Cross-talk between the fat body and brain regulates insect ... · Cross-talk between the fat body and brain regulates insect developmental arrest Wei-Hua Xua,1, Yu-Xuan Lua, and David

Cross-talk between the fat body and brain regulatesinsect developmental arrestWei-Hua Xua,1, Yu-Xuan Lua, and David L. Denlingerb,c,1

aState Key Laboratory of Biocontrol, School of Life Sciences, Sun Yat-Sen University, Guangzhou 510006, China; and Departments of bEntomologyand cEvolution, Ecology and Organismal Biology, Ohio State University, Columbus, OH 43210

Contributed by David L. Denlinger, July 27, 2012 (sent for review June 22, 2012)

Developmental arrest, a critical component of the life cycle inanimals as diverse as nematodes (dauer state), insects (diapause),and vertebrates (hibernation), results in dramatic depression of themetabolic rate and a profound extension in longevity. Althoughmany details of the hormonal systems controlling developmentalarrest are well-known, we know little about the interactions be-tween metabolic events and the hormones controlling the arrestedstate. Here, we show that diapause is regulated by an interplaybetween blood-borne metabolites and regulatory centers within thebrain. Gene expression in the fat body, the insect equivalent of theliver, is strongly suppressed during diapause, resulting in low levelsof tricarboxylic acid (TCA) intermediates circulating within the blood,and at diapause termination, the fat body becomes activated,releasing an abundance of TCA intermediates that act on the brainto stimulate synthesis of regulatory peptides that prompt productionof the insect growth hormone ecdysone. This model is supported byour success in breaking diapause by injecting a mixture of TCAintermediates and upstream metabolites. The results underscorethe importance of cross-talk between the brain and fat body asa regulator of diapause and suggest that the TCA cycle may bea checkpoint for regulating different forms of animal dormancy.

prothoracicotropic hormone | glucose | pyruvate

As days shorten in late summer and temperatures drop, manyinsects respond by entering an overwintering diapause,

a form of developmental arrest characterized by metabolic de-pression. The major endocrine events that regulate diapause arefairly well-understood. In larvae and pupae, the arrest is usuallya consequence of the brain’s failure to produce or release pro-thoracicotropic hormone (PTTH), a neuropeptide needed tostimulate the prothoracic gland to synthesize the steroid hor-mones ecdysteroids (20-hydroxyecdysone is the most active formand will be referred to hereafter as ecdysone) (1). Without ec-dysone, the insect remains locked in a developmental arrest thatpersists as long as ecdysone is absent.Specific patterns of gene expression and unique metabolic

profiles characterize diapause (2–4), but the interactions betweengenes, metabolites, and the major endocrine centers are poorlyknown. One of the most conspicuous metabolic patterns duringdiapause in insects and dormancy in other animals (5–7) is a shiftto anaerobic metabolism favoring glycolysis and gluconeogenesis.Although it is usually assumed that changes in abundance ofspecific metabolites are downstream responses to the diapauseprogram, the demonstration that elevated sorbitol is the cause,rather than the consequence, of developmental arrest in embryosof the silk moth (8) suggests the possibility that the metaboliteprofile itself may influence the diapause decision. This possibilityis tested here by monitoring changes in metabolite abundance inassociation with diapause and then showing that artificiallyboosting the abundance of nondiapause metabolites can elevatemRNA levels of PTTH in the brain and prompt the terminationof diapause. The results that we present for regulation of pupaldiapause in moths of the Heliothis/Helicoverpa complex of agri-cultural pests suggest cross-talk between the brain and fat body

that is based on the abundance of tricarboxylic acid (TCA)-related metabolites present in the hemolymph.

Results and DiscussionAs predicted from results with other diapausing pupae (2), thepupal diapause of H. armigera is characterized endocrinologicallyby a low ecdysone titer (Fig. S1). Pupae programmed by long daylength release a pulse of ecdysone that prompts continuous de-velopment, whereas those pupae that receive short days fail toproduce ecdysone and hence, remain locked into a develop-mental arrest and metabolic slowdown in the pupal stage.This metabolic slowdown is especially obvious in the fat body.

