Abstract
In Drosophila larvae, growth and developmental timing are regulated by nutrition in a tightly coordinated fashion. The networks that couple these processes are far from understood. Here, we show that the intestine responds to nutrient availability by regulating production of a circulating lipoprotein-associated form of the signaling protein Hedgehog (Hh). Levels of circulating Hh tune the rates of growth and developmental timing in a coordinated fashion. Circulating Hh signals to the fat body to control larval growth.
It regulates developmental timing by controlling ecdysteroid production in the prothoracic gland. Circulating Hh is especially important during starvation, when it is also required for mobilization of fat body triacylglycerol (TAG) stores. Thus, we demonstrate that Hh, previously known only for its local morphogenetic functions, also acts as a lipoprotein-associated endocrine hormone, coordinating the response of multiple tissues to nutrient availability.
Introduction
Organismal growth and development are regulated by the availability of nutrients. The reaction of a developing animal to nutrient deprivation involves a complex inter-organ response orchestrated by the nervous, endocrine and digestive systems, with the goal to allow basic cellular metabolism but to delay energy demanding processes. A major physiological response to starvation is the utilization of stored energy and the delay of growth and development.
The Drosophila fat body, a tissue analogous to mammalian white adipose tissue and liver, has an important role in coupling nutrient availability to growth. In response to changes in the availability of sugars, amino acids and lipids, it releases secreted factors that regulate growth by modulating systemic insulin signaling. (Arquier et al. 2008; Rajan and Perrimon 2012; Geminard et al. 2009; and reviewed in Hietakangas and Cohen 2009). It also acts as a depot for these nutrients, and can release them in response to starvation (reviewed in Telfer and Kunkel 1991; Arrese and Soulages 2010). The fat body directly senses circulating amino acid levels using the amino acid transporter Slimfast and the Target of Rapamycin (TOR) pathway (Colombani et al. 2003). However, how it senses the availability of other types of nutrients is unknown.
In addition to regulating growth, nutrient availability also controls developmental progression by influencing production of ecdysteroids in the prothoracic gland. During larval development, ecdysteroids are produced in pulses that regulate molting and initiate pupariation (reviewed in Thummel 1996). These pulses are tightly coupled to organismal growth. A key developmental decision occurs once larvae achieve a “critical weight” (reviewed in Mirth and Riddiford 2007). After this point, starvation no longer delays pupariation, although it affects the final size of resulting adults. Critical weight is associated with a small increase in ecdysteroid levels, which later rise dramatically to initiate pupariation (Warren et al. 2006). The commitment to and timing of pupariation are subject to multiple checkpoints and can be affected not only by nutrition, but also damage to imaginal tissues, circadian signals, and temperature (reviewed in Shingleton and Mirth 2012). Consistent with this, multiple signaling pathways in the prothoracic gland influence the decision to up-regulate ecdysteroid production (Mirth et al. 2005; Colombani et al. 2005; McBrayer et al. 2007; Rewitz et al. 2009; Gibbens et al. 2011; Cáceres et al. 2011; Ou et al. 2011) How these networks interact to couple ecdysteroids production to the size of the animal is far from understood.
In other organisms, the digestive system is an important node in the signaling network that controls metabolism. Signals originating from the stomach and the intestine activate neural and humoral pathways to regulate food intake and metabolic homeostasis (reviewed in Cummings and Overduin 2007). Less is known about signals produced by the Drosophila gastrointestinal tract, although it harbors neuroendocrine cells that can synthesize peptide hormones, and is connected to the nervous system (Cognigni et al. 2011; Miguel-Aliaga 2012; Lemaitre and Miguel-Aliaga 2013). The digestive system is well-placed to directly sense nutrient availability as opposed to circulating nutrients. However, whether the Drosophila gastrointestinal tract conveys this type of nutritional information is not known.
Hedgehog (Hh) signaling is known to regulate metabolic processes such as fat storage and glucose uptake in mice (Suh et al. 2006; Pospisilik et al. 2010; Teperino et al. 2012). But the source of the relevant Hh ligand has never been identified. Interestingly, we have shown that circulating lipoproteins in flies and mammals carry not only lipid cargo, but also the lipid-modified signaling molecule Hedgehog (Palm et al. 2013). The tissue source and function of circulating Hh is unknown. In this study, we investigate the source and the function of lipoprotein-associated systemically circulating Hh in Drosophila.
