Differential Effects of Tissue-Specific Deletion of BOSS on Feeding Behaviors and Energy Metabolism

Abstract

Food intake and energy metabolism are tightly controlled to maintain stable energy homeostasis and healthy states. Thus, animals detect their stored energy levels, and based on this, they determine appropriate food intake and meal size. Drosophila melanogaster putative G protein-coupled receptor, Bride of sevenless (BOSS) is a highly evolutionarily conserved protein that responds to extracellular glucose levels in order to regulate energy homeostasis. To address how BOSS regulates energy homeostasis, we characterized a boss mutant by assessing its food intake and stored energy levels. Boss mutants exhibited increased food intake but decreased stored triacylglyceride levels. Using boss-GAL4 drivers, we found that boss is expressed in select tissues that are involved in nutrient sensing and food intake, in a subset of neurons in brain and chemosensory organs, in fat body, and in endocrine cells in gut (enteroendocrine cells). Flies with tissue-specific boss knockdowns in these tissues had abnormal stored energy levels and abnormal food intake. These results suggest that BOSS in either neurons or peripheral nutrient-sensing tissues affects energy homeostasis in ways that relate to the sensing of nutrients and regulation of food intake.

About the Authors:

Ayako Kohyama-Koganeya

Affiliation: Molecular Membrane Neuroscience, Brain Science Institute, RIKEN, Wako-shi, Saitama, Japan

Mizuki Kurosawa

Affiliation: Molecular Membrane Neuroscience, Brain Science Institute, RIKEN, Wako-shi, Saitama, Japan

Yoshio Hirabayashi

* E-mail: [email protected]

Affiliation: Molecular Membrane Neuroscience, Brain Science Institute, RIKEN, Wako-shi, Saitama, Japan

Introduction

To maintain energy homeostasis, energy sensing tissues, such as brain, gut, and adipose tissue, translate changes in internal metabolic state into alterations in feeding behavior [1]. Understanding how these tissues communicate with each other is fundamental for understanding state-dependent modifications of behavior and also for elucidating the relevance of this communication to human obesity and associated diabetes and cardiovascular disease. Drosophila melanogaster, a widely used genetic model animal, provides a useful model system to study feeding behavior, because many molecules involved in the regulation of energy homeostasis are highly conserved between flies and mammals [2].

Drosophila BOSS is an orphan G protein-coupled receptor (GPCR) membrane protein that was first identified as a ligand for sevenless tyrosine kinase (SEVENLESS), which is involved in eye differentiation in Drosophila [3, 4]. Beside this function, we found that BOSS has an additional critical function: to respond to extracellular glucose levels and regulate energy homeostasis [5]. This function seems to be conserved during evolution, because knockout of the mice homologue GPRC5B protected the mice from obesity resulting from a high-fat diet [6, 7].

We previously reported that energy homeostasis is disrupted in a boss mutant, suggesting that BOSS may play a role in regulating food intake and energy metabolism [5]. Thus, we examined food intake and further metabolic phenotypes of boss mutant flies. We found that stored lipid levels are reduced, but food intake is increased in boss mutant flies. To elucidate how BOSS regulates food intake and energy metabolism, we first analyzed tissue-specific BOSS expression in Drosophila. BOSS is expressed in neurons of chemosensory organs and in the brain, in enteroendocrine cells in the gut, and in the fat body. Then, we generated tissue-specific boss knockdowns and showed that BOSS either in neurons or in the peripheral tissues (the fat body and enteroendocrine cells) regulates energy stores and food intake. Our findings suggest that BOSS is an important regulator of energy homeostasis in nutrient-sensing tissues.

Materials and Methods

Fly stocks and genetics

Boss-GAL4 lines: A 2.1kbp DNA fragment upstream of the putative transcriptional start site of boss [8] was subcloned into a pUAST vector (Drosophila Genomics Resource Center) and microinjected into w embryos (BestGene). Wild-type w1118 wild-type flies were used as controls. The following lines were also used: elav-Gal4, 10xUAS-mCD8GFP, UAS-mCherryNLS (Bloomington Drosophila Stock Center, Indiana University, Bloomington, IN, USA); FB-Gal4 (gift from Dr. R. Kuhnlein, Max Planck Institute for Biophysical Chemistry, Gottingen, Germany); prospero-Gal4 (gift from Dr. G. Suh, Skirball Institute of Biomolecular Medicine, NYU, New York, NY, USA); UAS-boss RNAi (#4365 and #4366, VDRC, Vienna, Austria). To visualize BOSS expression patterns, 10XUAS-mCDGFP/+;Boss-Gal4/+ and 10X UAS-mCD8GFP/Boss-Gal4, UAS-mCherryNLS were produced by crossing 10X UAS-mCD8GFP to Boss-Gal4 or Boss-Gal4, UAS-mCherryNLS. To generate tissue specific boss knockdown flies, adult flies carrying different Gal4 driver (elav-Gal4 for neuronal, prospero-Gal4 for enteroendocrine cells and FB-Gal4 for fat body knockdown) were crossed to boss-RNAi lines [boss-RNAi#1(#4365, on the second chromosome) or boss-RNAi#2 (#4366, on the third chromosome)].