A sampling of 60 randomly selected genes expressed in the brainof H. armigera showed a nearly equal proportion of up- anddown-regulated genes (27% up-regulated genes, 27% down-regulated genes, and 47% no change genes) (Table 1). Thisfinding is in marked contrast to results noted in the fat body.Among 55 genes that were sampled, only 7% were up-regulated,60% were down-regulated, and no change was noted in 33% ofgenes (Fig. S2). These results suggest that a slowdown in activityof the fat body is especially noteworthy during diapause, a resultconsistent with the shutdown of the liver in hibernating mammals(9). This conclusion is also supported by a comparison of COXactivity and ATP content in the brain and fat body (Fig. 1). Al-though COX activity and ATP content are lower during diapausein both the brain and fat body, suppression during diapause ismuch more pronounced in the fat body. The fat body, whichfunctions much like the mammalian liver, is the key site of inter-mediary metabolism (10, 11), and thus, the pronounced shut-down of this organ is likely responsible for the major metabolicshifts noted in association with diapause (2, 3, 12). The fact thatgene expression in the fat body rapidly responds to an injectionof ecdysone (Fig. S3) implies that the fat body is responsive tothe ecdysone titer, and low ecdysone is essential for the low levelsof fat body gene expression and enzyme activity during diapause.Metabolomic profiling of the hemolymph reveals numerous

distinctions between diapausing and nondiapausing pupae, es-pecially among intermediates in the TCA cycle (Fig. 2). Levels ofthe insect blood sugar, trehalose, as well as isocitrate are higherin the hemolymph of diapausing pupae, whereas levels of glu-cose, pyruvate, fumarate, and malate are lower in diapausingpupae (Fig. 2A). When diapause was terminated artificially with aninjection of ecdysone, levels of these hemolymph metabolites shif-ted: trehalose and isocitrate declined, whereas increases were notedfor glucose, pyruvate, fumarate, succinate, and malate (Fig. 2B).Monitoring metabolic profiles in the brain and hemolymph

from the time of pupation until the entry into diapause on day 10again points to distinctions between these two developmental

Author contributions: W.-H.X. and D.L.D. designed research; Y.-X.L. performed research;W.-H.X., Y.-X.L., and D.L.D. analyzed data; and W.-H.X. and D.L.D. wrote the paper.

The authors declare no conflict of interest.1To whom correspondence may be addressed. E-mail: [email protected] [email protected].

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1212879109/-/DCSupplemental.

www.pnas.org/cgi/doi/10.1073/pnas.1212879109 PNAS | September 4, 2012 | vol. 109 | no. 36 | 14687–14692

PHYS

IOLO

GY

Dow

nloa

ded

by g

uest

on

June

5, 2

020

states (Fig. S4). The most striking differences center on trehaloseand glucose. Brain levels of trehalose and glucose remain fairlylow and unchanged with time in nondiapausing pupae, but bothtrehalose and glucose levels progressively increase in the brainsof diapause-destined pupae. Within the hemolymph, trehaloselevels in diapause-destined pupae remain elevated, whereaslevels in nondiapausing pupae decline during this 10-d interval.Glucose levels differ significantly only on day 10, at which timeglucose is higher in nondiapausing pupae. Pyruvate, isocitrate,succinate, fumarate, and malate are all substantially moreabundant in the brains of nondiapausing pupae. Hemolymphdifferences are less pronounced, but the pyruvate level is againhigher in nondiapausing pupae.Glucose and pyruvate, which are upstream of the TCA cycle,

may act as major limiting factors to depress TCA cycle activityin the brain when they are present in low amounts in the he-molymph. This finding is consistent with the hypothesis thatdiapause entrance accompanies a decrease in intermediatesfrom the fat body, resulting in a significant reduction of TCA

activity in the brain, potentially leading to developmental arrestin the pupae.The differences in metabolite profiles of diapausing and

nondiapausing pupae of H. armigera suggest the possibility thatthis information may be used as a component of the signalingsystem that regulates diapause. We tested this idea by injectingglucose or a mixture of four metabolites that increased inabundance at diapause termination (Fig. 2B) (glucose, pyruvate,fumarate, and malate) into diapausing pupae to see if theseagents are capable of breaking diapause. As shown in Table 2,glucose by itself wasmoderately effective in breaking diapause, butthe effect was much more pronounced when the mixture of fourmetabolites was injected. Fewer than 10% of controls broke dia-pause, whereas>70% of pupae injected with a high concentrationmixture broke diapause, and the response was dose-dependent.We tested whether injection of metabolites can terminate

diapause in other species within this pest complex as well. Thefour-component mixture that was active in H. armigera was alsoeffective in terminating pupal diapause in the corn earworm,

