poordietary conditions (Matsuda et al., 2015; Yasugi et al., 2017).However, larval lethality inGlyS and GlyPmutants was not changedunder a yeast-only (Y-only) diet that largely lacks carbohydratescompared with that of mutants on a normal diet (ND) (Fig. 4A).Similarly, glycogen mutants were able to reach adulthood on a poordiet (1/5Y) without apparent defects in body growth (Fig. 4A,B).Thus, unlike Tps1 mutants, undernutrition does not appear to be themajor cause of larval lethality in GlyS and GlyP mutants.We next tested whether enriched foods could rescue the mutant

lethality. We found that larval lethality in GlyS, but not GlyP,mutants was partially rescued under high-yeast (2Y) and high-glucose (2G) diet conditions (Fig. 4A). Although GlyS mutants

grew normally under these dietary conditions, mutant larvae thatsurvived showed strong lethality during metamorphosis (Fig. 4A,B). These results suggest that enriched foods are able to overcomelarval lethality in GlyS mutants, but the diet induces deleteriouseffect(s) during metamorphosis.

To determine which organ is crucial for the observed larvallethality in glycogen mutants, we conducted tissue-specificknockdown experiments. Similar to the mutant phenotype,ubiquitous knockdown of GlyS and GlyP by Tub-Gal4 led tosemi-lethality in larvae, with a certain population becoming pupaeand adults (Fig. 4C). Glycogen is mainly stored in the muscles, fatbody and CNS in larvae (Yamada et al., 2018). Muscle-specificknockdown by Mef2-Gal4 induced strong lethality that variedconsiderably between vials and experiments (Fig. 4C). By contrast,pan-neuronal knockdown by Elav-Gal4 produced a slight reductionin the survival rate of larvae, and pan-glial knockdown by Repo-Gal4 and fat body-specific knockdown by Cg-Gal4 had modestimpacts, if any, on viability (Fig. 4C). The combination of muscle-and neuron-specific knockdown fully recapitulated the lethalityobserved when the RNAi was induced ubiquitously (Fig. 4C). Thus,larval lethality in GlyS and GlyP mutants is caused by synergisticdefects in multiple organs, including muscles and neurons.

Differences in glycogen metabolism and trehalosemetabolismWe next examined the consequences of GlyS and GlyPoverexpression. The GlyS and GlyP proteins overexpressed by

Fig. 4. Larval lethality in glycogen mutants is partly rescued by enriched foods. (A) The larval survival rate and eclosion rate of glycogen mutantsunder various dietary conditions. ND, normal diet; Y-only, yeast-only low-carbohydrate diet; 1/5Y, low-protein diet; 2Y, 2× yeast diet; 2G, 2× glucose diet. (B) Pupalvolume of glycogen mutants grown under various dietary conditions. (C) The larval survival rate was analyzed in tissue-specific knockdown under ND. Thetotal numbers of vials (A,C) and pupae of mixed gender (B) from at least two independent experiments are indicated in each graph. One-way ANOVA withDunnett’s post-hoc test with respect to ND in each genotype (A) or control in each diet (B) or Tukey’s post-hoc test (C); *P<0.05, **P<0.01, ***P<0.001;n.s., not significant.

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Tub-Gal4 were distributed uniformly in the cytoplasm in the larvalfat body (Fig. 5A). However, despite high-level overexpression ofGlyP, there was no visible impact on the level of stored glycogen inseveral tissues (Fig. 5B, Fig. S2A). These results were confirmed bybiochemical quantification in whole animals at the early third instarand in the wandering larvae (Fig. 5C, Fig. S2B). Furthermore,overexpression of GlyS slightly increased the amount of glycogen.Consistent with these observations, flies overexpressing either GlySor GlyP by Tub-Gal4 were viable.These glycogen results were in striking contrast to those

regarding trehalose metabolism. The overexpression of Tps1drastically increased trehalose (Fig. 5D) (Matsuda et al., 2015).Likewise, overexpression of the trehalose-hydrolysis enzyme Treh,especially secreted Treh (sTreh), strongly depleted trehalose(Fig. 5C,D). Chronic overexpression of cytoplasmic Treh (cTreh)had a marginal effect. However, the transient induction of cTrehusing heat-shock Gal4 (hsGal4) transiently decreased trehalose(Fig. 5D). Thus, the overexpression of Tps1 and Treh was sufficientto promote trehalose synthesis and breakdown, respectively. Theseresults suggest that the regulatory mechanisms for glycogenmetabolism and trehalose metabolism are different.

Stored glycogen in oocytes is required for trehalosesynthesis but largely dispensable for embryogenesisOur genetic analyses demonstrated that homozygous glycogenmutants are viable. Consistent with a previous report (Sieber et al.,

2016), glycogen accumulated in matured oocytes during oogenesis(Fig. S3A), suggesting that stored glycogen is a crucial energyreserve in embryogenesis. However, contrary to expectations, wefound that embryos laid by either GlyS and GlyP homozygousfemales exhibited only a slight reduction in hatching rates (Fig. 6A).Moreover, GlyS and GlyP homozygous mutants could bemaintained for more than a year. We noticed that the timing ofhatching was significantly delayed by 2 h in both GlyS and GlyPmutants (Fig. 6B), suggesting that glycogen metabolism plays a rolein timely progression of embryogenesis.