Husbandry

Drosophila melanogaster were grown using standard food comprising 10% glucose, 4% yeast, 4% corn meal, and 1% agar. They were maintained at 25°C, under a 12-hour light and 12-hour dark regime. Flies were collected at the middle of light regime and used for experiments.

Food intake Assays

For the blue-dye food assay, male flies aged 3–5 days were switched from standard food to blue-colored food (10% sucrose, 1% agar, 2% blue no.1 dye) for 1 h. After the feeding, 10 flies were immediately frozen in one tube using liquid nitrogen. Then, flies were homogenized in PBS buffer and centrifuged (13000 rpm) for 15 min. The supernatants were transferred to a new tube, and absorbance was measured at 625 nm.

We used a capillary feeder (CAFE) to measure ingestion by flies. For the CAFE assay, 10 male flies (aged 3–5 days) that were fed on standard food were transferred to an experimental vial containing a glass micropipette and allowed to feed on 10% sucrose solution. The amount consumed, minus evaporation (determined from a vial without flies), was measured. For the post-starvation response of the feeding assay, male flies starved for 18 h or 24 h in vials containing 1% agar were transferred to vials containing blue-colored food or CAFE assay vials.

Metabolic assay for triacylglyceride (TAG) and glycogen

Male flies aged 3–5 days were homogenized in PBS. Supernatant was collected after heat inactivation at 70°C for 5 min and centrifugation at 13000 rpm for 5 min. TAG levels were measured using a TAG assay kit (Sigma, TR0100). Glycogen levels were measured using a glycogen assay kit (Sigma, MAK016). Data were normalized to the total protein measured by a Bradford assay.

Hemolymph sugar measurements

Hemolymph from five groups each comprising 20 female flies aged 3–5 days was used for each condition. Hemolymph was diluted (1:10) in homogenization buffer (137 mM NaCl, 2.7 mM KCl, 5 mM Tris [pH 6.6]) and heated for 5 min at 70°C. Glucose and trehalose levels were measured using a glucose assay kit (Sigma, GAGO-20). To measure trehalose levels in the samples, porcine trehalase (Sigma, T8778) was used to convert trehalose into glucose and quantified with glucose assay kit. The background level of glucose present in the sample was subtracted.

Immunofluorescence

Larvae and flies were dissected in PBS, fixed in PBS containing 4% formaldehyde for 25 min at room temperature, and washed in PBS containing 0.2% Triton X-100 (PBT). Tissues were blocked for 15 min in Blocking One Histo (Nacalai Tesque Inc., Nakagyo-ku, Kyoto, Japan). Primary antibodies were incubated overnight at 4°C, and Alexa Fluor-conjugated secondary antibodies (Life Technologies) were incubated for 2 h at room temperature. Tissues were mounted in Vectashield mounting medium (Vector Laboratories, Inc.). Confocal images were collected using Olympus/FV1000 confocal system.

Antibodies.

We used chick anti-GFP (1:500, Abcam); rat anti-RFP (1:500, [5F8] Chromotek); mouse anti-nc82 (1:20, DSHB); and mouse anti-prospero (1:50, DSHB).

Quantitative real-time PCR (qRT-PCR)

RNA was collected from male flies aged 3–5 days. Samples (10 flies per sample) were immediately frozen in liquid nitrogen. Total RNA was extracted using Trizol Reagent (Invitrogen). Total RNA samples (1 μg per reaction) were reverse transcribed using oligo-dT and random primers and Superscript RT-III (Invitrogen). The generated cDNA was used for real-time RT-PCR (qPCR Mastermix Plus for SYBERGreen, Applied Biosystems). Three separate samples were collected from each condition, and triplicate measurements were conducted. Primers (S1 Table) were designed using the Universal Probe Library Assay Design Center (Roche Applied Science).

Western blotting

Ten male flies aged 3–5 days were used for each condition. The bodies were dissected out in Schneider’s Drosophila medium and homogenized in sample buffer (Nacali Tesque Inc.). For insulin treatment, the bodies were incubated with or without 5 μM recombinant human insulin (Sigma, I2643) for 1 h at 25°C before being homogenized in sample buffer. Rabbit anti-pAKT (1:1000, Cell signaling); rabbit anti-AKT (1:1000, Cell signaling); and mouse anti-tubulin (1:10,000, Sigma) were used for primary antibodies, and HRP-conjugated anti-rabbit or anti-mouse IgG antibodies (Cell Signaling Technology) were used for secondary antibodies. Blots were visualized using the Chemi-Lumi One Super reagent (Nacali Tesque Inc.).