Table 1. Changes of gene expression in brain and fat body of diapausing pupae compared withexpression levels in nondiapausing pupae

Genes examined Down-regulation in diapause Up-regulation in diapause Equivalent

Brain 60 16 (27%) 16 (27%) 28 (47%)Fat body 55 33 (60%) 4 (7%) 18 (33%)

Four cDNA libraries constructed from brains and fat body of H. armigera yielded 3,586 genes. RT-PCR was usedto monitor expression in 60 selected genes from the brain and 55 selected genes from the fat body. Genes werecategorized as being down- or up-regulated in diapause or showing equivalent levels of expression. A 1.5-folddifference in expression was defined as the cutoff for differential expression.

Brain

Fat body

A

B

Fig. 1. COX activity and ATP content in the (A) brain and (B) fat body of nondiapausing pupae (ND), diapausing pupae (D), and diapausing pupae injectedwith 1 μg 20-hydroxyecdysone (20E). Numbers represent hours after injection of 20E or distilled water (H2O). Control, no injection. Bars represent mean ± SDof three replicates. *P < 0.05; **P < 0.01 (determined by one-way ANOVA).

14688 | www.pnas.org/cgi/doi/10.1073/pnas.1212879109 Xu et al.

Dow

nloa

ded

by g

uest

on

June

5, 2

020

H. zea (Fig. S5), thus implying that this response is not restrictedto H. armigera.

If metabolite levels exert a direct effect on development, wemight also expect to be able to affect the progression of de-velopment in nondiapausing individuals by manipulating me-tabolite levels. This affect on progression does, indeed, occur.2-Deoxy-glucose (DOG) is a glucose derivative in which the2-hydroxyl group is replaced by hydrogen. Although DOG isreadily taken up by cells, the molecule is unable to undergoglycolysis. Development is delayed when DOG is injected intopupae programmed for nondiapause (Fig. 3A). Migration of thepupal stemmata is a good landmark for monitoring progressionof pupal development (13), and this stage is usually completed4–5 d after pupation. Pupae injected with DOG on day 1 re-quired 7–8 d to reach this same stage, thus implying that DOGinjection delays the normal progression of development.Diapause entrance is accompanied by a decrease in the release

of intermediates from the fat body, resulting in a significant re-duction of TCA activity in the brain. When we injected diapause-programmed pupae (4 d after pupation) with a mixture of fourmetabolites (two TCA cycle intermediates and two metabolitesupstream of the TCA cycle), diapause was averted, and the pu-pae proceeded with nondiapause development. Glucose alonedid not elicit this effect (Table 3); the entire mixture was needed.These observations are consistent with a critical role for themetabolic intermediates in affecting developmental decisions.As shown in Fig. 3B, injection of the metabolite mixture elicits

elevation of PTTH mRNA, which in turn, likely leads to syn-thesis and release of PTTH and subsequent stimulation of the

Nondiapause and diapause differences

Changes in diapausing pupae after injection of 20-hydroxyecdysone

A

B

Fig. 2. Differences in metabolic intermediates present in the hemolymph of (A) nondiapausing and diapausing pupae and (B) after diapause termination inresponse to an injection of 20E. Metabolic intermediates, including trehalose, glucose, pyruvate, and four substances in the TCA cycle, were compared: day 3nondiapause pupae (ND), day 21 diapausing pupae (D), and day 21 diapausing pupae injected with 1 μg 20E. Relative ratio is the ratio of the peak area ofmetabolic intermediate to the peak area of the internal standard (sucrose). Hemolymphs from 10 pupae were mixed as one sample; bars represent themean ± SD of four replicates. *P < 0.05; **P < 0.01 (determined by one-way ANOVA).

