Spatial Activation of TORC1 Is Regulated by Hedgehog and E2F1 Signaling in the Drosophila Eye.

Article

Spatial Activation of TORC1 Is Regulated by Hedgehog and E2F1 Signaling in the Drosophila Eye Graphical Abstract

Authors Wonho Kim, Yoon-Gu Jang, Jinsung Yang, Jongkyeong Chung

Correspondence [email protected]

In Brief How target of rapamycin complex 1 (TORC1), a critical activator for cell growth, is regulated during animal development is largely unknown. Kim et al. demonstrate that TORC1 is active in spatially restricted cells in the Drosophila eye, which requires Hedgehog-E2F1 signaling and is necessary for synchronous cell-cycle progression.

Highlights d

TORC1 is selectively active in the second mitotic wave (SMW) of Drosophila eye disc

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Hedgehog activates TORC1 independently of insulin and amino acid signaling

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TORC1 regulates G1/S transition downstream of Hh signaling in the SMW

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TORC1 regulation by Hedgehog is conserved in mammalian cells

Kim et al., 2017, Developmental Cell 42, 363–375 August 21, 2017 ª 2017 Elsevier Inc. http://dx.doi.org/10.1016/j.devcel.2017.07.020

Developmental Cell

Article Spatial Activation of TORC1 Is Regulated by Hedgehog and E2F1 Signaling in the Drosophila Eye Wonho Kim,1,3 Yoon-Gu Jang,1,2,3 Jinsung Yang,1,2 and Jongkyeong Chung1,2,4,* 1National Creative Research Initiatives Center for Energy Homeostasis Regulation and Institute of Molecular Biology and Genetics, Seoul National University, Seoul 08826, Republic of Korea 2School of Biological Sciences, Seoul National University, Seoul 08826, Republic of Korea 3These authors contributed equally 4Lead Contact *Correspondence: [email protected] http://dx.doi.org/10.1016/j.devcel.2017.07.020

SUMMARY

Target of rapamycin complex 1 (TORC1) regulates cell growth in response to nutrients and growth factors. Although TORC1 signaling has been thoroughly studied at the cellular level, the regulation of TORC1 in multicellular tissues and organs has remained elusive. Here we found that TORC1 is selectively activated in the second mitotic wave (SMW), the terminal synchronous cell division, of the developing Drosophila eye. We demonstrated that Hedgehog (Hh) signaling regulates TORC1 through E2F1 and the cyclin D/Cdk4 complex in the SMW, and this regulation is independent from insulin and amino acid signaling pathways. TORC1 is necessary for the proper G1/S transition of the cells, and the activation of TORC1 rescues the cell-cycle defect of Hh signaling-deficient cells in the SMW. Based on this evolutionarily conserved regulation of TORC1 by Hh signaling, we propose that Hh-dependent developmental signaling pathways spatially regulate TORC1 activity in multicellular organisms.

INTRODUCTION Target of rapamycin complex 1 (TORC1) signaling forms the central axis of cell growth and division regulation in response to diverse physiological information, including nutrient availability, hormones, and growth factors (reviewed in Dibble and Manning, 2013). TORC1, a kinase complex composed of TOR kinase, Raptor, and LST8, activates ribosomal protein S6 kinase (S6K) and inhibits eukaryotic initiation factor 4E-binding protein (4E-BP) via direct phosphorylation (Burnett et al., 1998; Chung et al., 1992). Activated S6K then phosphorylates ribosomal protein S6 (RpS6) to regulate protein translation (reviewed in Ruvinsky and Meyuhas, 2006). TORC1 kinase activity requires Rheb, a small GTP-binding protein (Saucedo et al., 2003), which is negatively regulated by the GTPase-activating protein domaincontaining TSC1/TSC2 complex (the TSC complex) (Garami

et al., 2003; Inoki et al., 2003). In particular, the TSC complex integrates multiple intra- and extracellular inputs, such as insulin, hypoxia, and energy deprivation, to regulate TORC1 signaling activity (reviewed in Laplante and Sabatini, 2012). The molecular mechanism for how extracellular cues regulate TORC1 has been vigorously studied in mammalian cells and model animals including Drosophila and Caenorhabditis elegans. Among them, insulin and amino acid availability have been most extensively studied. Firstly, insulin regulates TORC1 positively by evoking a signal cascade starting from insulin receptor. When insulin receptor tyrosine kinase is activated, phosphatidylinositol 3-kinase (PI3K) is recruited to the receptor via insulin receptor substrate (Chico in Drosophila) and converts phosphatidylinositol (4,5)-bisphosphate to phosphatidylinositol (3,4,5)-trisphosphate (PIP3) in the plasma membrane, which is counteracted by phosphatase and tensin homolog (PTEN), a phosphatase responsible for dephosphorylating PIP3 (reviewed in Song et al., 2012). Increased PIP3 recruits phosphoinositidedependent kinase 1 (PDK1) and Akt to the plasma membrane via their pleckstrin homology domains, and PDK1 subsequently phosphorylates and activates Akt, which inhibits the TSC complex by phosphorylation (Inoki et al., 2002; Potter et al., 2002). The importance of amino acid availability in mTORC1 activity in mammalian cells, which was reported almost two decades ago (Hara et al., 1998; Wang et al., 1998), has recently drawn high interest among researchers. RagA/C GTPases are responsible for conveying signals from amino acid to mTORC1 by binding to Raptor, and thereby recruiting the whole TOR complex to mTORC1-activating Rheb in the lysosome. (Kim et al., 2008; Sancak et al., 2008). The Ragulator complex localizes Rag GTPases at the lysosome and promotes the GTP-GDP nucleotide exchange of RagA (Sancak et al., 2010). The GATOR1 and GATOR2 complexes sequentially regulate RagA in amino acid signaling. The GATOR1 complex induces GTP hydrolysis in RagA and inactivates mTORC1, and the GATOR2 complex inhibits the GATOR1 complex upon amino acid stimulation (Bar-Peled et al., 2013). The Drosophila eye is an excellent model system for understanding how signaling networks are organized in multicellular tissues (reviewed in Baker, 2007). It consists of approximately 750 repetitive units called ommatidia, and each ommatidium contains eight photoreceptor cells as well as non-neuronal