Results
Midgut derived Hh circulates systemically
To investigate the source of circulating Hh, we characterized the tissue expression patterns of a variety of different Gal4 drivers, and used them to knock down Hh production (suppl. Figure 1A). We then measured Hh levels in the larval hemolymph. Ubiquitous induction of hhRNAi using tubulin-Gal4 efficiently reduces the level of circulating Hh (suppl. Figure 1B). We next used en-GAL4, en105-Gal4 or hh-Gal4 to target hhRNAi to posterior compartments of larval ectoderm and imaginal discs, which express hh. These drivers are also active in the salivary glands. Knock-down using these drivers had no effect on the level of circulating Hh, despite strongly reducing hh expression in imaginal discs. (Figure 1A, and suppl. Figure 1C). Thus, circulating Hh does not derive from the imaginal discs, larval ectoderm or salivary gland. Therefore, we used immunofluorescence to identify other tissues that might express hh. We noted that Hh protein is present in the larval midgut and its attached gastric caeca (Figure 1B). In the midgut, Hh localizes to the enterocytes, which function in nutrient absorption (Figure 1C). To ask whether this region of the gut also transcribes hh, we performed in-situ hybridization and RT-PCR on dissected midgut samples. These experiments showed that hh mRNA is present in the midgut, and that it localizes to enterocytes but not gastric caeca (Figure 1D,E). Thus, Drosophila midgut enterocytes transcribe and translate Hh. The Hh protein found in gastric ceaca must be derived from other sources.
To ask whether circulating Hh originates from the larval midgut, we used the myo1A-Gal4 driver. This driver is active in the midgut enterocytes and the salivary gland, and is also expressed in a few cells in the fat body, CNS, and disc peripodial membrane (suppl. Figure 1A). Driving hhRNAi with myo1A-Gal4 reduces the amount of Hh in the larval hemolymph, although it is not completely eliminated (Figure 1A). To ask whether the activity of myo1A-Gal4 in fat body or CNS underlies its ability to reduce Hh levels in the hemolymph, we induced hhRNAi in these tissues more completely using lpp-Gal4 or elav-Gal4 (which drive strong expression in the fat body and CNS, respectively). Neither reduced systemic Hh levels when used to drive hhRNAi (Figure 1A). These results strongly suggest that midgut enterocytes are a major source of systemic Hh, although minor amounts may come from other tissues.
Circulating Hh inhibits systemic growth and slows developmental timing
Although larvae lacking circulating Hh can give rise to normally patterned adults, these animals are somewhat smaller than controls. We therefore wondered whether circulating Hh might control growth or developmental timing. To address this, we manipulated Hh production in the midgut and quantified the larval growth rate, and the timing of pupariation. Specific RNAi-mediated knock-down of Hh production in the midgut, which reduces but does not eliminate circulating Hh, causes earlier pupariation but doesn’t significantly affect the larval growth rate (Figure 2A). The resulting adults and pupae are smaller because they grow at a similar rate but pupariate too early. To ask whether a complete lack of circulating Hh would cause a stronger growth phenotype, we examined larvae mutant for the temperature sensitive allele hhts2. When these larvae are grown at the restrictive temperature, they pupariate earlier than control larvae and also grow faster (Figure 2B), giving rise to pupae of normal size.
To study the effects of raising circulating Hh levels, we expressed hh under the control of two different Gal4 drivers: myo1A-Gal4 and npc1b-Gal4. Npc1b-Gal4-driven expression is strictly midgut specific and strongly increases the level of circulating Hh. Myo1A-Gal4 is expressed more highly than npc1b-Gal4 in the midgut, but is also active a few other tissues. Myo1A-Gal4 driven hh expression increases circulating Hh levels more dramatically than npc1b-Gal4 (Palm et al. 2013). Pupariation of npc1b>hh larvae is delayed by 20 hours, compared to control npc1b/+ larvae (Figure 2C). Myo1A-driven hh has a stronger effect, delaying pupariation by 4 days compared to controls (Figure 2C). Delayed pupariation in hh over-expressing animals is accompanied by a proportional reduction in larval growth, such that these animals pupariate at close to a normal size (Figure 2C). Taken together, these data show that the amount of Hh circulating in the hemolymph controls both the larval growth rate and the speed of pupariation in a proportional fashion.
Circulating Hh promotes survival under starvation
Growth and developmental timing are modulated in response to nutrient availability. Circulating Hh is produced in the tissue where nutrient absorption occurs. Therefore, we investigated whether its production responded to nutrient uptake. Quantitative PCR and Western blot analysis show that midgut hh expression is significantly elevated upon starvation in both pre and post-critical weight larvae (Figure 3A,B). To distinguish the effects of protein, sugar and lipids on hh expression, we asked which nutrients could reverse the upregulation of hh expression in starving larvae. Adding 5% sugar (fructose or glucose) to starvation medium (PBS/Agar) did not reverse hh upregulation (Figure 3C). Adding 10% chloroform-extracted yeast autolysate as a source of amino acids, and other water-soluble molecules, partially reversed hh upregulation (Figure 3C). However, adding both sugar and protein consistently and completely reversed Hh upregulation in the midgut (Figure 3C). These data suggest that midgut cells deprived either of sugar or of amino acids increase hh expression. Interestingly, dietary lipids do not appear to be required to keep hh expression low in the midgut.