Table 2. Termination of pupal diapause in response to aninjection of metabolites

n Percent diapause termination ± SD

No treatment 63 4.4 ± 4.4H2O 118 8.7 ± 5.225 μg glucose 77 37.3 ± 11.8*100 μg glucose 77 41.8 ± 6.2*150 μg glucose 58 51.7 ± 3.9*5× mixture 88 32.7 ± 5.6*20× mixture 90 53.0 ± 6.7*20× mixture† 65 63.5 ± 7.4*30× mixture 80 72.7 ± 8.8*

Diapause pupae were kept at 20 °C for 21 d after pupation, injected witha 5 μL solution, and then incubated at 22.5 °C. Diapause termination wasdetermined by examining the pupal stemmata after injection of metabo-lites. The 1× mixture contained 5 μg glucose, 4 μg pyruvate sodium, 10 μgmalate, and 2 μg fumarate, and it is equal to the physiological concentrationin diapause pupa. The 20× and 30× mixtures contained only 20 μg fumarate,because fumarate did not dissolve when the concentration was higher than20 μg. Percentage diapause termination was calculated 15 d after injection.Diapause termination was compared with the injection of distilled water.*P < 0.01 (determined by one-way ANOVA).†The 20× mixture contained 30× glucose, but the other three metaboliteswere at 20×.

Xu et al. PNAS | September 4, 2012 | vol. 109 | no. 36 | 14689

PHYS

IOLO

GY

Dow

nloa

ded

by g

uest

on

June

5, 2

020

prothoracic gland to synthesize ecdysone, an event well-known tobreak diapause (1).The results suggest complexity in the regulation of insect dor-

mancy that goes beyond the simple prevailing models that suggestthat diapause is a simple shutdown in the release of a singlehormone. Work with Drosophila indicates that the fat body affectsgrowth and development by regulating synthesis and release of theneurohormone insulin from the brain (14, 15), and our experi-ments with H. armigera show that the fat body can exert an effecton the brain regulatory center by altering sugar levels and levels of

TCA metabolites. Injection of ecdysteroids can artificially breakdiapause, a response accompanied by elevation of COX activityand ATP content in the fat body as noted (Fig. 1), and causea switch in hemolymph metabolite levels from the diapause to thenondiapause profile (Fig. 2B). Together, the results describedhere are consistent with cross-talk between biochemical processesin the fat body and brain (Fig. 3C). Elevation of glucose, pyruvate,and two TCA cycle intermediates, products of the fat body,prompts termination of diapause by elevating PTTH in the brain,leading to synthesis of ecdysone in the prothoracic gland.

a. Nondiapause b. Diapause

A B

C

Fig. 3. Manipulations of development with a nonusable glucose derivative and a metabolic mixture and a regulatory scheme defining documentedinteractions. (A) Developmental delay caused by DOG injection. Nondiapause pupae were kept at 20 °C for 1 d after pupation, injected with a 5 μL solution ofDOG, and incubated at 20 °C. The developmental delay was determined by examining the location of the pupal stemmata on different days after injection.No treatment, n = 40; distilled water (H2O), n = 29; 0.2 mM DOG, n = 31; 2 mM DOG, n = 30. (B) Change in PTTH mRNA expression after injection ofa metabolite mixture into 21-d-old diapausing pupae. The 30× mixture is described in Tables 2 and 3. Bars represent the mean ± SD of three replicates. *P <0.05 (determined by one-way ANOVA). (C) A schematic representation showing the action of metabolites on the brain, prothoracic glands (PGs), and fat bodyin the regulation of (a) nondiapause and (b) diapause. Black lines indicate demonstrated pathways (pathway 1), and red lines (pathways 2 and 3) indicateresults from this study. Arrows and broken arrows indicate activation and no activation, respectively.

14690 | www.pnas.org/cgi/doi/10.1073/pnas.1212879109 Xu et al.