One possibility is that GlyS and GlyP mutants compensate forenergy production by oxidizing other sources, such as stored lipidsand amino acids. To understand the global change in metabolismcaused by defective glycogen metabolism during embryogenesis,we conducted a comparative metabolomics analysis of water-soluble metabolites and TAGs (Table S2, Fig. S3B). PC analysisusing 161 water-soluble metabolites revealed a separation betweenearly embryo [0-2 h after egg laying (AEL)] and late embryo(22-24 h AEL) in the direction of PC1, whereas PC2 and PC3separated three genotypes (i.e. control w−, GlyS and GlyP mutants)(Fig. 6C). Venn diagram comparisons using the metabolomics dataincluding water-soluble metabolites and TAGs revealed that GlySand GlyP mutants showed common metabolic changes, especiallyat the late embryo stage (Fig. 6D, Fig. S3C), suggesting thatglycogen mutants induce metabolic stress by the progressionof embryogenesis.

Fig. 5. Differences in glycogen and trehalose metabolism. (A) Localization of overexpressed GlyS, GlyP and Treh in the fat body. The punctate localizationof sTreh-Flag suggests secretory vesicles. Scale bar: 50 μm. (B) The effects of GlyS and GlyP overexpression on the levels of stored glycogen as visualizedby PAS staining at the wandering stage. Scale bar: 100 μm. (C) The effects of GlyS, GlyP and Treh overexpression on glycogen and trehalose levels at thewandering stage. The total numbers of batches in each genotype are indicated. (D) The effects of transient induction of genes involved in glycogen andtrehalose metabolism using a heat shock-Gal4 (hs-Gal4). Early third instar larvae (0 h) on a normal diet were heat-treated for 1 h and analyzed after 6 h or 18 h.Heat shock itself induced the mobilization of glycogen. Values shown are mean±s.e.m. n=3 batches. One-way ANOVAwith Dunnett’s post-hoc test (C), two-wayANOVA with Dunnett’s post-hoc test with respect to values in control (D); **P<0.01, ***P<0.001.

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We confirmed that embryos laid by GlyS mutant females did notretain glycogen (Fig. 6E). Embryos laid by GlyP mutant femaleshad more glycogen than control embryos. Interestingly, a significantreduction of stored glycogen was observed in GlyP mutants duringembryogenesis, suggesting that GlyP-independent degradation ofglycogen occurs. We further found that consumption of TAGsduring embryogenesis did not vary between control and glycogenmutants, although significant changes in the intermediates of fattyacid oxidation, such as even-chain acylcarnitines andacetylcarnitine, were detected (Fig. 6E). The increase of uric acid,a nitrogen-containing waste product of amino acid and purinecatabolism, occurred normally in GlyS and GlyP mutants.Furthermore, reduction of ATP was not observed in glycogenmutants (Fig. 6E). These results suggest that loss of glycogenmetabolism does not force a detectable switch to fatty acid or aminoacid oxidation.Interestingly, the increase of trehalose and glucose at late

embryo was strongly impaired in GlyS and GlyP mutants

(Fig. 6E), indicating that stored glycogen is utilized as a carbonsource to produce trehalose and glucose during embryogenesis.Taken together, these results support the conclusion that glycogenmetabolism, per se, is largely dispensable for embryogenesis;however, glycogen mutants do exhibit metabolic alterations.

Glycogen metabolism is crucial to maintain adult fitnessunder starvation conditionsTo further understand the physiological significance of glycogen inadult fitness, we next analyzed physical ability. Adult flies attemptto climb to the top of a vial, in opposition to gravity, as an innatebehavior (Rhodenizer et al., 2008). By recording their locomotion,we measured the average speed at which individual flies climbed.We found that GlyS mutant males had significantly decreasedclimbing ability, which was fully rescued by ubiquitous expressionof GlyS (Fig. 7A). By contrast, GlyP mutants did not showremarkable deficits in climbing speed. Aging slightly reducedphysical activity in GlyP mutants (Fig. 7B). We next conducted a

Fig. 6. Stored glycogen in oocyte is required for trehalose synthesis but is largely dispensable for embryogenesis. (A) The hatching rate of embryoslaid by homozygous GlyS and GlyP mutant females. The numbers of batches and the total number of embryos (in parentheses) are indicated. (B) Glycogenmutants exhibited 2-h delays in timing of larval hatching. (C) PC analysis of water-solublemetabolites at early (0-2 h) and late (22-24 h) embryonic stages. Ellipsesof clusters show the 95% confidence regions for each sample group. (D) Numbers of significantly different metabolites (q<0.001 with respect to control) betweenGlyS and GlyP mutants at each stage. (E) Relative changes in the amounts of metabolites during embryogenesis. Metabolite levels were directly analyzedby LC-MS/MS or after enzymatic treatment (for glycogen). TAG levels indicate the sum of detected TAG lipid classes. Control (A-E): a genetically matched strain,w1118. n=8 batches (C-E); one-way ANOVA with Dunnett’s post-hoc test (A) or Tukey’s post-hoc test (E), Kolmogorov–Smirnov test (B); **P<0.01, ***P<0.001.