Control larvae can survive for 48 hours when they are starved before reaching critical weight (Figure 3D). To ask whether Hh production by the midgut was important for survival under starvation conditions, we reduced Hh function in larvae starved before critical weight by driving hhRNAi in the midgut with myo1A-Gal4. Although myo1A-driven hhRNAi does not reduce survival of normally fed animals, it reduces survival on starvation to only 24 hours (Figure 3D, and suppl. Fig 2A). Thus, even the incomplete reduction of circulating Hh caused by myo1A -induced hhRNAi strongly decreases survival upon starvation. This suggests that circulating Hh promotes survival under starvation conditions.
To reduce systemic Hh levels more completely, we blocked Hh function using the temperature sensitive Hh allele hhts2. Hhts2 larvae cultured at the restrictive temperature (29°C) from hatching onwards complete larval development with no increased lethality when fed normally, but die during early pupal stages (suppl. Fig 2B). Thus, Hh function is not required for larval survival or pupariation of normally fed animals. However, when starved before critical weight, these animals die 38 hours earlier than wild type controls (Figure 3E). This reduction in viability is due to the temperature sensitive mutation because hhts2 larvae do not differ from wild type controls when starved at the permissive temperature (16°C) (Figure 3E). We wondered whether circulating Hh acted at the time of starvation to increase larval survival. Alternatively, loss of circulating Hh during larval growth might affect nutrient storage, making them more sensitive to later starvation. To distinguish these possibilities, we shifted hhts2 larvae to 29°C coincident with starvation, and monitored their survival. These animals die 24 hours earlier than wild type (Figure 3F). These data confirm the phenotypes generated by hhRNAi in the midgut cells, and show that increased production of circulating Hh by the midgut is essential for normal survival under starvation conditions.
Hh produced in the midgut is not required for midgut homeostasis or midgut tracheal branching
We wondered whether Hh produced in the midgut was required locally to influence intestinal function. To ask whether altered Hh levels in the midgut might affect its structure or homeostasis, we assayed morphology, proliferation and apoptosis in the midgut. We also counted the number of undifferentiated adult midgut progenitor (AMPs) cells within a cluster, and total AMPs clusters in a given region of interest. Neither hh overexpression nor RNAi-mediated knock-down of hh in the midgut induced apoptosis or changed enterocyte cell size or morphology (suppl. Figure 3A). Furthermore, the total number of AMP clusters and the number of within a given cluster remained constant (suppl. Figure 3A,B), consistent with previous results (Jiang and Edgar 2009). Thus, Hh produced by midgut enterocytes does not locally influence their morphology or homeostasis.
To more directly assess whether Hh signaling might affect the function of ECs or AMPs, we autonomously increased or decreased pathway activity in these cell types by expressing RNAi or dominant negative/active constructs for Hh pathway components under the control of myo1A-Gal4 and esg-Gal4, respectively. The Hedgehog receptor, Patched (Ptc), represses signaling when Hh is not present. Binding of Hh to Ptc blocks Ptc-mediated repression of the 7-pass transmembrane protein Smoothened (Smo), which can mediate both transcriptional and non-transcriptional pathway outputs (van den Heuvel and Ingham 1996; Alcedo et al. 1996; Chen et al. 2001; DeCamp et al. 2000; Zhang et al. 2001; Ogden et al. 2008). We either activated Hh signaling using ptcRNAi, or inhibited it by smoRNAi in ECs or in AMPs. None of these manipulations altered cell morphology, apoptosis or proliferation in the midgut (suppl. Figure 3C,D). To further assess whether Hh signaling in ECs and AMPs could regulate systemic growth and developmental timing we measured the timing of pupariation and the resulting pupal volume after increasing or decreasing Hh signaling activity in midgut EC and AMPs. None of these manipulations affected pupal volume or the timing of pupariation (suppl. Figure 3E,F). Taken together, these results strongly suggest that Hh produced in the midgut is not required locally in either ECs or AMPs to regulate systemic growth and developmental timing.
Recently, the regulated branching of midgut trachea has been implicated in the regulation of systemic growth and developmental timing in response to metabolic changes (Linneweber et al. 2014).To test whether midgut hh expression might regulate systemic growth and developmental timing by affecting the structure of midgut trachea, we investigated their morphology in larvae with altered hh expression in the midgut. Examining trachea on 3 different regions of the midgut revealed no effect of hh overexpression or knock-down on tracheal size or morphology (suppl. Figure 3G). Thus, Hh produced by midgut enterocytes does not appear to influence the overlying trachea.