Dow

nloa

ded

by g

uest

on

June

5, 2

020

Conversely, a decrease of ecdysteroid levels by the down-regula-tion of PTTH causes a drop of fat body activity in diapause-typeindividuals, which is noted by COX activity and ATP content, andit results in a decrease in the levels of metabolites in the fat body.Down-regulation in the generation of fat body metabolites resultsin fewer TCA-related metabolites in the hemolymph and brain,thus suggesting a regulatory pathway linking TCA activity andhormone action in the regulation of insect diapause (Fig. 3C).Other complexities certainly contribute to diapause regulation aswell. For example, diapause in members of the Heliothis/Heli-coverpa complex can also be terminated by the neuropeptidediapause hormone (1, 16, 17), but we do not yet know how dia-pause hormone or other regulatory molecules, such as insulin (14,15), may be linked to the scheme presented in Fig. 3C.One attractive corollary suggested by the model that we

present is that metabolite levels could offer a mechanism forsensing energy reserves during diapause. One of the criticaldecisions that an insect needs to make during diapause is how tocarefully parse out those reserves so that it can bridge the wintermonths and still retain sufficient reserves for completing de-velopment in the spring (18). How an insect is able to monitorthose reserves and break diapause when the reserves are be-coming depleted remains unknown, but the cross-talk that wereport here could very well contribute to a scheme for achievingthis crucial form of energy management.In summary, several lines of evidence suggest that metabolites

fueling the TCA cycle may be a checkpoint for developmentalarrest in diverse animal groups. (i) Developmental arrest resultsin a similar phenotype of metabolic depression in nematodes,insects, and mammals. (ii) The fat body and its vertebrateequivalent, the liver, are the tissues most profoundly affectedduring dormancy, which was indicated by the high proportion ofdown-regulated genes in these tissues. (iii) Low TCA activity isa feature shared by both insects and the dauer state in Caeno-rhabditis elegans (19). (iv) Injection of DOG can mimic de-velopmental delay, which was noted in insects and mammalianhibernation (20). Together, these results suggest some commonalityin the schemes used by animals to regulate developmental arrest.

Materials and MethodsInsects. Colonies of H. armigera and H. zea were maintained in our labora-tories on an artificial diet at 20 °C and a photoperiod of light to dark of14:10 h to generate nondiapause pupae and 20 °C and short day length(light to dark of 10:14 h) to generate diapause pupae. After pupation, thebrain and fat body were dissected in a 0.75% NaCl solution and stored in−80 °C.

Measurement of COX Activity and ATP Content. COX activity was measured asdescribed (21). In brief, brains and fat bodies were homogenized in 110 μLradioimmunoprecipitation (RIPA) buffer (50 mM Tris·HCl, pH 8.0, 150 mMNaCl, 1% Nonidet P-40, 0.1% SDS, 0.5% sodium deoxycholate) and centri-fuged at 12,000 × g at 4 °C. COX activity in the brain and fat body wasmeasured using a COX assay kit (Sigma). The 50 μL COX extraction, 50 μLenzyme dilution buffer (10 mM Tris·HCl, pH 7.0, 250 mM sucrose), and 950 μLassay buffer (10 mM Tris·HCl, pH 7.0, 120 mM KCl) were mixed, 50 μL fer-roytochrome c substrate solution were added to the mixture, and the samplewas read at A550/min immediately at room temperature.

Brains and fat bodies were homogenized in 400 μL 6% HClO4 andcentrifuged at 12,000 × g at 4 °C. Then, 450 μL supernatant were transferredto a new 1.5-mL microtube, 60 μL 3 M K2CO3 were slowly added, and themixture was centrifuged at 12,000 × g at 4 °C. ATP content in the brain andfat body was measured using an ATP Bioluminescent Assay Kit (Sigma); 50 μLATP assay mix solution were added to a 96-well plate, swirled, and incubatedfor 3 min at room temperature. A 5-μL sample and 45 μL distilled H2O wereadded, swirled briskly to mix, and immediately measured to read theamount of light produced by VICTOR ×5 (Perkin-Elmer).