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flight performance test in 5-day-old males. Approximately 30% ofGlyS mutants showed compromised flight ability, whereas GlyPmutants were largely normal (Fig. 7C). We further examinedadult lifespan and found that GlyS mutant males had a slightlyshortened lifespan (6-day difference in the median survival),whereas GlyP mutants had a normal lifespan (Fig. 7D). Takentogether, these results suggest that GlyS is required for normalphysical performance and adult fitness, whereas GlyP is largelydispensable under normal dietary conditions.These results raise the question of why onlyGlySmutants showed

an apparent deficit in physical activity under fed conditions. GlySmutants exhibited a specific decrease in circulating trehalose andglucose (Fig. 3A), so perhaps a reduction in circulating sugars

explains the lower physical activity. To test this hypothesis, weexamined a hypomorphic allele in Tps1 (Matsuda et al., 2015). Asexpected, Tps1MIC homozygous adults showed a significantdecrease in climbing ability (Fig. 7E). These results support thepossibility that the lower physical fitness in GlyS mutants is anindirect consequence of metabolic defects caused by impairedglycogenesis rather than the lack of glycogen metabolism as afuel source.

If glycogen metabolism plays a role in maintaining energyhomeostasis in times of metabolic need,GlyPmutants may decreasetheir physical activity after starvation. Indeed, we found that bothGlyS and GlyP mutants had a lower survival rate after starvation(Fig. 7F). Control flies maintained their climbing activity 1 day

Fig. 7. Glycogen metabolism is crucial for adult fitness under starvation conditions. (A) GlyS, but not GlyP, mutant males showed impaired physicalfitness. Adult locomotion ability was assessed by a climbing assay. The average speeds of 5-day-old flies are shown. (B) Aging slightly enhanced the physicaldefect in GlyP mutants. The percentage of flies that climbed up to 10 cm was plotted. (C) A certain population of GlyS, but not GlyP, mutant males displayedcompromised flight ability. The x-axis represents the vertical positions where the flies landed in the cylinder, and the y-axis represents the percentage of flies.The gray region indicates the bottom of the cylinder. (D) Adult lifespan in GlyS and GlyP mutant males. (E) Tps1MIC mutant males showed impaired physicalactivity. The average climbing speeds of 5-day-old flies are shown. (F) Both GlyS and GlyPmutants showed reduced survival rates after starvation. Ten-day-oldflies were used for the starvation assay. (G) One-day starvation decreased the climbing activity in GlyP mutant males. The average speeds of 5-day-old fliesare shown. Control (A-G): a genetically matched strain, w1118. The total numbers of flies from at least two independent experiments are indicated. One-wayANOVAwith Dunnett’s post-hoc test (A,E) or Tukey’s post-hoc test (G), Kruskal–Wallis test with Dunn’s correction (B,C), log-rank test (D,F); *P<0.05, ***P<0.001;n.s., not significant.

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after starvation compared with those under fed conditions (Fig. 7G,Fig. S4). However, both GlyS and GlyP mutants had decreasedclimbing speed when climbing performance was tested 1 day afterstarvation. Taken together, these results demonstrate that glycogenas a carbon source contributes to the maintenance of physicalperformance under fluctuations of dietary conditions.

DISCUSSIONIn Drosophila, the importance of glycogen in homeostatic control,such as starvation tolerance, hypoxia tolerance, muscle function,protection from glucose toxicity, and aging, has been analyzed intissue-specific overexpression and knockdown experiments (Duranet al., 2012; Paik et al., 2012; Zirin et al., 2013; Saez et al., 2014;Sinadinos et al., 2014; Garrido et al., 2015; Yamada et al., 2018).However, genetic null mutants of glycogen metabolism enzymeshave not been reported.In this study, we demonstrate that glycogen metabolism plays an

important role in larval survival in collaboration with the circulatingsugar trehalose. We also determine that muscle is an importantorgan for the observed lethality in GlyS and GlyP mutants.Interestingly, 90% of mice lacking muscle glycogen synthase(GYS1) die shortly after birth owing to abnormal cardiacdevelopment, but the surviving null mice grow up withoutobvious abnormalities, other than being smaller than wild-typemice (Pederson et al., 2004, 2005). Muscles are primarily defined asthe organs for movement. However, muscle-specific knockdown ofGlyS has no effect on larval locomotion under fed conditions (Zirinet al., 2013). Consistent with this, we did not detect a reduction infood intake, which depends on muscles for mouth hook movementand gut peristalsis (Min et al., 2017). Thus, the larval lethal effectappears not to rely on the movement function of muscles. Becauseof its mass and high metabolic rate during exercise, muscle has aprofound influence on body metabolism. Recently, muscle has beenshown to orchestrate many aspects of animal physiology, includingbody growth and lifespan, by regulating organismal energyhomeostasis and governing systemic signaling networks(Droujinine and Perrimon, 2016; Rai and Demontis, 2016). Forexample, muscle-derived growth factors and cytokines, known asmyokines, mediate the endocrine functions of muscles on othertissues. Thus, glycogenmetabolism inmuscles is important for larvaldevelopment, likely through a systemic effect on body homeostasis.Our results indicate that disrupting two enzymes acting in