Statistics

For all experiments, error bars represent standard error of the mean (SEM), and p values refer to results of a Student’s test performed in Microsoft Excel (version 14.4.4) (*p<0.05).

Results

Boss mutant exhibits abnormal energy homeostasis

A previous study found that boss mutant larvae exhibit abnormal sugar and lipid metabolism (i.e., elevated circulating sugar and lipid levels and impaired lipid mobilization to oenocytes), and are sensitive to nutrient deprivation stress [5]. Thus, we first measured energy store (TAG and glycogen) levels in adult boss mutant flies. In normal Drosophila, TAG and glycogen constitute the major forms of energy storage for carbohydrates and lipids, respectively. Adult boss mutant flies had reduced TAG levels (Fig 1A), but no difference in glycogen levels (Fig 1B). We also measured hemolymph sugar levels in adult flies. The boss mutant had high trehalose and glucose sugar levels (Fig 1C), levels that are comparable to larval hemolymph sugar levels, which we previously measured [5].

[Figure omitted. See PDF.]

Fig 1. BOSS controls energy metabolism.

(A and B) Relative lipid (A) and glycogen (B) content in WT and boss mutant adult flies (n = 3, 10 flies per replicate). WT, wild type; TAG, triacylglyceride. (C) Circulating sugar levels in the hemolymph of boss mutant adult flies. ACC, acetyl-CoA carboxylase (n = 3, 20 flies per replicate). (D) Amounts of ACC and PEPCK mRNA in boss mutant flies relative to control flies (n = 3, 10 flies per replicate). PEPCK, phosphoenolpyruvate carboxykinase. Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.g001

Acetyl-CoA carboxylase [9] is the rate-limiting enzyme for fatty acid synthesis [9]. We found that ACC expression was reduced in boss mutant flies (Fig 1D). Furthermore, we found that expression of phosphoenolpyruvate carboxykinase (PEPCK), which catalyzes the rate-limiting step of gluconeogenesis and is used as a marker for starvation, was increased in the boss mutant flies (Fig 1D). Overall, these results demonstrate that energy metabolism is disrupted in the boss mutant.

Insulin signaling pathway is downregulated in the boss mutant

Insulin signaling regulates lipid and glucose metabolism in Drosophila similarly to that in vertebrates [10]. In line with the aberrant lipid and glucose metabolism in boss mutant flies, we examined the activity of insulin signaling in these flies. First, we measured phosphorylated AKT (pAKT) levels. Under normal feeding conditions, the boss mutant flies had reduced pAKT levels (Fig 2A). With insulin stimulation, activated pAKT is used as an indication of insulin resistance [11]. Thus, we examined the ability of exogenous insulin to stimulate the phosphorylation of AKT. The boss mutant flies responded to insulin, leading to an increase in pAKT similar to that in control flies (Fig 2A). This ex vivo experiment demonstrates that the insulin-signaling machinery of boss mutant flies is functional but that the insulin-signaling is not activated in vivo. We speculated that Drosophila insulin-like peptides (DILPs) activity might be reduced. In Drosophila, eight DILPs (DILP1-8) are identified, and DILP2, 3, 5 and 6 have been shown to play a physiological role in metabolic control [12]. We, therefore, measured these four dilps’ mRNA expressions by using qRT-PCR. The amounts of mRNA for dilp2 and dilp3 were reduced, but those for dilp5 and dilp6 were increased in the boss mutant flies (S1 Fig). Regulation of dilps expression is very complicated [10, 13, 14]. For example, starvation reduces expression of dilp3 and dilp5, but not dilp2. Moreover, knockdown of dilp2 expression is compensated for by increases in dilp3 and dilp5. Thus, further studies will be required to delineate whether BOSS directly coordinates the regulation of DILPs secretion.

[Figure omitted. See PDF.]

Fig 2. BOSS affects insulin signal activity.

(A) Immunoblot detecting AKT phosphorylation. Without insulin stimulation, pAKT levels are decreased in boss mutant flies. AKT phosphorylation levels in boss mutant flies after insulin stimulation. (B) Amounts of Sdr, dInR and Thor mRNA in boss mutant flies relative to control flies (n = 3, 10 flies per replicate). Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.g002

Recently, secreted decoy of InR (SDR) was identified as a negative regulator of insulin signaling [15]. SDR has similar structure to the extracellular domain of insulin receptor, and antagonizes insulin signaling via this domain. SDR is constantly secreted into the hemolymph and binds to DILPs. In boss mutant flies, Sdr mRNA expression was increased (Fig 2B), indicating that increased Sdr expression is one of the causes of insulin signaling suppression in these flies.