Sample Extraction and Derivatization for Metabolites. For hemolymph sampleextraction and derivatization, 100 μL hemolymph were added to 100 μLdistilled water (containing 0.1 mg/mL sucrose as an internal standard) and800 μL methanol. The solution was vortex-mixed for 10 s, incubated on icefor 1 h, and centrifuged at 20,000 × g at 4 °C. Then, 600 μL supernatant weretransferred to a GC vial and evaporated to dryness in a vacuum concentrator;90 μL freshly prepared methoxylamine hydrochloride (15 mg/mL in pyridine;Sigma) were added to the vial, vortex-mixed for 10 min, and incubated atroom temperature for 16 h. The sample was trimethylsilylated by the additionof 90 μL N-Methyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA) [containing1% trimethylchlorosilane (TMCS); Sigma] and incubated at room temperaturefor 1 h. Finally, 120 μL heptane were added to stop the reaction.

Thirty brains of each sample were extracted and derivatized as described(22). In brief, each sample was homogenized in a 600 μL methanol–chloro-form mixture (2:1) and sonicated for 15 min. Then, 200 μL chloroform and200 μL H2O (containing 0.01 mg/mL sucrose as an internal standard) wereadded to the sample and centrifuged at 20,000 × g for 10 min. The aqueouslayer was transferred to a GC vial followed by evaporation. The dried samplewas dissolved in 10 μL freshly prepared methoxylamine hydrochloride (15mg/mL in pyridine; Sigma), vortex-mixed for 5 min, and held at room tem-perature for 16 h. The sample was added to 10 μL MSTFA (containing 1%TMCS; Sigma) and incubated at room temperature for 1 h. Finally, 20 μLheptane were added to stop the reaction.

GC-MS Analysis for Metabolites. The GC-MS analysis program was described(22) and slightly modified. The 1 μL derivatization sample was autoinjectedusing a 1:10 split ratio to a Trace GC Ultra-DSQ II GC-MS (Thermo), with theinjector temperature held at 280 °C. The oven temperature ranged from50 °C to 300 °C (5 °C/min), and then, it was maintained at 300 °C for 5 min.The column was DB-5msUI (length = 30 m, i.d. = 25 mm; Agilent), and themass spectra were scanned from 50 to 450 m/z. Peak identification wascompared with retention time with authentic standards. A peak that lackedauthentic standards was identified by comparison with the National Instituteof Standards and Technologies standards, and a reverse similarity index (RSI)value > 700 was considered to be a good match. Individual integrated peakareas were compared with the internal standard (sucrose) peak area.

Additional details on construction of the cDNA library, sequencing,analysis and annotation the EST, gene expression analysis, and RIA forecdysteroids are presented in SI Materials and Methods.

ACKNOWLEDGMENTS. We thank Dr. Qirui Zhang (Ohio State University) forevaluating the response in Helicoverpa zea and Drs. Vladimir Kostal (Insti-tute of Entomology, Czech Republic), Daniel Hahn (University of Florida),S. Reddy Palli (University of Kentucky), and Xavier Belles (Institut de BiologiaEvolutiva, Barcelona, Spain) for critical reading of the paper. This study wassupported by Natural Scientific Foundation of China Grant-in-Aid 30730014,National Basic Research Program of China, Ministry of Science and Technol-ogy Grant 2012CB114101 (to W.-H.X.), and US Department of Agriculture(USDA)-National Institute of Food and Agriculture (NIFA) Grant 2011-67013-30199 (to D.L.D.).

1. Zhang Q, Denlinger DL (2012) Dynamics of diapause hormone and prothoracicotropichormone transcript expression at diapause termination in pupae of the corn ear-worm, Helicoverpa zea. Peptides 34:120–126.

2. Denlinger DL, Yocum GD, Rinehart JP (2005) Comprehensive Insect Molecular Science,eds Gilbert LI, Iatrou K, Gill SS (Elsevier, Amsterdam), Vol 3, pp 615–650.

3. Kostál V (2006) Eco-physiological phases of insect diapause. J Insect Physiol 52:113–127.

4. Ragland GJ, Denlinger DL, Hahn DA (2010) Mechanisms of suspended animation arerevealed by transcript profiling of diapause in the flesh fly. Proc Natl Acad Sci USA107:14909–14914.