opposite directions provokes at least in part similar phenotypictraits. Potentially, common phenotypic traits in both GlyS and GlyPmutants reveal problems in carbohydrate homeostasis, whereasspecific phenotypes in either GlyS or GlyP mutants reveal theconsequence of lack or excess of glycogen granules. Interestingly,GlyS and GlyP mutants showed distinct metabolic defects in a lifestage-specific manner. One possibility is that the lack or over-accumulation of glycogen influences intracellular signaling andtranscription, resulting in changes in sugar and lipid metabolism.This idea is supported by the fact that glycogen inhibits the kinaseactivity of AMPK by binding to the glycogen-binding domain of theβ-subunit (McBride et al., 2009). Moreover, glycogen granules actas a signaling scaffold via physical interaction with a variety ofproteins (Graham, 2009; Philp et al., 2012; Stapleton et al., 2013).Thus, the flux of either glycolysis or fat oxidation may be promotedthrough a metabolic shift to compensate for the energy deficit in amutant-specific manner. Considering that metabolism is intricatelyconnected to diverse pathways (Kamleh et al., 2008; St Clair et al.,2017), impaired glycogenesis and glycogenolysis may also lead todistinct metabolic outcomes. Although the detailed mechanisms

remain largely unknown, accumulating evidence has revealed aprofound difference in carbohydrate metabolism between larvaeand adults (Mattila and Hietakangas, 2017). Further analysis will berequired to clarify the mutant- and life stage-specific metabolicchanges.

We found that the majority of glycogen mutant embryossuccessfully hatch, but with a 2-h delay. In contrast to glycogen,the amount of trehalose increases during embryogenesis (An et al.,2014; Tennessen et al., 2014; Matsuda et al., 2015). Our resultsindicate that the carbon source of trehalose synthesis duringembryogenesis is highly dependent on stored glycogen in oocytes.Nevertheless, trehalose synthesized in the embryo is dispensable forembryogenesis. This result is consistent with our previous reportshowing that maternal and zygotic mutation of Tps1 does not resultin embryonic lethality (Matsuda et al., 2015). Because Tps1 andTreh mutant larvae are extremely sensitive to dietary deprivation(Matsuda et al., 2015; Yoshida et al., 2016), the physiologicalsignificance of trehalose synthesis during embryogenesis is the mostlikely explanation for survival immediately after hatching. Theseresults are consistent with previous reports showing that globalexpression of genes involved in aerobic glycolysis occurs at the endof the embryonic stage for metabolic adaptation in larvae(Tennessen et al., 2011, 2014). In this regard, a major purpose ofglycogen deposition in matured oocytes is to distribute a source ofglucose for larval survival but not a fuel source for embryogenesis.The amount of trehalose should be spatiotemporally regulated inembryogenesis because ectopic trehalose synthesis potentiallyinduces uncontrolled osmotic swelling that results in lethality(Matsuda et al., 2015; Yoshida et al., 2016) and that precludes directdeposition of trehalose in oocytes. These observations further implythat stored TAGs are likely an indispensable fuel source to completeembryogenesis. Interestingly, GlyP mutants deposit normal levelsof TAGs in embryos, despite a significant reduction in the wholebody levels of TAGs in adults, suggesting that adult females are ableto control the correct deposition of stored lipids in oocytes,presumably through amechanism involved in local steroid signaling(Sieber and Spradling, 2015).

Our results also indicate that stored glycogen is partly consumedduring embryogenesis in GlyP mutants. It appears that GlyP-independent glycogenolysis occur at this stage, most likely vialysosome/autophagy-dependent degradation of glycogen (Roachet al., 2012; Zirin et al., 2013). The partial mobilization of storedglycogen inGlyPmutants is strongly supported by the fact that GlyPmutants synthesize trehalose to some degree during embryogenesis.In addition, relatively weak embryonic lethality in GlyP mutantscompared with GlyS mutants could also be attributed to the partialglycogenolysis.

We found that the ectopic induction of GlyS has a modest impacton glycogen levels, whereas that ofGlyP is not sufficient to promoteglycogen mobilization in larvae. Because glycogen metabolism istightly regulated by allosteric and post-translational modifications(Bouskila et al., 2010; Roach et al., 2012), it is reasonable to assumethat increased levels of GlyS and GlyP fail to facilitate eitherglycogenesis or glycogenolysis. Levels of glycogenin, a coreprotein for glycogen synthesis, may also restrict the amount ofglycogen (Lomako et al., 2004; Roach et al., 2012). It should benoted, however, that GlyP overexpression reduces glycogen inadults (Post et al., 2018) and the expression of human GYS1increases the amount of glycogen in Drosophila adult neurons(Sinadinos et al., 2014). Furthermore, GlyS and GlyP expression iselevated in whole larvae on a high-sugar diet (Musselman et al.,2011; Garrido et al., 2015; Mattila et al., 2015). Thus,