Circulating Hh does not influence local wing imaginal disc Hh signaling
We wondered whether circulating Hh influenced Hh signaling in the imaginal discs. The growth of imaginal discs exerts strong non-autonomous effects on larval developmental timing (Garelli et al. 2012; Colombani et al. 2012). Changes in imaginal disc Hh signaling due to circulating Hh might thereby indirectly affect larval developmental timing. We ask whether endogenously circulating Hh was required for normal patterning in the wing disc, we reduced circulating Hh levels in the gut using myo1A-Gal4 and monitored Hh levels and pathway activity in the wing disc. Hh is locally expressed in the posterior compartment of the wing disc, and spreads into the anterior compartment where its receptor ptc is expressed. Hh signaling stabilizes the full-length form of the Cubitus interruptus (Ci) transcription factor (Ci155), and upregulates transcription of target genes including ptc (Tabata and Kornberg 1994; Aza-Blanc et al. 1997). Reducing Hh levels in the hemolymph did not affect the distribution of Hh in the wing disc (Figure 4A,B). Neither did it change the range of Ci155 stabilization or ptc expression (Figure 4A,B), and flies emerged with normally patterned wings (not shown). Thus, endogenous levels of circulating Hh are not required for local patterning in wing imaginal discs. While we have previously observed that strong elevation of circulating Hh levels perturbs local patterning in the wing imaginal disc (Palm et al. 2013), more moderate increases produced by expressing Hh under the control of npc1b-Gal4 have no effect (Figure 4A,B). Nevertheless, npc1b-driven hh expression reduces larval growth and delays pupariation (Figure 2C). Thus, circulating Hh does not act indirectly through imaginal discs.
Circulating Hh does not influence the frequency of mouth hook contractions
The mammalian intestine produces hormones that act on the brain to regulate feeding behavior (Sandoval et al. 2008; Strader and Woods 2005). To ask whether circulating Hh produced by the midgut might influence larval feeding, we counted larval mouth hook contractions. Neither overexpression nor RNAi-mediated knock-down of hh in the midgut changed the frequency of larval mouth hook contractions (suppl. Fig 4). Thus, changes in growth and developmental timing are not a simple consequence of less enthusiastic feeding.
Circulating Hedgehog promotes neutral lipid mobilization in the fat body upon starvation
Since Hh signaling can reduce TAG levels in both flies and mammals, we wondered whether the influence of circulating Hh on starvation resistance, growth rate or pupariation timing was mediated through effects on fat metabolism. We first examined how loss or gain of circulating Hedgehog influenced TAG levels in normally fed animals, using shotgun mass spectrometry. We observed no change in the amount of stored TAG under either condition – either in the fat body or in whole animals (Figure 5A,B). Thus, altering the levels of circulating Hh is insufficient to change TAG metabolism in well-fed animals. These observations suggest that Hh influences growth and developmental timing by mechanisms that do not depend on TAG metabolism.
We next investigated whether circulating Hh might influence TAG metabolism under starvation conditions. In control larvae, starvation initially increases levels of whole larval TAG within 12 hours by almost 2-fold (Figure 5C). Interestingly, TAG’s do not increase in the fat body at this time - this increase must occur in other tissues (Figure 5D). Over the next 12 hours, both whole body and fat body TAG’s drop dramatically (Figure 5C,D). Wild type larvae normally die about 24 hours after TAG stores have dropped (48 hours after starvation) (Figure 4D).
Reducing the levels of circulating Hh during starvation completely blocks mobilization of fat body TAG’s (Figure 5D). However it does not prevent the disappearance of TAG’s from other tissues; whole body TAG’s drop between 12-24 hours to levels almost as low as those in control animals (Figure 5C). Residual whole body TAG at this time likely reflects failed mobilization of fat body stores. These larvae die 24 hours earlier than control larvae – right after the time that wild type animals would have mobilized fat body TAG stores. Thus, circulating Hh is required to mobilize fat body TAG stores under starvation conditions. However elevating circulating Hh in well-fed animals is not sufficient by itself to reduce TAG’s in the fat body – other signals active during starvation must work together with Hh to accomplish this.
Fat body Hh signaling regulates starvation resistance and larval growth
We wondered whether some of the effects of circulating Hh might be mediated by direct signaling to the fat body – indeed, the fat body expresses Hh pathway components, but not hh itself (Suh et al. 2006; Palm et al. 2013; Pospisilik et al. 2010 and suppl. Fig 5A). To investigate this, we first asked whether Hh protein was detectable in the fat body, despite the fact that it is not expressed there. Immunoblotting shows that the fat bodies of feeding third instar larvae indeed contain small amounts of Hh protein (Figure 6A). Fat body Hh protein levels, but not hh transcription, increase upon starvation (Figure 6A, and suppl. Figure 5B). These are precisely the conditions under which Hh production is upregulated in the midgut (Figure 3A,B). To determine whether Hh in the fat body was derived from the midgut, we reduced Hh production there and monitored Hh protein in the fat body of starving larvae by Western blotting. Indeed, hh knock-down in the midgut reduces Hh protein levels in the fat body (Figure 6B). Thus, the midgut is the source of Hh ligand found endogenously in the fat body. Furthermore, driving hh over-expression in the gut using npc1b-Gal4 elevates the amount of Hh in the fat body (Figure 6A). Thus, the amount of Hh accumulating in the fat body directly reflects the amount of Hh produced by the midgut.