Table 3. Developmental fate of diapause-programmed pupaeinjected with glucose or a metabolite mixture before the onset ofdiapause

n Percent development ± SD

No treatment 26 19.0 ± 3.4H2O 34 20.7 ± 5.7150 μg glucose 35 34.1 ± 7.21,500 μg glucose 35 31.6 ± 5.930× mixture 35 65.4 ± 10.3*

Diapause-programmed pupae were kept at 20 °C for 4 d after pupation,injected with a 5 μL solution as reported in Table 2, and then held at 22.5 °C.Pupal development was determined on day 15 by examining the location ofthe pupal stemmata.*P < 0.01 (determined by one-way ANOVA).

Xu et al. PNAS | September 4, 2012 | vol. 109 | no. 36 | 14691

PHYS

IOLO

GY

Dow

nloa

ded

by g

uest

on

June

5, 2

020

5. Hu PJ (2007) The C. elegans Research Community, WormBook. Available at http://

www.wormbook.org. Accessed August 8, 2007.6. Storey KB, Heldmaier G, Rider MH (2010) Dormancy and Resistance in Harsh Envi-

ronments, eds Lubzens E, et al. (Springer, Berlin), pp 227–252.7. Guppy M, Withers P (1999) Metabolic depression in animals: Physiological per-

spectives and biochemical generalizations. Biol Rev Camb Philos Soc 74:1–40.8. Horie Y, Kanda T, Mochida Y (2000) Sorbitol as an arrester of embryonic development

in diapausing eggs of the silkworm, Bombyx mori. J Insect Physiol 46:1009–1016.9. Williams DR, et al. (2005) Seasonally hibernating phenotype assessed through tran-

script screening. Physiol Genomics 24:13–22.10. Haunerland NH, Shirk PD (1995) Regional and functional differentiation in the insect

fat body. Annu Rev Entomol 40:121–145.11. Colombani J, et al. (2003) A nutrient sensor mechanism controls Drosophila growth.

Cell 114:739–749.12. Yamashita O (1996) Diapause hormone of the silkworm, Bombyx mori: Structure,

gene expression and function. J Insect Physiol 42:669–679.13. Phillips JR, Newsom LD (1966) Diapause in Heliothis zea and Heliothis virescens

(Lepidoptera: Noctuidae). Ann Entomol Soc Am 59:154–159.14. Géminard C, Rulifson EJ, Léopold P (2009) Remote control of insulin secretion by fat

cells in Drosophila. Cell Metab 10:199–207.

15. Sousa-Nunes R, Yee LL, Gould AP (2011) Fat cells reactivate quiescent neuroblasts viaTOR and glial insulin relays in Drosophila. Nature 471:508–512.

16. Xu WH, Denlinger DL (2003) Molecular characterization of prothoracicotropic hor-mone and diapause hormone in Heliothis virescens during diapause, and a new rolefor diapause hormone. Insect Mol Biol 12:509–516.

17. Zhang TY, et al. (2004) The diapause hormone-pheromone biosynthesis activatingneuropeptide gene of Helicoverpa armigera encodes multiple peptides that break,rather than induce, diapause. J Insect Physiol 50:547–554.

18. Hahn DA, Denlinger DL (2011) Energetics of insect diapause. Annu Rev Entomol 56:103–121.

19. O’Riordan VB, Burnell AM (1989) Intermediary metabolism in the dauer larva ofthe nematode Caenorhabditis elegans-1. Glycolysis, gluconeogenesis, oxidativephosphorylation and the tricarboxylic acid cycle. Comp Biochem Physiol B 92:233–238.

20. Dark J, Miller DR, Licht P, Zucker I (1996) Glucoprivation counteracts effects of tes-tosterone on daily torpor in Siberian hamsters. Am J Physiol 270:R398–R403.

21. Yang J, Zhu J, Xu WH (2010) Differential expression, phosphorylation of COX subunit1 and COX activity during diapause phase in the cotton bollworm, Helicoverpa ar-migera. J Insect Physiol 56:1992–1998.

22. Zhang Q, Lu YX, Xu WH (2012) Integrated proteomic and metabolomic analysis oflarval brain associated with diapause induction and preparation in the cotton boll-worm, Helicoverpa armigera. J Proteome Res 11:1042–1053.

14692 | www.pnas.org/cgi/doi/10.1073/pnas.1212879109 Xu et al.

Dow

nloa

ded

by g

uest

on

June

5, 2

020