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transcriptional regulation of GlyS and GlyP may facilitate glycogenmetabolism depending on life stages or in response to dietaryconditions. By contrast, the overexpression of Treh is sufficientto hydrolyze circulating trehalose. Considering that Tps1overexpression increases trehalose and leads to pupal lethality(Matsuda et al., 2015), transcriptional regulation is likely sufficientto facilitate trehalose metabolism. It will be interesting to explorefurther how the distinct regulatory systems in glycogen andtrehalose metabolism mirror the functional relevance of thesestorage sugars throughout the life cycle.A previous report indicated a significant drop in flight

performance in GlyP mutants (Eanes et al., 2006), but we did notobserve any such abnormality in ourGlyPmutants. This discrepancymight arise from the difference in experimental conditions; namely, atethered flight performance test focusing on wing beat frequency (inthe previouswork) versus a release-based flight test without tethering(in the present study). Alternatively, dietary conditionsmay affect theflight performance because starvation significantly decreasesclimbing activity in GlyP mutants. Interestingly, loss of neuronalglycogen improves locomotion ability during aging in Drosophila(Sinadinos et al., 2014). These observations raise the possibility thatlocal defects in glycogen metabolism result in distinct outcomescompared with inherited mutations. In humans, several inheritedmetabolic disorders, collectively referred to as glycogen storagediseases, are caused by deficiencies of enzymes involved in glycogensynthesis or breakdown (Özen, 2007; Adeva-Andany et al., 2016).An explicit comparison of GlyS and GlyP mutants will advance ourunderstanding of life stage-specific requirements for glycogenmetabolism with respect to carbohydrate metabolism and bodyhomeostasis.

MATERIALS AND METHODSDrosophila strainsThe following Drosophila melanogaster strains were used: w1118 (used as acontrol), UAS-Tps1, Tps1d2, Tps1MI03087 (Matsuda et al., 2015). y2 cho2 v1,P{nos-Cas9, y+, v+}/FM7c, KrGal4, UAS-GFP (CAS-0002) was obtainedfrom the National Institute of Genetics (NIG) Drosophila Stock Center.Df(3R)ED10561 (a deficiency strain of the GlyS locus, no. 150359) andDf(2L)ED119 (a deficiency strain of the GlyP locus, no. 150067) wereobtained from the Kyoto Drosophila Stock Center. The following stockswere obtained from the Bloomington Drosophila Stock Center (BDSC):GlyS-RNAi (34930),GlyP-RNAi (33634), hs-Gal4 (1799), tub-Gal4 (5138),Mef2-Gal4 (27390), elav-Gal4 (8760, 8765), Repo-Gal4 (7415), Cg-Gal4(7011), P{lacW}GlyPk07918 (10692), Mi{MIC}GlyPMI00957 (34131) andMi{MIC}GlySMI01490 (34440).

Of note, the original GlySMI01490 and GlyPMI00957 lines from the BDSCshowed complete lethality in the early larvae or embryo; however,transheterozygotes of each line over a deficiency line exhibited semi-lethality in larvae that resulted in the emergence of escaping adult flies,indicating that these Minos-insertion lines had second-site mutation(s).

Generation of the GlyS mutantsGeneration of the GlyS mutant allele was carried out using the CRISPR/Cas9 system. Sense and antisense oligonucleotides corresponding tosgRNA target sequences were annealed and inserted into BbsI-digestedpBFv-U6.2 vector (obtained from NIG). The GlyS sgRNA vector wasinjected into embryos carrying attP40 and nos-phiC31, and transgenicstrains were generated (BestGene). The nos-Cas9-based gene targeting wascarried out as previously described (Kondo and Ueda, 2013). Independentisogenized strains for each sgRNA construct were established. Indelmutations were analyzed via genome DNA extraction and PCRamplification of the DNA fragment including the target site, followed bysequence analysis. We isolated several frameshift mutations for GlyS usingtwo different sgRNAs. All isolated GlyS mutants were semi-viable; that is,approximately 10-30% survived to adulthood. We chose one strain, GlyS8,

for further analyses after backcrossing four times to the w− control strain.The backcrossed GlyS8 homozygous mutants in w− background were usedin all experiments unless otherwise noted.

Characterization and generation of GlyP mutantsA transposable P-element insertion P{lacW}k07918 (BDSC, 10692) wasbackcrossed to the w− control strain to remove the lethal mutation in the ftlocus as described previously (Tick et al., 1999). Consistent with this finding,transheterozygotes of the original P{lacW}k07918 line with a deficiencyallele lacking the GlyP locus, but not the ft locus, displayed homozygousviability. The backcrossed P-element line (w+) was used to generate GlyPmutants by imprecise excision. The progenywere first screened for the loss ofan eye color marker (w+), and the extent of the deletion in each mutant wasdetermined by PCR and subsequent DNA sequencing.GlyP3-11 andGlyP3-13

carried deletions of 1463 base pairs and 1710 base pairs, respectively, both ofwhich resulted in the lack of the entire first exon, including the transcriptionaland translational start sites. GlyP3-13 was backcrossed three times to the w−

control strain. The backcrossed GlyP3-13 homozygous mutants in w−

background were used in all experiments unless otherwise noted.

Fly foodDrosophila were reared on fly food (normal diet, ND) containing 8 g agar,100 g glucose, 45 g dry yeast, 40 g corn flour, 4 ml propionic acid and0.45 g butylparaben (in ethanol) per liter (1× recipe). For the analysis ofrestricted or enriched dietary conditions, fly food was prepared as describedpreviously (Matsuda et al., 2015; Yasugi et al., 2017). In brief, fly foodcontaining a reduced or increased amount of yeast or glucose was used (0.2×or 2× the amount used for the 1× recipe). The yeast-only diet was preparedaccording to the 1× recipe. No yeast paste was added to the fly tubes in anyof the experiments. All the experiments were conducted under non-crowdedconditions at 25°C. The food intake assay was carried out as describedpreviously (Yasugi et al., 2017).

qRT-PCR experimentsqRT-PCR analysis was performed as described previously (Okamoto et al.,2012; Okamoto and Nishimura, 2015). The primers used in this study havealso been described before (Yamada et al., 2018). The primers for GlyS andGlyP genes were located in the C terminus of the coding region.