To ask whether Hh signaling in the fat body might mediate effects on growth, pupariation and starvation resistance, we autonomously increased or decreased pathway activity in this tissue by expressing RNAi or dominant negative/active constructs for Hh pathway components under the control of lpp-Gal4. We first examined whether Hh signaling in the fat body controlled TAG metabolism or starvation resistance. Although manipulation of the pathway at the level of the systemic Hh ligand does not alter TAG stores, autonomous pathway activation and inactivation in the fat body cause opposing changes in whole animal TAG – consistent with previous reports (Suh et al. 2006; Pospisilik et al. 2010 and suppl. Fig 5C). Interestingly, both pathway activation and pathway repression reduced survival upon starvation (Figure 6C,D). These animals may die for opposite reasons. When Hh signaling is permanently activated in the fat body, larvae have less stored TAG to mobilize. When it is inactivated, they may be unable to mobilize it.
Is the fat body responsible for the alterations in growth and/or developmental timing caused by perturbing hh expression in the midgut? To test this, we monitored the timing of pupariation and resulting pupal volume after increasing or decreasing Hh signaling in the fat body. Activating the pathway by RNAi-mediated ptc knock-own decreased the larval growth rate (Figure 6E), but did not affect pupariation timing (Figure 6E,G). As a consequence, pupal volume was decreased (Figure 6E,F). Pathway inhibition caused by smoRNAi increased larval growth rate (Figure 6E), again without affecting the timing of pupariation (Figure 6E,G), resulting in larger pupae (Figure 6F). Similar results were obtained when Hh signaling was blocked by expression of the constitutively active ptcΔloop2 (Figure 6F,G). Interestingly, these effects of Ptc and Smo in the fat body appear to be exerted by a non-canonical pathway that does not involve processing of the transcription factor Cubitus Interruptus (Ci) (suppl. Text 1 and suppl. Figure 6). Instead, up and down-regulation of Hh causes opposing changes in specific fat body diacylglycerol species (suppl. Figure 6H). Taken together, these data show that the fat body mediates the effects of circulating Hh on larval growth, and does so by a non-canonical pathway. However, its effects on developmental timing must be exerted through a different tissue.
Hh signaling in the prothoracic gland regulates developmental timing
The timing of Ecdysteroid production by the prothoracic gland regulates developmental transitions including pupariation. We therefore asked whether Hh might signal to the prothoracic gland to control developmental timing. We first investigated whether Hh pathway components were present in this tissue using immunofluorescence. We detected Ptc, Smo and Ci155 specifically in the prothoracic gland, but not in the corpus allatum or corpus cardiacum, which produce other hormones (Figure 7A). Furthermore, Lpp, which carries circulating Hh, also accumulates specifically on cells of the prothoracic gland (Figure 7A). However Hh itself could not be detected by immunofluorescence (data not shown). To ask whether Hh produced in the midgut was able to travel to the ring gland, we expressed either hh-gfp or membrane tethered cd8-hh-gfp under the control of npc1b-Gal4 and stained ring glands with an antibody to GFP. Hh-GFP is clearly detected on cells of the prothoracic gland, but not when the midgut expresses the membrane-tethered version (Figure 7B). Thus, Hh made in the midgut can travel to the prothoracic gland, which expresses Hh pathway components.
To investigate the autonomous effects of Hh signaling in the prothoracic gland, we used phm-GAL4 to drive the expression of RNAi and constitutive pathway constructs in this tissue. We then monitored the rate of larval growth and timing of pupariation. Autonomously activating Hh signaling by RNAi-mediated knock-down of ptc in the prothoracic gland does not significantly affect the larval growth rate, but almost completely blocks pupariation (Figure 7C,D). Inhibiting Hh signaling by smo-RNAi expression only slightly delays puparation (8-12 hours after wild type controls) and larvae pupariate at the same size (Figure 7C,D). These data suggest that loss of Hh signaling alone is not sufficient to induce early pupariation, but that increased Hh signaling in the prothoracic gland inhibits pupariation. In contrast to its function in the fat body, Hh signaling in the prothoracic gland does not strongly influence larval growth (Figure 7C). Thus, the physiological effects of elevating circulating Hh can be accounted for by its combined action in the fat body and prothoracic gland – Hh signaling in the fat body reduces larval growth, and Hh signaling in the prothoracic gland delays pupariation.
How does Hh signaling in the prothoracic gland block pupariation? Pupariation is normally initiated by a rise in the level of ecdysteroid hormones. At early wandering stages, the prothoracic gland increases transcription of ecdysteroid biosynthetic enzymes and elevates production of ecdysone, which is hydroxylated by other tissues to 20-OH ecdysone (Petryk et al. 2003). To ask whether Hh signaling in the prothoracic gland blocked pupariation by preventing production of ecdysteroids, we quantified ecdysteroids over the course of the third instar and in early pupae using LC-MS/MS. In wild type third instar larvae, we detect both 20-OH ecdysone and the related hormone, Makisterone A, but not ecdysone or methyl-ecdysone (the precursor of Makisterone A) (Figure 7E,F). This suggests that 20-hydroxylation must be rapid and efficient at this stage. 20-OH ecdysone and Makisterone A levels begin to rise 56 hours after the molt from L2 to L3, and peak in white prepupae (Figure 7E,F). In contrast, larvae with constitutively active Hh signaling in the prothoracic gland fail to produce high levels of either 20-OH ecdysone or Makisterone A (Figure 7E,F). Only a slight rise in hormone levels is apparent even 96 hours after the molt from L2 to L3, long after wild type larvae have pupariated. Thus, constitutive Hh signaling in the prothoracic gland blocks 20-OH ecdysone and Makisterone A production. Quantitative RT-PCR shows that ptcRNAi in the prothoracic gland reduces mRNA levels for phantom, an ecdysteroid biosynthetic enzyme (Warren et al. 2004) (Figure 7G). This suggests that reduced transcription of biosynthetic enzymes contributes to lower ecdysone/methyl-ecdysone biosynthesis in the prothoracic gland.