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accessory cells (Ready et al., 1976). During the early third-instar larval stage, an indentation called the morphogenetic furrow (MF) is formed at the posterior end of the Drosophila eye disc, which then progresses anteriorly (Wolff and Ready, 1991). In the far anterior of the MF, undifferentiated cells proliferate at random until they are arrested in G1 phase (Ready et al., 1976). After G1 phase, some precursor cells in the MF begin to sequentially differentiate into R8, R2/5, and R3/4 photoreceptor neurons (Ready et al., 1976), while other precursor cells enter S phase for an additional round of cell division, which is referred to as the SMW, and then differentiate into R1/6 and R7 photoreceptor neurons, cone cells, pigment cells, and interommatidial bristles (Wolff and Ready, 1991). Several signaling pathways coordinate various events during Drosophila eye development, such as cell fate determination, pattern formation, and morphogenesis. Hedgehog (Hh) signaling is required for both MF initiation/progression and photoreceptor neuron differentiation, and is also critical for the S phase entry of the cells in the SMW (Dominguez and Hafen, 1997; Firth and Baker, 2005). In the absence of Hh ligand, Patched (Ptc) suppresses Smoothened (Smo) by promoting its endocytosis and degradation, thereby inhibiting the Hh signaling (Denef et al., 2000; Li et al., 2012). The kinesin-like protein Costal2 moves along microtubules and serves as a scaffold for the Hh signaling complex (HSC), connecting Cubitus interruptus (Ci), a transcription factor, to its regulators (Farzan et al., 2008). In the absence of Hh stimulation, the HSC is associated with microtubules and Ci is sequentially phosphorylated by multiple kinases at its C-terminal region, leading to its partial degradation by Slimb E3 ligase to form a transcriptional repressor of Ci (CiR) (reviewed in Jiang, 2006). CiR then enters the nucleus and represses transcription of the target genes of Hh signaling. Upon binding of the Hh ligand, however, Ptc is not able to induce the degradation of Smo, which then associates with the HSC and inhibits the degradation of Ci (Denef et al., 2000; Zhu et al., 2003). Consequently, the fulllength Ci (CiFL) is activated by further post-translational modifications and translocates into the nucleus to induce the expression of target genes of Hh signaling (reviewed in Briscoe and Therond, 2013). In this study, we investigated the regulatory mechanisms of TORC1 signaling during Drosophila eye development. By examining the phosphorylation of RpS6, we found that TORC1 is specifically activated in the SMW cells of the Drosophila eye imaginal disc. Interestingly, Hh and E2F1-cyclin D/Cdk4 signaling pathways activated TORC1 in the eye disc, independently to the previously known TORC1 regulators in insulin and amino acid signaling pathways. These results provide a regulatory mechanism for the tissue-specific activation of TORC1 during animal development. RESULTS RpS6 Phosphorylation Is Modulated by TORC1 Signaling in Drosophila To examine the regulatory mechanisms of TORC1 in Drosophila tissues, we generated an antibody reagent that would allow us to monitor Drosophila TORC1 activity through immunostaining. Mammalian TORC1 activity has been successfully analyzed in mammalian cells by immunostaining for phosphorylated RpS6 364 Developmental Cell 42, 363–375, August 21, 2017

(Hong et al., 2012). Interestingly, the C-terminal regions of Drosophila and human RpS6 contain highly conserved amino acid sequences (Figure S1A). Moreover, it was previously reported that insulin treatment increases the phosphorylation of five serine residues (Ser233, Ser235, Ser239, Ser242, and Ser245) in the C-terminal region of Drosophila RpS6. In addition, this was blocked by rapamycin treatment in Drosophila cells (Radimerski et al., 2000), suggesting that TORC1 is fully responsible for RpS6 phosphorylation in Drosophila. Based on the previous studies mentioned above, we generated a phosphospecific antibody against a peptide consisting of Drosophila RpS6 amino acids 227–238 phosphorylated at Ser233 and Ser235 (Figure S1A). By immunoblotting, we found that the antibody recognized wild-type RpS6 but not mutant RpS6, in which Ser233 and Ser235 were switched to alanines (data not shown). To confirm that RpS6 phosphorylation is modulated by TORC1 activity, amino acids were depleted from S2 cells in order to inhibit TORC1, and replenished later to reactivate it. In accordance with previous reports, supplying amino acids increased S6K phosphorylation at Thr398 (Figure 1A) (Kim et al., 2008; Sancak et al., 2008). Similarly, RpS6 phosphorylation was induced by the treatment of amino acids (Figure 1A). Moreover, S6K phosphorylation was inhibited by treatment with double-stranded RNA (dsRNA) for TOR and Raptor, which are the components of TORC1 (Figure 1B). In addition, dsRNAmediated knock down of Rheb, which is required for TORC1 activity, blocked S6K phosphorylation, whereas knock down of tsc2, a negative regulator of Rheb, increased S6K phosphorylation (Figure 1B). Likewise, RpS6 phosphorylation was inhibited by knock down of TOR, Raptor, and Rheb, and was increased by knock down of tsc2 (Figure 1B). Furthermore, knock down of S6K, which directly phosphorylates RpS6, almost completely blocked the phosphorylation of RpS6 (Figure 1B). These results indicated that RpS6 phosphorylation can be used as a specific marker to monitor TORC1 activity in Drosophila. TORC1 Activity Is Selectively Increased in S-Phase Cells of the SMW in the Eye Disc Next, we immunostained various tissues with our anti-phosphospecific RpS6 antibody to examine the pattern of TORC1 activity in Drosophila tissues. We observed that RpS6 was highly phosphorylated near the MF in the eye disc of wandering larvae (Figures 1C and 1D). To support this result, we examined RpS6 phosphorylation with another antibody. It was previously reported that human RpS6 ectopically expressed in Drosophila cells was also phosphorylated upon TORC1 activation (Lindquist et al., 2011). Therefore, we generated a transgenic fly in which human RpS6 was expressed throughout the entire body under control of the tubulin promoter (tub-hS6) (Figure S1B). We confirmed the expression of human RpS6 in the whole body by immunoblotting, and in the eye disc by immunostaining, using the antibody for human RpS6, which does not detect Drosophila RpS6 (Figures S1C–S1E). Then we observed that human RpS6 was phosphorylated at Ser235 and Ser236 in the transgenic fly by immunoblotting with a commercially available anti-phosphospecific human RpS6 antibody (Figure S1C). Parallel to the results with the Drosophila RpS6, we found that the ectopically expressed human RpS6 was highly phosphorylated near the MF (compare Figures S1F and S1G with Figures 1C and 1D).