Developmental staging and survival rateDevelopmental staging was performed essentially as previously described(Okamoto et al., 2012; Okamoto and Nishimura, 2015). Early third instarlarvae were defined as 0-6 h after second ecdysis. A defined number ofnewly hatched larvae were placed on each vial, and the numbers of pupaeand adults were counted. Embryos laid over a time period of 4 h were used todetermine the timing of pupariation. Pupariation was visually assessed twotimes per day, and the average time was calculated for each vial.Experiments were repeated at least twice, and all data were summed.

Quantification of weight and body sizeQuantification of adult weight was performed as previously described(Okamoto et al., 2013). Adult flies, pupae and larvae were photographedunder a Zeiss Stemi 2000-C stereomicroscope equipped with a CanonPowerShot G15 digital camera (Canon). The volumes of larvae and pupaewere determined using the formula 4/3π(L/2)(l/2)2, where L is the lengthand l is the diameter.

Plasmid constructionThe cDNAs encoding sTreh, cTreh, GlyS and GlyP were cloned by RT–PCR using sequenced strains (BDSC, 2057) and then subcloned intothe pCR-BluntII-TOPO vector (Invitrogen). The cDNAs were subclonedinto a modified pUAST vector that contained a C-terminal 1xFlagtag. Transformants were obtained using a standard injection method(BestGene).

Measurements of protein, TAG and sugar levelsProtein, TAG, trehalose, glycogen and glucose levels were measured asdescribed previously, with minor modifications (Matsuda et al., 2015).

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Frozen samples in tubes were homogenized using a pellet pestle in 100 µlof cold PBS containing 0.1% Triton X-100, immediately heat-inactivated at80°C for 10 min, and then cooled to room temperature (RT). Samples werefurther crushed to obtain uniform homogenates with 1× φ3-mm zirconiabeads using an automill (Tokken) at 41.6 Hz for 2 min. A portion ofhomogenate was mixed with triglyceride reagent (Sigma-Aldrich,),incubated at 37°C for at least 30 min, and then cleared by centrifugationat 20,000 g for 10 min. The supernatant was used for measurement of TAGusing free glycerol reagent (Sigma-Aldrich). A triolein equivalent glycerol(Sigma-Aldrich) was used as the standard. The amount of TAG wasnormalized to the total protein level.

Homogenate samples were further used to determine the glycogen levels,and the cleared samples after centrifugation at 10,000 rpm (9000 g) for10 min at RT were used to determine trehalose and glucose levels. A portionof homogenate was incubated with PBS containing bacterially producedrecombinant His-tagged Drosophila cTreh (Yoshida et al., 2016) oramyloglucosidase (Sigma-Aldrich) at 37°C overnight. A portion of samplewas incubated with PBSwithout enzymes in parallel for the determination ofglucose levels. The amounts of samples were adjusted based on priorexperience to obtain linearity within the range of standards. The reaction wascarried out in a 15-µl assay mixture. Glucose levels were determined using aglucose assay kit (Sigma-Aldrich). A serial dilution of glucose was used as astandard. The trehalose and glycogen concentrations for each sample weredetermined by subtracting the values of free glucose in the untreated samples.Trehalose, glycogen and glucose levels were normalized to the protein level,which was determined by a BCA protein assay kit (Thermo Scientific).

Metabolite extraction and a widely targeted metabolomicsprofileFrozen samples in 1.5 ml plastic tubes were homogenized in 300 µl of coldmethanol with 1× φ3-mm zirconia beads using an automill (Tokken) at41.6 Hz for 2 min. The homogenates were mixed with 200 µl of methanol,200 µl of H2O and 200 µl of CHCl3 and then vortexed for 20 min at RT. Thesamples were centrifuged at 15,000 rpm (20,000 g) for 15 min at 4°C. Thesupernatant was mixed with 350 µl of H2O and vortexed for 10 min at RT.The aqueous phase was collected after centrifugation and dried in a vacuumconcentrator. The samples were re-dissolved in 2 mM ammoniumbicarbonate (pH 8.0) and analyzed by liquid chromatography withtandem mass spectroscopy (LC-MS/MS). The insoluble pellets werewashed with 90% ethanol, re-dissolved with PBS containing 0.1% TritonX-100, and used for quantification of glycogen after treatment withamyloglucosidase. The samples were further mixed with two volumes of0.2 NNaOH, heat-denatured, and used to quantify total protein using a BCAprotein assay kit (Thermo Scientific).