To confirm insulin-signaling activity in the boss mutant flies, we examined the activity of dFOXO. Insulin signaling inhibits FOXO activity [10]. We found that expression of Drosophila Insulin receptor (dInR) and Thor/4E-BP, which are FOXO target genes, were reduced in the boss mutant (Fig 2B). Although it remains to be solved how insulin signaling is downregulated in the boss mutant, these data suggest that the insulin signaling pathway is inactivated in boss mutant flies.

Boss mutant flies exhibit increased food intake

Changes in food intake behavior can affect energy homeostasis in flies. Thus, we examined levels of food intake in the boss mutant by using the CAFE assay (Fig 3A) and blue-dye feeding assay (Fig 3B). Boss mutant flies consumed significantly more food than control flies under free-feeding and starving conditions. These results demonstrate that boss mutant flies display marked increases in food intake (Fig 3).

[Figure omitted. See PDF.]

Fig 3. Boss mutant flies are hyperphagic.

(A) Food consumption measured by the CAFE assay (n = 10, 10 flies per replicate). (B) Food consumption measured by Blue-dye ingestion assay. The ingestion of dye was quantified after feeding male flies for 1h in the light cycle, and then by measuring the absorbance of ingested dye (n = 10, 10 flies per replicate). Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.g003

BOSS is expressed in nutrient-sensing tissues

Food intake is a necessary behavior that provides energy to survive, and this depends on body energy balance. Tissues communicate with each other and strive to maintain this balance [2, 16]. To elucidate the mechanisms of BOSS signaling, it is important to closely examine BOSS-expressing tissues. Thus, we analyzed boss expression by various tissues using a boss-GAL4 driver. We generated boss-GAL4 transgenic flies that contain 2.1kb upstream of the predicted transcription start site of the boss gene [8], and characterized the expression pattern of this Gal4 driver using UAS-mCD8GFP and UAS-mCherryNLS.

Live fluorescence imaging and immunohistochemistry showed that GFP and mCherry were expressed in tissues known to be involved in nutrient sensing and food intake (Fig 4). For example, GFP and mCherry expression were detected in chemosensory neurons of five specific chemosensory organs: the dorsal and ventral pharyngeal organ, the ventral pits of larvae, the antennae, the proboscis, and the legs of adults (Fig 4A–4C). Second, GFP and mCherry expression were detected in specific neurons of the central nervous system (CNS) (Fig 4D–4F). Third, GFP expression was detected in peripheral tissues, specifically, subsets of cells in the gut and the fat body (Fig 4G–4J). Most of these GFP-labeled cells in the midgut were co-labeled with the enteroendocrine cell marker prospero (93% in larva, and 98% in adult) (Fig 4G and 4H, S2 Fig), suggesting that boss-GAL4 expressing cells in the gut are subsets of enteroendocrine cells.

[Figure omitted. See PDF.]

Fig 4. BOSS is expressed in nutrient-sensing tissues.

Expression of boss in larval and adult flies was visualized by expression of 10xUAS-mCD8.GFP and UAS-mCherryNLS using a boss-GAL4 driver. (A-C) Expression in larval (A) and adult (B) sensory organs. (C) Adult foreleg. (D) Larval CNS. (E) Adult brain. (F) Adult thoracic ganglion. To identify the locations of boss-expressing cells, UAS-mCherryNLS (magenta) was coexpressed with 10xUAS-mCD8.GFP [9] in the brain and sensory organs. The CNS was also labeled with antibody nc82 (blue), a marker that labels synapses. Larval (G) and adult (H) midgut. The gut was counterstained with DAPI (blue) and a marker for enteroendocrine cells, prospero (magenta). Arrowheads show the prospero-positive GFP-expressing cells. See also S2 Fig. Larval (I) and adult (J) fat body. The fat body was counterstained with DAPI (blue), a cell nucleus marker. Scale bars; 500 μm in A, 30 μm in B, 200 μm in C and E, 100 μm in G, H and J, 80 μm in D, 50 μm in H.

https://doi.org/10.1371/journal.pone.0133083.g004

CNS and enteroendocrine cells express and secrete hormones and neuropeptides, some of which are involved in energy metabolism [17] [18] [19]. Chemosensory organs detect nutrients and send signals to other tissues to coordinate food intake [20]. The fat body of flies, which is similar in function to mammalian liver and white adipose tissue, stores and releases energy in response to energy demands [21]. The BOSS expression profile we observed, therefore, indicates that BOSS coordinates inter-organ/inter-tissue communication to regulate food intake and energy metabolism. Therefore, we next examined the function of BOSS in these tissues.

Tissue-specific boss knockdown flies exhibit contrasting food intake and metabolic phenotypes

To determine the function of BOSS in the tissues, neurons, enteroendocrine cells and fat bodies, we generated tissue-specific boss knockdown flies (S3 Fig) and measured their food intake under free-feeding and starved states. Neuronal-specific knockdown flies ingested decreased amounts of food (Fig 5A). One line of enteroendocrine cells-specific knockdown flies(RNAi #2)also showed reduced food ingestion, but the other line(RNAi #1)showed no difference (Fig 5B). We speculate this discrepancy may occur by positional effects of RNAi lines, since they were generated using P-element random insertion method [22]. Thus, there is a possibility that food intake phenotype was diminished by positional effects in RNAi #1 line. We need further experiments to confirm that boss expressed in enteroendocrine cells affects food intake regulation. In contrast, fat body-specific knockdown flies significantly and selectively increased their food ingestion in both states (Fig 5C).