Reducing Hh signaling by smo knock-down in the PG only slightly delays pupariation (Figure 7C). Consistent with this, these animals increase production of 20-OH ecdysone and MakisteroneA with slightly delayed kinetics compared to wild type (Figure 7E,F). Taken together, these data suggest that the increase in ecdysteroid biosynthesis that precedes and initiates pupariation does not occur if Hh signaling in the prothoracic gland is too high. However, reducing Hh signaling is not sufficient to initiate ecdysteroid biosynthesis. Lowered Hh signaling is likely only one of many inputs that regulate this important decision.
Discussion
The ability to couple nutrition to growth and developmental progression allows animals to thrive in unpredictable environments. Work in Drosophila has begun to outline how different organs communicate to coordinate these processes. Key interactions have been demonstrated between the fat body, prothoracic gland and central nervous system (Mirth et al. 2005; Colombani et al. 2005; Geminard et al. 2009; Delanoue et al. 2010). Our results highlight the importance of the intestine as another node in the signaling network. The intestine is well-placed to specifically sense intake of nutrients as opposed to their circulating levels, which may reflect other physiological conditions and not food availability alone.
We have shown that circulating Hh produced by midgut enterocytes inhibits ecdysteroid production in the prothoracic gland, delaying pupariation. However, blocking Hedgehog signaling in the prothoracic gland is insufficient to elevate ecdysteroid synthesis. Clearly, the information provided to the prothoracic gland by circulating Hh must be integrated with that of other signals. Insulin, TGFβ and PTTH signaling all act positively to increase ecdysteroid production (Mirth et al. 2005; Colombani et al. 2005; McBrayer et al. 2007; Rewitz et al. 2009; Gibbens et al. 2011).
Circulating Hh also signals to the fat body to slow larval growth, and to mobilize TAG’s during nutrient deprivation. Its action in the fat body and prothoracic gland are independent of each other. Growth and pupariation timing can be altered separately by autonomously activating Hedgehog signaling in either of these tissues by itself. Thus, circulating Hh couples systemic growth and developmental timing by its independent actions in the fat body and the prothoracic gland. While Hh acts directly on both these tissues, other studies have clearly shown that information can also flow back and forth between the ring gland and fat body. Humoral signals from the fat body, including the Leptin-related Unpaired2, promote release of Drosophila Insulin-like peptides (Dilp’s) release by the specific neurons in the brain (Rajan and Perrimon 2012; Geminard et al. 2009). This promotes insulin signaling in the ring gland to elevate ecdysteroid production (Mirth et al. 2005). Conversely, moderately increased levels of ecdysteroids act on the fat body to inhibit myc activity, which slows larval growth – possibly by reducing Dilp release (Colombani et al. 2005; Delanoue et al. 2010). Taken together, the signals flowing between the gut, fat body, IPC’s and prothoracic gland represent a robust information processing network that produces sensible decisions about growth and development.
We have highlighted the importance of intestinal derived circulating Hedgehog to regulate lipid mobilization and to couple systemic growth to developmental progression during nutritional stress. Interestingly, work by others shows that Hh transcription is increased by bacterial infection in the gut, and also by loss DHR96 (Chakrabarti et al. 2012; Bujold et al. 2010). This nuclear hormone receptor mediates the response to xeniobiotics and regulates TAG metabolism (King-Jones et al. 2006; Sieber and Thummel 2009). Thus it would be interesting to test whether circulating Hh might regulate growth and development in response to other physiological stresses.