Figure 1. TORC1 Is Selectively Activated in the S-Phase Cells of the SMW (A) S2 cells were subjected to amino acid and serum starvation for an hour, and then they were replenished with amino acid for 20 min. Lysate samples were prepared, separated by SDS-PAGE, and immunoblotted with the anti-pS6, -pS6K, -S6K, and -b-tubulin antibodies. (B) Specific dsRNAs were used to knock down S6K, TOR, Raptor, Rheb, or tsc2 in S2 cells as indicated. dsRNA for Luciferase was used as a control. After 4 days, the S2 cells were serumstarved for 4 hr. Lysate samples were separated by SDS-PAGE and subsequently immunoblotted by the anti-pS6, -pS6K, -S6K, and -b-tubulin antibodies. (C) The eye discs of wandering w1118 larvae were immunostained with Hoechst (DNA, blue) and the anti-pS6 (red) antibody. Arrowhead indicates the MF. (D) Boxed area of (C) was magnified. (E) The eye disc expressing a GFP-fused PCNA via endogenous PCNA promoter was immunostained with the antibody against pS6 (red). (F) The boxed area of (E) was magnified. (G and H) S6K (G) and TOR (H) mutant clones were generated and immunostained with the anti-pS6 antibody (red). The absence of GFP indicates the mutant clones. (I) GFP-positive Raptor RNAi-expressing clones were stained with the anti-pS6 antibody (red). (J) Rheb mutant clones marked by the absence of GFP were examined by immunostaining with antipS6 antibody (red). (K) GFP-positive Rheb-expressing clones were immunostained with the anti-pS6 antibody (red). (L) GFP-negative clones in which tsc2 was mutated were immunostained with the anti-pS6 antibody (red). (K and L) Due to the strong anti-pS6 immunostaining signals from clones, the pS6 signals in the SMW looked very faint. The boxed areas in (G)–(L) are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. Scale bars, 30 mm. See also Figures S1–S3.

Taken together, the results of the immunostaining experiments using two independent antibodies consistently indicated that RpS6 is selectively phosphorylated near the MF in the Drosophila eye disc. To investigate the regulation of TORC1 during Drosophila eye disc development, we first determined the identity of the cells with increased RpS6 phosphorylation. Our primary candidate was S-phase cells of the SMW, because they are known to locate near the MF. To verify our hypothesis, we utilized a

fly strain expressing GFP-fused PCNA driven by the endogenous promoter of PCNA so that the GFP signal can be used as a marker of the S-phase cells (Thacker et al., 2003). Indeed, phosphorylated RpS6 was detected in the PCNAGFP-positive cells (Figures 1E and 1F). As a result, we concluded that RpS6 is selectively phosphorylated in the S-phase cells of the SMW. Subsequently, we examined whether the phosphorylation of RpS6 is dependent on TORC1 activity in the eye disc. We first checked if S6K or TOR mutation altered the phosphorylation of RpS6 and found that the phosphorylation was completely blocked in S6K or TOR mutant clones generated by FRT-mediated recombination in the eye disc (Figures 1G and 1H, respectively). To prove that S6K exclusively modulates Developmental Cell 42, 363–375, August 21, 2017 365

RpS6 phosphorylation in the eye disc, we examined whether RpS6 phosphorylation was decreased by mutation of p90 ribosomal S6 kinase (RSK), another kinase known to phosphorylate RpS6 in mammals (Anjum and Blenis, 2008). Since RpS6 phosphorylation was not significantly decreased in RSK mutant eye disc (data not shown) (Kim et al., 2008), we concluded that RpS6 phosphorylation depends solely on S6K, not RSK. In addition, to determine which of the two TOR kinase-containing complexes, TORC1 and TORC2, is responsible, we clonally expressed Raptor RNAi by flip-out recombination to specifically inhibit TORC1. As a result, we observed that Raptor RNAi expression dramatically decreased RpS6 phosphorylation in the SMW (Figure 1I). Collectively, these results demonstrated that TORC1 is selectively activated in the S-phase cells of the SMW to induce phosphorylation of RpS6 during Drosophila eye development. Next, we investigated whether Rheb and the TSC complex regulate TORC1 activity in the eye disc. When we generated Rheb mutant clones or clones simultaneously overexpressing TSC1 and TSC2, TORC1 activity was completely inhibited (Figures 1J and S2A, respectively), whereas clonal overexpression of Rheb or tsc2 mutation resulted in strong activation of TORC1 in the eye disc (Figures 1K and 1L, respectively). When S6K was mutated simultaneously with tsc2, RpS6 phosphorylation was completely blocked, providing additional evidence that the TORC1 activation in the tsc2 mutant cells caused the hyperphosphorylation of RpS6 (Figure S2B). Taken together, these results indicated that Rheb and the TSC complex regulate TORC1 activity in the eye disc. TORC1 Activity Is Spatially Regulated in Various Drosophila Tissues We further examined RpS6 phosphorylation in other larval and adult tissues. We first observed that human RpS6 expressed in the ovary of tub-hS6 flies was phosphorylated specifically in the follicle cells (Figure S3A), whereas in the fat body cells of the wild-type larvae RpS6 was uniformly phosphorylated in all cells (Figure S3B). We also found that human RpS6 was phosphorylated in the wandering larval stage at several peptidergic cells in the brain marked by expression of a bHLH transcription factor, dimmed, which has a role in the differentiation of peptidergic cells (Figure S3C) (Park et al., 2008). In the adult brain, human RpS6 was highly phosphorylated in the neuropeptide neurons including insulin-producing cells and Pdf-expressing circadian neurons (Figures S3D–S3F). In agreement with our observation, TORC1 activity in Pdf-expressing cells was reported to modulate circadian behaviors (Zheng and Sehgal, 2010). Moreover, we observed that nutrient restriction significantly decreased RpS6 and human RpS6 phosphorylation in the fat body and the adult brain, respectively, suggesting that TORC1 activity can be monitored by RpS6 phosphorylation in these tissues (Figures S3G–S3J). Insulin and Amino Acid Signaling Pathways Are Required for Activation of TORC1 in the SMW In Drosophila and mammalian cells, insulin signaling regulates TORC1 activity positively. Therefore, we decided to investigate TORC1 regulation by insulin signaling in the eye disc. Firstly, by immunostaining with an antibody against phosphorylated 366 Developmental Cell 42, 363–375, August 21, 2017