[Figure omitted. See PDF.]

Fig 5. Effect of tissue-specific expression of boss RNAi on food intake.

Flies expressing boss RNAi in neurons from the elav-GAL4 (A), enteroendocrine cells from proepero-GAL4 (B), and fat body from FB-GAL4 (C) were used for blue-dye ingestion assay. The ingestion of dye was quantified after feeding male flies for 1 h, and thereafter by measuring the absorbance of ingested dye. Two transgene insertions were used: boss-RNAi #1 and #2 (n = 10, 10 flies per replicate). Assays were performed under two conditions [no starvation (upper row) and 24h starvation (lower row)]. Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.g005

Next, we examined stored TAG and glycogen levels. Neuronal-specific knockdown flies had reduced stored TAG levels (Fig 6A and 6B). Enteroendocrine cells-specific knockdown flies had reduced stored TAG levels but increased stored glycogen levels (Fig 6C and 6D). On the other hand, fat body-specific knockdown flies had significantly increased stored TAG levels but not glycogen levels (Fig 6E and 6F). These results indicate that peripheral and neuronal BOSS is differentially involved in the regulation of energy stores.

[Figure omitted. See PDF.]

Fig 6. Effect of tissue-specific expression of boss RNAi on lipid and glycogen levels.

TAG (upper row) and glycogen (lower row) levels were normalized to protein levels and expressed as ratios to the control. The flies’ genotypes are the same as those flies used in the blue-dye ingestion assay.

https://doi.org/10.1371/journal.pone.0133083.g006

Discussion

The question of how animals control their feeding and energy metabolism and maintain energy homeostasis is one of the most important issues we currently need to understand. To approach this question, it is necessary to identify the molecule(s) that control and coordinate feeding and energy metabolism. Here, we present evidence that BOSS regulates food intake and energy metabolism. Without BOSS, food intake and energy storage levels in the body are altered, and energy homeostasis is disrupted. Furthermore, tissue-specific boss knockdown mutants flies exhibited the opposite food intake and metabolic phenotypes, indicating that BOSS coordinates the interaction between metabolic tissues for maintaining energy homeostasis (S4 Fig).

BOSS contributes to energy homeostasis in tissues where it is expressed

We have shown that a boss null mutant and tissue-specific boss knockdown flies show aberrant energy metabolism (Figs 1 and 2) and food intake (Fig 3). One of the more important findings is that the phenotype of the fat-body-specific boss knockdown flies is opposite to that of the neuronal-specific or enteroendocrine-cell-specific boss knockdown flies. The fat-body-specific boss knockdown flies increased stored lipids and food intake. In contrast, the neuronal-specific and enteroendocrine-cell-specific boss knockdown flies reduced their stored lipids and food intake.

At present, few reports have shown that the input of a gene or signaling pathway from some tissue is counteracted by the input of that from another tissue. Cyclic AMP responsive element binding protein (CREB) is an evolutionarily conserved transcription factor involved in many important physiological functions [23]. CREB activity either in neurons or in the fat body controls energy stores and feeding behavior, but the effects are opposite [24]. Another example of this counteraction scenario is found in circadian clock genes. Flies lacking circadian clock (clock knockdown or dominant negative form of cycle overexpression) in the fat body display increased food intake but decreased stored glycogen and TAG. On the other hand, flies lacking circadian clocks in neurons display decreased food intake but increased stored glycogen and TAG [25]. Thus, it is an especially interesting issue to investigate the correlation between BOSS and CREB or circadian clocks in the future.

BOSS is expressed in nutrient-sensing tissues

There is growing evidence that physiological communication between tissues coordinate food intake and utilization of nutrient stores with energy requirements [16]. For example, the brain functions as a key integrator of energy homeostasis that occurs in other tissues, such as in the fat body and in muscle [16, 26]. How do these tissues communicate with each other?

One of the major systems involved in the communication among tissues is the endocrine signaling network [2]. Hormones and neuropeptides—many of which are conserved between Drosophila and mammals—are secreted and used for inter-organ and inter-tissue communication [16]. Certain tissues are known to secret endocrine signals and transmit information about food or body energy condition. In the gut, enteroendocrine cells that are chemosensory cells, for example, secrete peptides in response to the nutritional status of the body [27]. In mammals and Drosophila, taste receptors are expressed in enteroendocrine cells, implying that these taste receptors have chemosensory roles in the regulation of physiological functions such as food intake, nutrient absorption, and sugar homeostasis [17, 28].