A variety of different Hh secretion forms have been reported(Zeng et al. 2001; Chen et al. 2004; Tanaka et al. 2005; Panáková et al. 2005; Creanga et al. 2012; Ohlig et al. 2012; Palm et al. 2013). We have previously studied two different secretion forms – one that is sterol-modified and lipoprotein associated, and another that is monomeric and not sterol modified (Palm et al. 2013). These different forms exert different and synergistic effects on the signaling pathway in Drosophila imaginal discs (Palm et al. 2013). In contrast, signaling in the fat body is mediated entirely by the Lpp-associated form. In disc, the Lpp form of Hh blocks the repressive activity of lipoprotein lipids on Smoothened stability, but even high levels do not influence target gene activation by Cubitus Interruptus – this requires the non-sterol modified form (Palm et al. 2013). Consistently, the action of lipoprotein-associated Hedgehog in the fat body requires Ptc and Smo, but does not appear to act through Ci-dependent transcription. These findings emphasize the physiological importance of the lipoprotein-associated Hh form, and suggest that the fat body will be a good system to further study its unique effects on the Hh pathway. It will be interesting to know whether Lpp-associated Hh might directly influence lipid storage, or regulate the availability of signaling lipids in the fat body, as suggested by work in imaginal discs. (Khaliullina et al. 2009; Palm et al. 2013).
Recently, the a non-canonical Hedgehog signaling pathway has been implicated in regulating lipid and sugar metabolism in adult animals – including mice and flies (Pospisilik et al. 2010; Teperino et al. 2012). However, whether this occurs in response to a Hedgehog ligand, and what the source of such a ligand might be, was completely unknown. Here we show that circulating Drosophila Hh, produced by the intestine, regulates lipid and steroid hormone metabolism and adapts the animal to its nutritional environment. Since the presence of Hedgehog in circulation is conserved (Palm et al. 2013), it may be that circulating mammalian Shh exerts a similar function. It will be particularly interesting to also examine the impact of circulating Shh on adrenal steroidogenesis, given its function in Drosophila and the dramatic effects of glucocorticoids on lipid and sugar metabolism (Walker 2007). Interestingly, the production of Hh proteins in the gut is widely conserved – even in Cnidarians (van den Brink 2007; Matus et al. 2008). Thus, it seems possible that a metabolic function for circulating and lipoprotein-associated Hedgehog proteins might have arisen early in evolution.
Material & Methods
Fly Stocks
The following stocks were used: lpp-Gal4 (Brankatschk and Eaton 2010), en(105)-Gal4, en-Gal4 (Eugster et al. 2007), hh-Gal4, tub-Gal4, myo1A-Gal4 (Jiang et al. 2009), npc1b-Gal4 (Voght et al. 2007), phm-Gal4 (Ono et al. 2006), hhts2, UAS-hh (Strigini and Cohen 1997), UAS-ptcΔloop2 (Briscoe et al. 2001), esg-Gal4, UAS-cd8-gfp, wild-type Oregon R (OreR) and w- were obtained from Bloomington. UAS-Dicer2, smoRNAi, ptcRNAi were obtained from VDRC, Vienna, and hhRNAi was obtained from DGRC, Kyoto.
Antibodies
Antibodies used: Hh (Eugster et al. 2007), Cv-D (Palm et al. 2012), Ci 1C2 (Wang and Holmgren 1999), Ci 2A1,Ptc, Smo, Fu, Dlg, tubulin (DSHB), phospho-HistoneH3 (Cell Signaling) and GFP (Invitrogen)
Survival upon starvation
To measure survival, larvae from 4h egg collections were kept on normal food until 60h after egg laying (AEL). For experiments with hhts2 animals, eggs were collected at 22°C for 4h and were kept at permissive temperature (16°C) for 48h to allow embryonic development. Hatched hhts2 and control larvae were shifted to restrictive temperature either at 48h AEL or just prior to starvation. 2nd instar larvae were washed in PBS and transferred to 24 well plates (a single larva per well) filled with Agar/PBS 0.8% (w/v). Dead larvae were counted every 8h and the percentage of surviving animals from individual 24 well plates was plotted over time. For each genotype, three 24 well plates were quantified.
Growth rates and developmental timing analysis
Growth rate and developmental timing analysis was performed as described (Delanoue et al. 2010; Colombani et al. 2005). In brief, larvae of different genotypes from a 4h egg collection were synchronized at hatching (23h AEL), cultured in petri dishes (6 cm) filled with normal food at a density of 50 larvae per dish. Digital images were captured every 24h and the larval or pupal volume was estimated by quantifying the area of the length and the width of the animals in Fiji, then estimating the corresponding volume using the formula (L = larval length, w = larval width). The average volume of a least 15 animals was plotted over time in biological triplicates. Timing of pupariation was defined as the time at which 50% of the larvae have pupariated.
Hemolymph preparation
Diluted hemolymph was obtained by bleeding larvae with a tungsten needle in ice-cold PBS. Hemocytes were removed by centrifugation for 30min at 1500g, 4°C. To remove cellular fragments, the supernatant was subsequently spun for 30min at 16000g, 4°C. The supernatant from the second centrifugation step was snap-frozen in liquid nitrogen and stored at 80°C until use.