Akt, which is induced upon active insulin signaling in Drosophila (Kockel et al., 2010), we explored which cells in the eye disc had active insulin signaling and found that Akt was phosphorylated evenly in all cells of the eye disc without any patterns (Figure 2A). To verify the specificity of this antibody, we immunostained phosphorylated Akt in the mosaic eye disc containing clones expressing a dominant negative form of PI3K, and observed specifically reduced Akt phosphorylation in such clones (Figure S4A). On the contrary, loss-of-function mutation of PTEN highly increased Akt phosphorylation (Figure S4B). These results indicated that insulin signaling is uniformly activated in the eye disc. To determine whether insulin signaling is required for TORC1 activity in the eye disc, we clonally expressed dominant negative forms of the insulin receptor and PI3K and Akt RNAi. As a result, we found that TORC1 activity was inhibited in these cells (Figures 2B–2D, respectively). Collectively, these results indicated that insulin signaling is required for TORC1 activity in the SMW. Next, the regulation of TORC1 by amino acid signaling was tested in the eye disc. In mammalian and Drosophila cells, amino acids are known to regulate TORC1 through GATOR1 and 2, Ragulator, and RagA/C GTPases. In the Drosophila eye, expression of a dominant negative form of RagA and a loss-of-function mutation of RagC blocked TORC1 activation in the SMW, indicating that RagA/C GTPases are required for TORC1 activity (Figures 2E and 2F, respectively). In addition, the RNAi-based downregulation of HBXIP and p18, the components of the Ragulator complex, in the dorsal region of the eye disc using DE-GAL4 inhibited TORC1 activity (Figures 2G and 2H, respectively). Next, when RNAi for mio, a component of the GATOR2 complex, was expressed, TORC1 activity was severely decreased (Figure 2I). Moreover, larvae with homozygous mutation in seh1, another component of the GATOR2 complex, also showed inhibited TORC1 activity in the SMW (Figure 2J). Collectively, these results indicated that amino acid signaling, which has been thoroughly delineated in mammalian cells, is required for TORC1 activity in the eye disc of Drosophila in a conserved manner. Hh Signaling Is Indispensable for TORC1 Activity in the SMW by Preventing CiR Formation Independently of Notch Signaling Next, the regulation of TORC1 through developmental signaling pathways, in addition to the insulin and amino acid signaling, was examined in the SMW of the eye disc. During Drosophila eye development, Hh signaling plays critical roles in many processes, including not only MF initiation/progression and photoreceptor neuron differentiation, but also cell-cycle progression in the SMW (Dominguez and Hafen, 1997; Heberlein et al., 1993). Therefore, we investigated whether Hh signaling is required for TORC1 activity in the SMW. For this purpose, we generated mutant clones of Smo, a transmembrane protein critical for Hh signaling, and observed that TORC1 was inhibited when Smo was mutated in the SMW (Figure 3A). This result suggested that Hh signaling is required for TORC1 activity in the SMW. Because Hh signaling regulates target gene expression by controlling Ci, we examined the role of Ci in the regulation of TORC1 activity. Without Hh, the C-terminal region of CiFL is partially degraded to generate CiR, the repressive form of the

Figure 2. Insulin and Amino Acid Signalings Are Required for TORC1 Activity in the SMW (A) Wild-type eye disc was immunostained with the anti-pAkt antibody (red). (B–D) InRDN- (B), PI3KDN- (C), and Akt RNAi (D)-expressing clones were marked by GFP and immunostained with the anti-pS6 antibody (red). (E) A dominant negative form of RagA was clonally expressed in the eye disc, which is indicated by GFP. RpS6 phosphorylation (red) was examined. (F) Phosphorylated RpS6 (red) was immunostained in the eye disc in which RagC mutant clones (GFP-negative) were generated. (G–I) RNAi targeting HBXIP (G), p18 (H), or mio (I) was expressed using DE-GAL4, and RpS6 phosphorylation (red) was examined. Dicer2 was expressed together for the higher knockdown efficiency. (J) The eye disc of seh1 mutant larvae was immunostained with the anti-pS6 antibody (red). (A and G–J) DNA was marked by Hoechst (blue). (B–F) The boxed areas are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. (G–I) The dorsal regions in which transgenes were expressed by DE-GAL4 are above the dotted lines. Scale bars, 30 mm. See also Figure S4.