The fat body functions not only as an energy store organ but also as an endocrine organ. The fat body secrets TAG under starving conditions and functions to maintain energy homeostasis [29]. Recently, Upd2 was identified as a leptin homologue secreted from the fat body [30]. In Drosophila, many peptides that regulate feeding and energy metabolism have been identified, and many endocrine signals (hormones and neuropeptides) are conserved between Drosophila and mammals [31].

In this study, we demonstrated that neuronally expressed BOSS affects food intake and energy metabolism. In addition, we showed that BOSS is expressed in neuronal and peripheral tissues that also express neuropeptides and hormones (Fig 4). Thus, it will be of great interest to investigate in the future how BOSS regulates endocrine signals and coordinate energy metabolism.

We also found that BOSS is expressed in both gustatory and olfactory organs (Fig 4A–4C). Feeding behavior is determined by taste and other sensory information, such as smell, touch, vision, and ambient temperature. One report describes how odor affects taste behavior through the maxillary palp [32]. Thus, in this context, further detailed analysis of neuronal BOSS’s function will reveal more precise physiological roles of neuronal BOSS. It will be interesting to examine whether and how BOSS expressed in the olfactory organs, the antenna, and the maxillary palp modulates feeding behavior.

BOSS is an evolutionarily conserved protein [33]. The expression of GPRC5B, a mammalian BOSS homologue, is similar to that of BOSS [6, 7, 34]. GPRC5B is broadly expressed in brain, gut, and adipose tissues. Recently, aberrant expression of GPRC5B was identified as an obesity risk factor [9]. The metabolism of Drosophila and mammals shows similarities with evolutionarily conserved signaling pathways and analogous tissues that regulate energy metabolism [35]. Thus, specific analysis of Drosophila BOSS will open the door for understanding the broader physiological functions of the highly conserved membrane protein BOSS/GPRC5B.

Supporting Information

[Figure omitted. See PDF.]

S1 Fig. Expression of Drosophila insulin-like peptides (dilps) in boss mutant flies.

Expression of Drosophila insulins (dilp2, 3, 5, and 6) were quantified by qRT-PCR (n = 3, 10 flies per replicate). Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.s001

(TIFF)

S2 Fig. BOSS-expressing cells in the gut.

(A) Expression of boss in the gut was visualized by expression of 10xUAS-mCD8.GFP using boss-GAL4 driver. To mark enteroendocrine cells, the antibody against prospero was used. Arrowheads show the prospero-positive GFP-expressing cells. Scale bars; 100 μm. (B) Total number of GFP (BOSS)- and prospero-positive cells in the larval and adult midgut. Cells were counted from immunofluoresence images (n = 3 each)

https://doi.org/10.1371/journal.pone.0133083.s002

(TIFF)

S3 Fig. BOSS expression in tissue-specific boss knockdown (KD) flies.

To confirm that boss knockdowns were effective, expression of boss mRNA was quantified by qRT-PCR (n = 3, 10 flies per replicate). (A) Neuronal-specific boss KD. (B) Enteroendocrine cells-specific boss KD. (C) Fat body-specific boss KD. To generate tissue specific boss KD flies, adult flies carrying different Gal4 driver [elav-Gal4 for neuronal KD, prospero-Gla4 for enteroendocrine cells KD and FB-Gal4 for fat body KD] were crossed to boss-RNAi lines [boss-RNAi#1(v4365) or boss-RNAi#2 (v4366)]. mRNA were collected from head for neuronal KD, whole gut for enteroendocrine cells KD, and body for fat body KD, since isolation of these tissues alone from whole flies was technically difficult. Data are represented as mean ±SEM (*P<0.05).

https://doi.org/10.1371/journal.pone.0133083.s003

(TIFF)

S4 Fig. BOSS expression in tissue-specific boss knockdown flies.

(A) Summary of BOSS-expressing organs/tissues in larva and adult. (B) Schematic model proposed to explain the roles of BOSS in regulating food intake and energy storage. BOSS-expressing tissues sense nutrition (glucose) in food or hemolymph, induce hormones and neuropeptides for coordinating inter-organ communication (arrows), and maintain energy homeostasis. BOSS expression in neurons and gut promotes food intake and energy storage, but BOSS expression in fat body suppresses food intake and energy storage.

https://doi.org/10.1371/journal.pone.0133083.s004

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S1 Table. Primer sequences for qRT-PCR.

https://doi.org/10.1371/journal.pone.0133083.s005

(TIFF)

Acknowledgments

We would like to thank the Bloomington Drosophila Stock Center (Indiana University), the Vienna RNAi Center (VDRC), the Developmental Studies Hybridoma Bank (University of Iowa) and FLYBASE for providing fly lines, antibodies and information.