Immunohistochemistry on larval tissues
Tissues were dissected form larvae at the indicated time AEL in 1x PBS on ice, fixed in 4% paraformaldehyde (w/v) in PBS for 20min at RT. Tissues were permeabilized with 0.05% (v/v) Triton X-100 (PBX) for wing discs, 0.1% PBX for ring glands and fat bodies and 0.2% PBX for midguts. Wing discs were blocked with 0.1% (w/v) bovine serum albumin in PBX. Brain-ring gland complexes, fat bodies and midguts were block in 10% (v/v) normal goat serum in PBX. Tissues were incubated with primary antibody over night in blocking solution at 4°C. After several washes in with the respective blocking solutions, tissues were incubated with secondary antibody and/or with rhodamin-phalloidin (Invitrogen) for 3-4h at RT. Wing disc were blocked additionally with 0.1% (w/v) and 4% (v/v) normal goat serum for 30 min prior to the incubation with secondary antibody. Tissues were washed with PBX several times and incubated with DAPI for 5 min and mounted in either Prolong Gold Antifade reagent (Invitrogen) for wing discs or VectaShield (Vectalabs) for other tissues.
Lipids extraction and shotgun mass spectrometry
Lipids were extracted from fat body and from whole larvae and quantified by shotgun mass spectrometry as described in (Carvalho et al. 2012) with modifications. Briefly, samples were homogenized in 150 mM ammonium bicarbonate buffer. From each homogenate, a volume containing ∼ 2 nmol of lipid material was removed and extracted using a modified Folch protocol at 4°C (WL n=3, FB n=4 fed to 12h of starvation, n=6 for 24h of starvation experiment).
Mass spectrometry analyses were performed on a Q Exactive instrument (Thermo Fisher Scientific, Bremen, Germany) equipped with a robotic nanoflow ion source TriVersa NanoMate (Advion BioSciences,Ithaca, NY) using chips with the diameter of spraying nozzles of 4.1 mm. The ion source was controlled by Chipsoft 8.1 software. Ionization voltage was set at +0.95 kV and -0.95 kV and back pressure was set at 1.25 and 0.6 psi in positive and negative modes, respectively. The temperature of ion transfer capillary was 200°C; tube voltages were 90 V (MS+) and -150V (MS-). Acquisitions were performed at the mass resolution Rm/z400=140 000. AGC control was set at 106 ions and maximum injection time to 250 ms.
Dried lipid extracts were re-suspended in 100 µL chloroform-methanol mixture 1:2 (v/v). For the analysis, 10 µL of samples were loaded onto 96-well plate (Eppendorf, Hamburg) and diluted either with 13 µL of 13.3 mM ammonium acetate in 2-propanol for positive ion mode analysis or with 10 µL of 0.01% methylamine in methanol for negative ion mode analysis. The plate was sealed with aluminum foil. Each sample was analyzed for ca 1min in positive and negative ion modes: PE, PC, PC-O, TAG, CerPE and DAG were quantified in positive and PI, PS, PG, PE, PEO-, Cer, HexCer in negative mode.
Lipids were identified by LipidXplorer (Herzog et al. 2011) software by matching m/z of their monoisotopic peaks to the corresponding elemental composition constraints. Molecular Fragmentation Query Language (MFQL) queries compiled for all the aforementioned lipid classes are available at the LipidXplorer wiki site: https://wiki.mpicbg.de/wiki/lipidx/index.php/Main_Page. Mass tolerance was 5 ppm and intensity threshold was set according to the noise level reported by Xcalibur software (Thermo Scientific).
The content of glycerolipids was normalized per polar lipids and expressed in pmol lipid x/pmol polar lipids. Lipids standards were purchased from Avanti Polar Lipids (Alabaster, Alabama). Solvents were purchased from Sigma-Aldrich (Taufkirchen, Germany).
Quantification of ecdysteroids by LC-MS/MS
Larvae of different genotypes from a 4h egg collection were synchronized at hatching (23h AEL), cultured in petri dishes filled with normal food at a density of 50 larvae each plate. Larvae were staged again at the 2nd to 3rd instar transition (AEL3). Every 4h groups of 4 animals were collected in 100 µl H2O and smashed with plastic pestle and extracted overnight with 1 ml of cold (6°C) methanol. Samples were centrifuged and supernatant was collected. The pellet was twice re-extracted with 1 ml methanol for 1 hour, the extracts combined, cleaned by solid-phase extraction, dried down and analyzed by LC-MS/MS as described in (Hikiba et al. 2013) with minor modifications. The detection limit of the method was ca 5 pg per individual ecdysteroid. Lipids standards were purchased from Avanti Polar Lipids (Alabaster, Alabama). Solvents were purchased from Sigma-Aldrich (Taufkirchen, Germany).
For additional Material & Methods see supplemental material
Acknowledgments
We are grateful to Bruce Edgar, Michael O’Connor, Isabell Guerrero and the Bloomington Stock Center for fly stocks. We gratefully acknowledge Robert Holmgren and the Developmental Studies Hybridoma Bank for providing antibodies. We thank Sven Ssykor and Marko Brankatschk for help with transgenesis. We thank Andreas Sagner, Marko Brankatschk and Jochen Rink for critical comments on the manuscript.
Footnotes
Contact: eaton{at}mpi-cbg.de phone +49 351 210-2526 fax +49 351 210-1309