protein, whereas the binding of the Hh ligand to its receptor stabilizes and activates CiFL (Aza-Blanc et al., 1997). Loss of Smo would roughly have two consequences on Ci: reduction of CiFL and accumulation of CiR. It has been reported that Ci mutant clones show normal eye disc development, suggesting that CiFL does not have a critical role in the process (Pappu et al., 2003). On the other hand, simultaneous mutation of Ci rescues the cell-cycle progression defect in the Smo mutant cells (Firth and Baker, 2005). These reports suggest the possibility that Hh signaling functions through inhibiting CiR accumulation rather than stabilizing CiFL during Drosophila eye development. To figure out whether CiR accumulation is responsible for the TORC1 inactivation in the Smo mutant cells, RNAi for Ci was expressed in Smo mutant clones using MARCM, which restored RpS6 phosphorylation (Figure 3B). Furthermore, the clonal expression of CiR resulted in suppressed phosphorylation of

RpS6 (Figure 3C). As expected, the expression of RNAi for Ci by DE-GAL4 to induce a condition mimicking reduced CiFL resulted in no difference in phosphorylated RpS6 level (Figure 3D). Although these data do not fully explain how TORC1 is activated in the eye disc in a patterned manner, from the results above we concluded that the amount of CiR, rather than CiFL, is responsible for keeping TORC1 active at downstream of Hh signaling in the SMW. Since Hh signaling has been reported to regulate Notch (N) signaling in the SMW (Baonza and Freeman, 2005; Firth and Baker, 2005), we tested if N signaling mediates TORC1 regulation of Hh signaling in the eye disc. When N signaling was manipulated by generating N mutant clones, TORC1 activity was not reduced (Figure S5A). Any other possible roles of N ligands Delta (Dl) and Serrate in TORC1 regulation were examined. When the clones of cells with amorphic mutant alleles of both ligands Developmental Cell 42, 363–375, August 21, 2017 367

Figure 3. Hh Signaling Is Required for TORC1 Activity through Inhibiting the Formation of CiR (A) RpS6 phosphorylation (red) was examined in Smo mutant clones (GFP-negative). (B) Smo mutant clones expressing Ci RNAi (GFPpositive) were generated in the eye disc, which was stained with the anti-pS6 (red) and -Ci (yellow) antibodies. (C) CiR was clonally expressed with GFP in the eye disc, and phosphorylated RpS6 (red) was immunostained. (D) The eye disc in which Ci RNAi and Dicer2 were expressed using DE-GAL4 was immunostained with the anti-pS6 (red) and -Ci (green) antibodies. The dorsal region in which transgenes were expressed by DE-GAL4 is above the dashed line. (A–C) The boxed areas are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. Scale bars, 30 mm. See also Figure S5.

were generated, TORC1 activity was not reduced (Figure S5B). Furthermore, to test the possibility that reduction of N signaling activity caused TORC1 inactivation in Smo mutant cells, the intracellular domain of N (Nintra) was ectopically expressed. However, RpS6 phosphorylation was not restored in the Nintra-expressing Smo mutant cells (Figure S5C). These results suggested that N signaling is not responsible for TORC1 regulation at downstream of Hh signaling. In addition, the role of TORC1 activity in N signaling downstream of Hh signaling was also examined. Because it was reported that Dl expression was reduced in Smo mutant cells, which led to reduced N signaling activity (Baonza and Freeman, 2005; Firth and Baker, 2005), we immunostained Dl in mosaic eye discs containing TOR mutant cells, and observed that Dl staining was not reduced (Figure S5D), suggesting that TORC1 does not regulate N signaling at the downstream of Hh signaling. From these experiments, we concluded that TORC1 is regulated by Hh signaling independently of N signaling. E2F1 and the Cyclin D/Cdk4 Complex Positively Regulate TORC1 Downstream of Hh Signaling We further investigated how Smo-Ci signaling activates TORC1 in the SMW. As shown in Figures 1E and 1F, TORC1 was activated in PCNA-GFP-positive cells. Because the PCNA gene is a target of the E2F1 transcription factor (Thacker et al., 2003), we hypothesized that E2F1 regulates TORC1. To test this hypothesis, we expressed E2F1 RNAi in the dorsal region and observed a decrease in RpS6 phosphorylation (Figure 4A), indicating that E2F1 is indeed required for TORC1 activity. Consistently, downregulation of Rbf1, a negative regulator of E2F1, by RNAi expression caused ectopic activation of TORC1 in the proximal portion of the S-phase cells in the SMW (Figure 4B). These results indicated that E2F1 positively regulates TORC1 activity in the eye disc. To further examine whether E2F1 regulates TORC1 downstream of Hh signaling, we generated Smo mutant clones and 368 Developmental Cell 42, 363–375, August 21, 2017

evaluated PCNA-GFP protein levels. Interestingly, PCNA-GFP was significantly decreased in the clones, suggesting that Hh signaling is required for the transcriptional activity of E2F1 (Figure 4C). To test whether E2F1 activation could restore the TORC1 inhibition phenotype in cells with Smo mutation, Smo mutant clones were generated in the eye disc in which Rbf1 RNAi was expressed in the dorsal region. Interestingly, TORC1 was clearly rescued by the knock down of Rbf1 in the clones (Figure 4D), indicating that E2F1 is an important mediator in TORC1 regulation by Hh signaling. Taken together, we concluded that Hh signaling regulates TORC1 through E2F1 in the SMW. To find the next downstream effector of Hh-E2F1 signaling in TORC1 regulation, various signaling molecules involved in cell growth and cell-cycle regulation were investigated, and the cyclin D/Cdk4 complex was considered to be a possible candidate. When RNAi for cyclin D or Cdk4 was expressed by DEGAL4, RpS6 phosphorylation was dramatically reduced in the dorsal region of the eye discs in both cases (Figures 4E and 4F, respectively), demonstrating that both cyclin D and Cdk4 are indispensable for TORC1 activity in the SMW. As a next step, we examined whether ectopic expression of cyclin D and Cdk4 can also allow elevated activity of TORC1 by overexpressing them with DE-GAL4. Similar to the results from Rbf1 RNAi expression, RpS6 phosphorylation was increased significantly in the proximal region of the SMW (Figure 4G; see also Figure 4B). Owing to the results so far, we hypothesized that the cyclin D/Cdk4 complex is downstream of E2F1, activating TORC1 in the SMW. To verify the hypothesis, we overexpressed cyclin D and Cdk4 along with RNAi for E2F1 using a DE-GAL4 driver, and observed that RpS6 phosphorylation was recovered in the dorsal region of the eye disc (Figure 4H). From these results, we suggest that the cyclin D/Cdk4 complex mediates Hh-E2F1 signaling to activate TORC1 in the SMW. It is notable, however, that the RpS6 phosphorylation pattern is not the same in the cases of cyclin D and Cdk4 overexpression with or without E2F1 knockdown. Given that E2F1 knockdown did affect RpS6 phosphorylation in cells with cyclin D and Cdk4