This work was supported in part by grants from the RIKEN Brain Science Institute (to Y. H.) and by 2010 General Research Grant from Nestle Nutrition Council, Japan (to AK). We thank to T. Simizu and E. Oshima for technical assistance. We are grateful to the Support Unit for Bio-material Analysis, RIKEN Brain Science Institute Research Resources Center, for help with the nucleotide sequencing analysis.

Author Contributions

Conceived and designed the experiments: AK YH. Performed the experiments: AK MK. Analyzed the data: AK MK. Wrote the paper: AK YH.

Citation: Kohyama-Koganeya A, Kurosawa M, Hirabayashi Y (2015) Differential Effects of Tissue-Specific Deletion of BOSS on Feeding Behaviors and Energy Metabolism. PLoS ONE 10(7): e0133083. https://doi.org/10.1371/journal.pone.0133083

References

1. Morton GJ, Meek TH, Schwartz MW. Neurobiology of food intake in health and disease. Nat Rev Neurosci. 2014;15(6):367–78. pmid:24840801; PubMed Central PMCID: PMC4076116.

2. Padmanabha D, Baker KD. Drosophila gains traction as a repurposed tool to investigate metabolism. Trends Endocrinol Metab. 2014. pmid:24768030.

3. Hart AC, Kramer H, Van Vactor DL Jr., Paidhungat M, Zipursky SL. Induction of cell fate in the Drosophila retina: the bride of sevenless protein is predicted to contain a large extracellular domain and seven transmembrane segments. Genes Dev. 1990;4(11):1835–47. pmid:2276620.

4. Kramer H, Cagan RL, Zipursky SL. Interaction of bride of sevenless membrane-bound ligand and the sevenless tyrosine-kinase receptor. Nature. 1991;352(6332):207–12. pmid:1857416.

5. Kohyama-Koganeya A, Kim YJ, Miura M, Hirabayashi Y. A Drosophila orphan G protein-coupled receptor BOSS functions as a glucose-responding receptor: loss of boss causes abnormal energy metabolism. Proc Natl Acad Sci U S A. 2008;105(40):15328–33. pmid:18832180; PubMed Central PMCID: PMC2563099.

6. Kim YJ, Sano T, Nabetani T, Asano Y, Hirabayashi Y. GPRC5B activates obesity-associated inflammatory signaling in adipocytes. Sci Signal. 2012;5(251):ra85. pmid:23169819.

7. Sano T, Kim YJ, Oshima E, Shimizu C, Kiyonari H, Abe T, et al. Comparative characterization of GPRC5B and GPRC5CLacZ knockin mice; behavioral abnormalities in GPRC5B-deficient mice. Biochem Biophys Res Commun. 2011;412(3):460–5. pmid:21840300.

8. Sunio A, Metcalf AB, Kramer H. Genetic dissection of endocytic trafficking in Drosophila using a horseradish peroxidase-bride of sevenless chimera: hook is required for normal maturation of multivesicular endosomes. Mol Biol Cell. 1999;10(4):847–59. pmid:10198042; PubMed Central PMCID: PMC25206.

9. Speliotes EK, Willer CJ, Berndt SI, Monda KL, Thorleifsson G, Jackson AU, et al. Association analyses of 249,796 individuals reveal 18 new loci associated with body mass index. Nat Genet. 2010;42(11):937–48. pmid:20935630; PubMed Central PMCID: PMC3014648.

10. Teleman AA. Molecular mechanisms of metabolic regulation by insulin in Drosophila. Biochem J. 2010;425(1):13–26. pmid:20001959.

11. Musselman LP, Fink JL, Narzinski K, Ramachandran PV, Hathiramani SS, Cagan RL, et al. A high-sugar diet produces obesity and insulin resistance in wild-type Drosophila. Dis Model Mech. 2011;4(6):842–9. pmid:21719444; PubMed Central PMCID: PMC3209653.

12. Kannan K, Fridell YW. Functional implications of Drosophila insulin-like peptides in metabolism, aging, and dietary restriction. Frontiers in physiology. 2013;4:288. pmid:24137131; PubMed Central PMCID: PMC3797364.

13. Gronke S, Clarke DF, Broughton S, Andrews TD, Partridge L. Molecular evolution and functional characterization of Drosophila insulin-like peptides. PLoS Genet. 2010;6(2):e1000857. pmid:20195512; PubMed Central PMCID: PMC2829060.

14. Nassel DR, Kubrak OI, Liu Y, Luo J, Lushchak OV. Factors that regulate insulin producing cells and their output in Drosophila. Front Physiol. 2013;4:252. pmid:24062693; PubMed Central PMCID: PMC3775311.

15. Okamoto N, Nakamori R, Murai T, Yamauchi Y, Masuda A, Nishimura T. A secreted decoy of InR antagonizes insulin/IGF signaling to restrict body growth in Drosophila. Genes Dev. 2013;27(1):87–97. pmid:23307869; PubMed Central PMCID: PMC3553286.