An in-house platform for a widely targeted analysis was established bymaking scheduled multiple reaction monitoring (MRM) methods usingindividual authentic compounds and biological samples. Chromatographicseparations in an Acquity UPLC H-Class System (Waters) were carried outunder reverse-phase conditions using an Acquity UPLCHSS T3 column andunder HILIC conditions using an Acquity UPLC BEH Amide column. Theionized compounds were detected using a Xevo TQD triple quadrupole massspectrometer coupled with an electro-spray ionization source (Waters). Thepeak area of a target metabolite was analyzed using MassLynx 4.1 software(Waters).Metabolite signals were then normalized to the total protein level ofthe corresponding sample after subtracting the values from the blank sample.P-values were calculated by two-tailedWelch’s t-test using Microsoft Excel.An estimate of the false discovery rate (FDR), which is given by the q-value,was calculated to take into account the multiple comparisons. Furtherstatistical analyses were performed using MetaboAnalyst 4.0 (Xia et al.,2009). For multivariate analysis, PC analysis was performed to identifybiologically relevant classifications (Worley and Powers, 2013). Data werenormalized to the median per sample. Heat map and hierarchical clusteringwere generated using Pearson correlations and Ward’s method. Enrichmentanalysis was performed using MetaCore software ver. 6.37 (ClarivateAnalytics). Networks with In Data>2 are shown in Table S1.

For the TAG analysis, the organic phase was collected after the additionof hexane. Dried samples were re-dissolved in isopropanol and analyzed byLC-MS/MS using an Acquity BEH C18 column (Waters).

Preparation of 13C-labeled glycogenControl pupae raised on a normal diet were transferred to a new vialcontaining a diet composed of 5% yeast and either 5% D-glucose (WakoChemical) or 5% D-[U-13C6] glucose (Cambridge Isotope Laboratories).Adult flies were collected 4 days after eclosion and transferred to 1.5 mltubes (100 flies per tube). Samples were homogenized in 1 ml of 10%trichloroacetic acid and cleared by centrifugation at 15,000 rpm (20,000 g)for 10 min at RT. The supernatant was diluted with 10 times volume ofethanol in a 50 ml tube. The pellet containing glycogen was recovered bycentrifugation at 15,000 rpm (20,000 g) for 10 min at 4°C and washed oncewith 90% ethanol. Glycogen was dissolved in 1× PBS, filtered through anAmicon filter (100 kDa MWCO), and washed five times with 1× PBS toremove low-molecular weight contaminants.

The purity of glycogen was confirmed by a high-performance liquidchromatography (Alliance 2695, Waters) coupled with a refractive indexdetector (Waters 2414). Size-exclusion chromatographywas performed at 40°C using a Shodex OHpak SB-806M HQ column (8.0×300 mm, Shodex).Elution was carried out with 1× PBS at a flow rate of 0.5 ml/min isocratic.Pullulan standards were obtained from Shodex. The proportion of glucoseisotopologues in purified glycogen was analyzed by LC-MS/MS aftertreatment with amyloglucosidase (Sigma).

GlyS enzyme assayTen early third instar larvae were dissected in PBS. Dissected tissues werehomogenized in cold 100 µl 2× homogenize buffer [2× PBS, 4 mMMgCl2,0.2% Triton X-100, 2× complete proteinase inhibitor cocktail (Roche) and1 mM PMSF] with motorized pestles. Samples (10 µl) were mixed on icewith 10 µl of 2× GlyS assay mixture containing 3.4 mM UDP-[U-13C6]glucose (Omicron Biochemicals), 76 mM glucose-6-phosphate (WakoChemical) and 200 µg rabbit liver glycogen (Wako Chemical) inH2O. Reactions were started by placing samples on a 30°C blockincubator for the indicated times and stopped by the addition of 200 µl10% trichloroacetic acid. The samples were cleared by centrifugation at15,000 rpm (20,000 g) for 10 min at RT. The supernatant was diluted with1 ml ethanol. The pellet containing glycogen was recovered bycentrifugation at 15,000 rpm (20,000 g) for 10 min at 4°C and washedoncewith 90% ethanol. The pellet was dissolved in 20 µl 1× PBS containingamyloglucosidase (Sigma). After incubation overnight at 37°C, sampleswere mixed with 500 µl acetonitril containing 50 ng 13C1-mannitol (Sigma)as an internal standard and cleared by centrifugation at 15,000 rpm (20,000g) for 10 min at 4°C. The supernatant was loaded onto a MonoSpin Amidecolumn (GL Science). The bound metabolites were eluted with 300 µl H2Oand then vacuum dried. Samples were re-dissolved in 2 mM ammoniumbicarbonate and analyzed by UPLC-MS/MS using an Acquity BEH Amidecolumn (Waters). The MRM transitions (negative ion mode) were asfollows: 13C1-Mannitol m/z 182>89, 13C6-glucose m/z 185>92. Theincorporation rate of 13C6-glucose into glycogen was determined afternormalization with values of 13C1-mannitol.

GlyP enzyme assayTissue homogenates were prepared as described above. Samples (10 µl)were mixed on ice with 10 µl 2× GlyP assay mixture containing 200 µg13C6-labeled glycogen in H2O. Reactions were started by placing sampleson a 30°C block incubator for the indicated times and stopped by theaddition of 500 µl acetonitrile containing 50 ng 13C1-mannitol (Sigma). Thesamples were cleared by centrifugation at 15,000 rpm (20,000 g) for 10 minat 4°C. The supernatant was loaded onto a MonoSpin Amide column (GLScience). The bound metabolites were eluted with 300 µl H2O and vacuumdried. Samples were re-dissolved in 2 mM ammonium bicarbonate andanalyzed by LC-MS/MS using an Acquity HSS T3 column (Waters). MRMtransitions (positive ion mode) were as follows: 13C6-glucose-phosphate(mixture of glucose-1-phosphate and glucose-6-phosphate) m/z 267>99.The release rate of 13C6-glucose-phosphate from 13C6-labeled glycogen wasdetermined after normalization with values of 13C1-mannitol.