Figure 4. E2F1 Transcription Factor Positively Regulates TORC1 Downstream of Smoothened (A) E2F1 RNAi was expressed in the eye disc using DE-GAL4, and RpS6 phosphorylation (red) was examined by immunostaining. (B) Rbf1 RNAi was expressed with Dicer2 in the eye disc using DE-GAL4, and phosphorylated RpS6 (red) was examined. (C) Smo mutant clones (absence of RFP) were generated in the eye disc in which a GFP-fused PCNA was expressed via endogenous PCNA promoter. (D) DE-GAL4 was used to express Rbf1 RNAi in the eye disc in which Smo mutant clones (absence of GFP) were generated. The eye disc was immunostained with the anti-pS6 antibody (red). (E and F) RNAi for cyclin D (E) or Cdk4 (F) was expressed with RFP (not shown in the figure) by DE-GAL4, and RpS6 phosphorylation (red) was examined. (G) Cyclin D and Cdk4 were expressed with RFP (not shown in the figure) by DE-GAL4, and the eye disc was immunostained with the anti-pS6 antibody (red). (H) E2F1 RNAi was expressed with cyclin D and Cdk4 by DE-GAL4, and RpS6 phosphorylation (red) was examined. EGFP was expressed together as a marker (not shown in the figure). (A, B, and D–H) The dorsal regions in which transgenes were expressed by DE-GAL4 are above the dotted lines. (A, B, and E–H) DNA was marked by Hoechst (blue). (C and D) The boxed areas are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. Scale bars, 30 mm.

expression, it can be concluded that the cyclin D/Cdk4 complex is not the sole mediator of Hh-E2F1 signaling in regulating TORC1 activity in the eye disc. Genetic Epistasis of Hh-E2F1 Signaling to the TSC Complex and Insulin/Amino Acid Signaling Next, genetic epistasis among Hh signaling, E2F1, the TSC complex, and Rheb was investigated. Interestingly, when we generated Rheb-overexpressing Smo mutant cells by MARCM, Rheb overexpression significantly increased TORC1 activity in the clones (Figure 5A). Moreover, when Smo mutant clones were generated in the eye disc in which tsc2 RNAi was expressed in the dorsal region, Smo mutation could not inhibit the TORC1 activity induced by tsc2 RNAi expression (Figure 5B). Similarly, expression of E2F1 RNAi did not suppress TORC1 activity in tsc2 RNAiexpressed cells (Figures 5C and 5D). These results demonstrated that Hh signaling and E2F1 are not downstream, but upstream of, or in parallel with, the TSC complex in TORC1 regulation. Moreover, we questioned whether Hh signaling and E2F1 regulate TORC1 through insulin or amino acid signaling. First, we examined the insulin signaling activity in Smo mutant cells, and observed that Smo mutation did not affect Akt phosphorylation (Figure 5E). In addition, E2F1 or Rbf1 RNAi expression using DE-GAL4 did not alter Akt phosphorylation (Figures 5F and 5G,

respectively). Furthermore, we tested whether elevating insulin signaling activity could restore the TORC1 activity defect in Smo mutant cells by manipulating PTEN. When PTEN mutant clones were generated there was an increased TORC1 activity near the SMW (Figure 5H), whereas when Smo and PTEN double-mutant clones were generated the cells in the clones elicited an absence of TORC1 activity (Figure 5I). It can be interpreted that hyper-activation of insulin signaling had no effect on restoring the TORC1 activity in the Smo mutant cells. Overall, these results indicated that Hh signaling and E2F1 do not affect insulin signaling to activate TORC1. We also tested whether Hh-E2F1 signaling regulates TORC1 through amino acid signaling. In mammalian cells and Drosophila, expression of RagA mutant (RagAQ61L), which is restricted to the GTP-bound conformation, constitutively activates TORC1 even in the absence of amino acids (Kim et al., 2008; Sancak et al., 2008). Clonal expression of RagAQ61L resulted in activated TORC1 near the SMW (Figure S2C), which is similar to PTEN mutation (Figure 5H). However, the suppressed TORC1 activity in the cells with E2F1 knockdown was not restored when RagAQ61L was expressed together (Figure 5J), suggesting that the amino acid signaling does not mediate the function of E2F1 in TORC1 regulation. Based on these results, we concluded that Hh signaling and E2F1 regulate TORC1 upstream of, or in parallel with, the TSC Developmental Cell 42, 363–375, August 21, 2017 369