16. Rajan A, Perrimon N. Drosophila as a model for interorgan communication: lessons from studies on energy homeostasis. Dev Cell. 2011;21(1):29–31. pmid:21763605; PubMed Central PMCID: PMC3560414.

17. Park JH, Kwon JY. Heterogeneous expression of Drosophila gustatory receptors in enteroendocrine cells. PLoS One. 2011;6(12):e29022. pmid:22194978; PubMed Central PMCID: PMC3237578.

18. Li S, Torre-Muruzabal T, Sogaard KC, Ren GR, Hauser F, Engelsen SM, et al. Expression patterns of the Drosophila neuropeptide CCHamide-2 and its receptor may suggest hormonal signaling from the gut to the brain. PLoS One. 2013;8(10):e76131. pmid:24098432; PubMed Central PMCID: PMC3788761.

19. Reiher W, Shirras C, Kahnt J, Baumeister S, Isaac RE, Wegener C. Peptidomics and peptide hormone processing in the Drosophila midgut. J Proteome Res. 2011;10(4):1881–92. pmid:21214272.

20. Melcher C, Pankratz MJ. Candidate gustatory interneurons modulating feeding behavior in the Drosophila brain. PLoS Biol. 2005;3(9):e305. pmid:16122349; PubMed Central PMCID: PMC1193519.

21. Arrese EL, Soulages JL. Insect fat body: energy, metabolism, and regulation. Annu Rev Entomol. 2010;55:207–25. pmid:19725772; PubMed Central PMCID: PMC3075550.

22. Dietzl G, Chen D, Schnorrer F, Su KC, Barinova Y, Fellner M, et al. A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila. Nature. 2007;448(7150):151–6. pmid:17625558.

23. Altarejos JY, Montminy M. CREB and the CRTC co-activators: sensors for hormonal and metabolic signals. Nat Rev Mol Cell Biol. 2011;12(3):141–51. pmid:21346730.

24. Iijima K, Zhao L, Shenton C, Iijima-Ando K. Regulation of energy stores and feeding by neuronal and peripheral CREB activity in Drosophila. PLoS One. 2009;4(12):e8498. pmid:20041126; PubMed Central PMCID: PMC2795867.

25. Xu K, Zheng X, Sehgal A. Regulation of feeding and metabolism by neuronal and peripheral clocks in Drosophila. Cell Metab. 2008;8(4):289–300. pmid:18840359; PubMed Central PMCID: PMC2703740.

26. Badisco L, Van Wielendaele P, Vanden Broeck J. Eat to reproduce: a key role for the insulin signaling pathway in adult insects. Frontiers in physiology. 2013;4:202. pmid:23966944; PubMed Central PMCID: PMC3735985.

27. Janssen S, Depoortere I. Nutrient sensing in the gut: new roads to therapeutics? Trends Endocrinol Metab. 2013;24(2):92–100. pmid:23266105.

28. Jang HJ, Kokrashvili Z, Theodorakis MJ, Carlson OD, Kim BJ, Zhou J, et al. Gut-expressed gustducin and taste receptors regulate secretion of glucagon-like peptide-1. Proc Natl Acad Sci U S A. 2007;104(38):15069–74. pmid:17724330; PubMed Central PMCID: PMC1986614.

29. Billeter JC, Atallah J, Krupp JJ, Millar JG, Levine JD. Specialized cells tag sexual and species identity in Drosophila melanogaster. Nature. 2009;461(7266):987–91. pmid:19829381.

30. Rajan A, Perrimon N. Drosophila cytokine unpaired 2 regulates physiological homeostasis by remotely controlling insulin secretion. Cell. 2012;151(1):123–37. pmid:23021220; PubMed Central PMCID: PMC3475207.

31. Audsley N, Weaver RJ. Neuropeptides associated with the regulation of feeding in insects. Gen Comp Endocrinol. 2009;162(1):93–104. pmid:18775723.

32. Shiraiwa T. Multimodal chemosensory integration through the maxillary palp in Drosophila. PLoS One. 2008;3(5):e2191. pmid:18478104; PubMed Central PMCID: PMC2364657.

33. Metpally RP, Sowdhamini R. Cross genome phylogenetic analysis of human and Drosophila G protein-coupled receptors: application to functional annotation of orphan receptors. BMC Genomics. 2005;6:106. pmid:16091152; PubMed Central PMCID: PMC1192796.

34. Robbins MJ, Michalovich D, Hill J, Calver AR, Medhurst AD, Gloger I, et al. Molecular cloning and characterization of two novel retinoic acid-inducible orphan G-protein-coupled receptors (GPRC5B and GPRC5C). Genomics. 2000;67(1):8–18. pmid:10945465.

35. Smith WW, Thomas J, Liu J, Li T, Moran TH. From fat fruit fly to human obesity. Physiol Behav. 2014. pmid:24508822.