HistochemistryPolysaccharide staining, including glycogen, was performed as describedpreviously (Yamada et al., 2018). Larval tissues were dissected in PBS

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containing 1% bovine serum albumin (PBSB), fixed using 3.7%formaldehyde in PBS for 20 min, washed twice in PBSB, incubated withperiodic acid solution (Merck) for 5 min, and washed twice in PBSB.Samples were stained with Schiff’s reagent (Merck) for 15 min, washedtwice in PBSB, and mounted in 50% glycerol in PBS. Images were acquiredwith a Zeiss Primo Star microscope equipped with an AxioCam ERc(Zeiss). The experiments were conducted independently at least twice, andrepresentative images obtained from several larvae are presented.

ImmunohistochemistryLarval tissues were dissected in PBS, fixed for 10 min in 3.7% formaldehydein PBS containing 0.2%TritonX-100, and processed as previously described(Wirtz-Peitz et al., 2008). Mouse anti-Flag M2 antibody (Sigma, F1804,1:500) was used as the primary antibody. Alexa-conjugated secondaryantibodies (A11029, 1:500) and phalloidin (Thermo Fisher Scientific) wereused. Images were acquired with a Zeiss LSM700 confocal microscope andprocessed in Photoshop (Adobe Systems).

Climbing assayGroups of 15-20 male flies, reared under conditions of controlled populationdensity, were collected within 24 h of eclosion. The flies were transferred tofresh vials every second or third day until the indicated ages, without furtherexposure to CO2. Before testing, the flies were transferred to a 100 ml plasticgraduated cylinder and allowed to rest for 10 min. The flies were tappeddown to the bottom of the cylinder, and their locomotion was recorded usinga Canon PowerShot G1X digital camera (Canon). The first five trials wereconducted for habituation to the environment, and the following three trialswere used for the analysis. The average climbing speed (cm/s) wasdetermined in ImageJ software by tracing the position of flies every seconduntil they reached the top. The experiment was repeated with at least twoindependently reared replicates, and all data were summed.

Flight testA φ20×100 cm acrylic cylinder was used for the flight test. Groups of tenmale flies (2-5 days after eclosion) were placed in the top of the cylinderthrough a funnel to initiate flight. The height at which flies landed in thecylinder was recorded by a Canon PowerShot G15 digital camera (Canon)and judged as their flight ability. The experiment was repeated with at leasttwo independently reared populations, and all data were summed.

Lifespan and transient starvation assayFlies were reared under conditions of controlled population density andtemperature. Groups of 20 flies (males for lifespan assay, males and femalesseparately for starvation assay) were collected within 24 h of eclosion. Theflies were transferred to fresh vials twice per week, and the number of deadflies was recorded. For the transient starvation assay, 10-day-old flies weretransferred to a vial that contained 0.8% agar, and the number of dead flieswas recorded. For each genotype, at least four independent cohorts of flieswere analyzed. Kaplan–Meier lifespan analysis was performed, andP-values were calculated using log-rank statistical analysis.

Statistical analysis and data presentationStatistical tests were performed using Microsoft Excel or GraphPad Prism 6software. Box and violin plots were drawn online using the BoxPlotRapplication (http://boxplot.tyerslab.com/). For box plots, centerlines showthe medians, box limits indicate the 25th and 75th percentiles, whiskersextend 1.5 times the interquartile range from the 25th and 75th percentiles,and outliers are represented by dots. For violin plots, white circles show themedians, box limits indicate the 25th and 75th percentiles, whiskers extend1.5 times the interquartile range from the 25th and 75th percentiles, andpolygons represent density estimates of data and extend to extreme values.

AcknowledgementsWe thank the Bloomington Drosophila Stock Center, the Kyoto Stock Center, andthe National Institute of Genetics Drosophila Stock Center for the fly stocks. We alsothank members of fly laboratories in RIKEN BDR for their valuable support anddiscussion; the genome resource and analysis unit in RIKEN BDR for their technicalsupport; and K. Hironaka, R. Niwa and S. K. Yoo for discussions and comments onthe manuscript.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsConceptualization: T.Y., T.N.; Validation: T.Y., O.H., T.N.; Formal analysis: T.Y.,O.H., Y.Y., R.M., H.K., Y.N., T.N.; Investigation: T.Y., O.H., Y.Y., R.M., H.K., T.N.;Writing - original draft: T.N.; Writing - review & editing: T.Y., Y.Y., Y.N.; Visualization:T.Y., T.N.; Supervision: T.N.; Project administration: T.N.; Funding acquisition: T.N.

FundingThis work was supported, in part, by the Japan Society for the Promotion of Science(JSPS) [KAKENHI grants JP17K19433 and JP17H03658 to T.N.].

Supplementary informationSupplementary information available online athttp://dev.biologists.org/lookup/doi/10.1242/dev.176149.supplemental

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