Figure 5. Genetic Epistasis of Hh-E2F1 Signaling to the TSC Complex and Insulin/ Amino Acid Signaling (A) Smo mutant clones in which Rheb was overexpressed with GFP were generated in the eye disc, and RpS6 phosphorylation (red) was examined. (B) Smo mutant clones (GFP-negative) were generated in the eye disc in which tsc2 RNAi and Dicer2 were expressed using DE-GAL4, and the eye disc was stained with the anti-pS6 antibody (red). (C) tsc2 RNAi was expressed with Dicer2 using DE-GAL4, and RpS6 phosphorylation (green) was examined. (D) E2F1 RNAi and tsc2 RNAi were simultaneously expressed with Dicer2 using DE-GAL4, and phosphorylated RpS6 (green) was immunostained. (E) The eye disc in which Smo mutant clones (GFPnegative) was generated and immunostained with the anti-pAkt antibody (red). (F and G) E2F1 RNAi (F) or Rbf1 RNAi (G) was expressed with Dicer2 by DE-GAL4, and the eye disc was immunostained with the anti-pAkt antibody (green). (H and I) PTEN mutant clones (absence of GFP) (H) or Smo and PTEN double-mutant clones (absence of GFP) (I) were generated and immunostained with anti-pS6 antibody (red). (J) RagACA and E2F1 RNAi were expressed with Dicer2 by DE-GAL4 and phosphorylated RpS6 (red) was immunostained. (B–D, F, G, and J) The dorsal regions in which transgenes were expressed by DE-GAL4 are above the dotted lines. (C, D, F, G, and J) DNA was marked by Hoechst (blue). (A, B, E, H, and I) The boxed areas are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. Scale bars, 30 mm. See also Figure S2.

complex, but the regulation is not through insulin or amino acid signaling. TORC1 Is Required for the G1/S Transition in the SMW Downstream of Hh Signaling Subsequently, the developmental role of TORC1 in the SMW was investigated. Because TORC1 was activated selectively in the S-phase cells of the SMW, the function of TORC1 in the G1/S transition of these cells was examined in detail. To monitor the S-phase cells, we incorporated bromodeoxyuridine (BrdU), a thymidine analog, into the eye disc and immunostained using an antibody against BrdU. Interestingly, TOR mutant cells entered S phase in the more posterior region than did the control cells in the SMW, indicating that their G1/S transition was delayed, and that TORC1 was required for the properly synchronized G1/S transition in the SMW (Figure 6A). In addition, we as370 Developmental Cell 42, 363–375, August 21, 2017

sessed whether S6K and 4E-BP, the two direct targets of TORC1 phosphorylation, mediate the G1/S transition downstream of TORC1. However, S6K mutation did not delay the transition of the cells in the SMW (Figure 6B), and 4E-BP mutation did not rescue the G1/S transition timing defect induced by TOR mutation (Figure 6C). Furthermore, by expressing wild-type 4E-BP in the cells with S6K mutation and monitoring the S-phase cells (Figure 6D), we abrogated the possibility of S6K and 4E-BP redundantly functioning in the G1/S transition. These results suggested that neither S6K nor 4E-BP is a critical mediator of TORC1’s role in promoting the G1/S transition. Because a previous study proposed that the two cyclins, cyclin E and A, regulate the G1/S transition of the SMW (Baonza and Freeman, 2005), we generated TOR mutant clones and immunostained for these cyclins to examine the involvement of TORC1 in their regulation. As expected, the protein levels of both cyclins were decreased in TOR mutant clones (Figures 6E and 6F). The reduction in protein levels was not the result of a decrease in general mRNA translation caused by the loss of TORC1

Figure 6. TORC1 Promotes the G1/S Transition of the Cells in the SMW Downstream of Hh Signaling (A–C) BrdU was incorporated in the eye disc in which mutant clones of TOR (A), S6K (B), or both TOR and 4E-BP (C) (GFP-negative) were generated, and the eye disc was immunostained with the anti-BrdU antibody (red). (D) 4E-BP-expressing S6K mutant clones, which were marked by GFP, were generated and BrdU incorporation (red) was examined. (E and F) The eye disc in which TOR mutant clones (GFP-negative) were generated was immunostained with the anti-cyclin E (E) or -cyclin A (F) antibodies (red). (G) Smo mutant clones (GFP-negative) were generated in the eye disc and the larvae were fed with BrdU for 2 hr. The eye disc was fixed and immunostained with the anti-BrdU (magenta) and -pS6 (red) antibodies. Arrows indicate Smo mutant cells at the clone boundary which entered S phase. Note that the TORC1 activity is still inhibited in these cells (no red signal in GFP-negative clones regardless of BrdU incorporation), as it is in other Smo mutant cells. (H) tsc2 RNAi was expressed with Dicer2 using DE-GAL4, and Smo mutant clones (absence of GFP) were generated in the eye disc. BrdU was incorporated in the eye disc, and the eye disc was immunostained with the anti-BrdU antibody (red). The dorsal region in which transgenes was expressed by DE-GAL4 is above the dotted line. (A–H) The boxed areas are magnified in the rightmost panels, where the boundary of the clones is indicated by white dotted lines. Scale bars, 10 mm (A–C, E, and F) or 30 mm (D, G, and H). See also Figure S6.

activity, because the protein levels of other cell-cycle regulators, such as Cdk1 and Dacapo (Drosophila p21/27 Cdk inhibitor), were not changed in TOR mutant cells (data not shown). Moreover, it is known that cyclin E hypomorphic mutant cells show decreased S phase progression in the SMW (Secombe et al., 1998), and we observed that cyclin A mutant cells did not enter the S phase in the SMW (Figure S6). These results suggest that the both cyclins are indispensable downstream effectors of TORC1 in the G1/S transition of the SMW. Next, we questioned whether Hh signaling is responsible for the cells to enter S phase in time through regulating TORC1. We found that Smo mutant cells in the SMW did not enter S phase (Figure 6G). Moreover, we observed that some mutant cells at the boundary of the clones entered S phase (indicated by arrows in Figure 6G), which was consistently reported by a previous publication (discussed below) (Firth and Baker, 2005). Based on our results, we hypothesized that the S phase entry defect of Smo mutant cells was caused by TORC1 inactivation. To test this hypothesis, we expressed tsc2 RNAi transgene in the dorsal region of the eye disc and generated Smo mutant clones. Surprisingly, tsc2 downregulation restored the defect in S phase entry of Smo mutant cells (Figure 6H). Based on these results, we

concluded that TORC1 is an important downstream mediator of Hh signaling in regulation of the G1/S transition in the SMW. Hh-E2F1-Cyclin D/Cdk4-mTORC1 Signaling Axis Is Conserved in Mammalian System Evolutionary conservation of TORC1 regulation by Hh-E2F1cyclin D/Cdk4 signaling was investigated in mammalian cells. For this purpose, we treated cells with sonic Hedgehog (Shh), one of the mammalian hedgehogs. Amino acid starvation dramatically decreased mTORC1 activity as shown by RpS6 phosphorylation (Figure 7A). However, in the Shh-treated cells, amino acid starvation could not decrease mTORC1 activity (Figure 7A). Purmorphamine (Pur), an agonist of Smo, was also used to pharmacologically activate Hh signaling pathway. We first confirmed that Pur activates Hh signaling in HeLa cells by observing induction of Gli1 mRNA upon treatment (Figure S7A). Consistent with the Shh experiments, we found that treatment of Pur made mTORC1 activity resistant to amino acid starvation (Figure 7B). Next, we tested whether E2F1, CDK4, and cyclin D mediate TORC1 regulation of Hh signaling in mammals. Drosophila E2F1, Cdk4, and cyclin D are conserved in the mammalian system as E2F1/2/3, CDK4/6 and cyclin D1/2/3, respectively. We silenced Developmental Cell 42, 363–375, August 21, 2017 371

Figure 7. Hh-E2F1-cyclin D/Cdk4-mTORC1 Signaling Axis Is Conserved in Mammalian Cells (A) HeLa cells were treated with recombinant sonic Hedgehog (Shh, 4 mg/mL) or starved with amino-acid-free DMEM (stv) for 2 hr and the lysate samples were immunoblotted with anti-pS6, -S6, and -b-tubulin antibodies. (B) HeLa cells were treated with purmorphamine (Pur, 20 mM) and/or starved with amino-acid-free DMEM. The lysate samples were immunoblotted with the same antibodies as in (A). (C–E) E2F1/2/3 (C), CDK4/6 (D), or cyclin D1/2/3 (E) knockdown cells were treated with Pur and/or starved with amino-acid-free DMEM. The lysate samples were immunoblotted with the same antibodies as in (A). (F) Comparison of cyclin D1 mRNA expression levels in E2F1/2/3 knockdown HeLa cells with or without Pur treatment (n = 3). Significance was determined by Student’s two-tailed t test (***p < 0.001; ns, not significant). Error bars indicate SD. (G) Cyclin D1-overexpressed and/or E2F1/2/3 knockdown cells were treated with Pur and/or starved with amino-acid-free DMEM. The lysate samples were immunoblotted with anti-pS6, -S6, -HA, and -b-tubulin antibodies. (B–G) The indicated cells were treated with 20 mM Pur for 72 hr. See also Figure S7.

each gene set at the same time (Figures S7B–S7D) and observed mTORC1 activity by examining phosphorylated RpS6. Interestingly, the starvation resistance of mTORC1 activity upon Pur treatment was abolished when any one set of the genes among E2F1/ 2/3, CDK4/6, and cyclin D1/2/3 was silenced (Figures 7C–7E, respectively). In the Drosophila eye disc, we observed that overexpression of cyclin D and Cdk4 partially restored TORC1 activity in E2F1 knockdown cells. Therefore, we examined whether Hh signaling regulates cyclin D transcript level through E2F1/2/3 in mammalian cells. Expectedly, Pur treatment increased cyclin D1, 2, and 3 mRNA levels, and simultaneous knock down of E2F1/2/3 blocked this Pur effect (Figures 7F, S7E, and S7F, respectively). These results indicated that Hh signaling positively regulates cyclin D mRNA transcription in an E2F1/2/3-dependent 372 Developmental Cell 42, 363–375, August 21, 2017

manner in mammalian cells. Finally, cyclin D was restored in the E2F1/2/3 knockdown cells to verify if this is sufficient for the rescue of the mTORC1 activity. As predicted, ectopic expression of cyclin D1 made mTORC1 activity resistant to amino acid starvation upon Pur treatment, even when E2F1/2/3 were downregulated (Figure 7G), suggesting that Hh-E2F1 signaling positively regulates mTORC1 activity through cyclin D. Collectively, these results suggest that the Hh-E2F1-cyclin D/Cdk4-mTORC1 axis is conserved in mammalian system and Drosophila. DISCUSSION The regulatory mechanism of TORC1 has been highly investigated at the cellular level using mammalian and yeast cells,

whereas TORC1 regulation in multicellular tissues remains largely unknown. To investigate this issue, we chose the Drosophila eye, an excellent model system for investigating how signaling networks are organized during animal development. By examining the phosphorylation of RpS6, a downstream target of TORC1, we found that TORC1 was selectively activated in the S-phase cells of the SMW. In agreement with mammalian cell-based studies, insulin signaling was required for the TORC1 activity, but, unexpectedly, insulin signaling was uniformly activated in the whole eye disc and did not match with the activity pattern of TORC1 in the SMW. This implies that TORC1 is regulated by other signaling molecules in addition to insulin signaling molecules in spatially specific cells during Drosophila eye development. Hh signaling induces patterned gene expression during development of many organisms (reviewed in Jiang and Hui, 2008). In this study, we demonstrated that Hh signaling activates TORC1 in the S-phase cells of the SMW. Moreover, we proposed the molecular mechanism of how Hh signaling activates TORC1 in the eye disc, by elucidating that Hh signaling activates E2F1 transcription factor in the S-phase cells of the SMW to induce TORC1 signaling. Furthermore, we searched for the downstream component of E2F1 using various genetic approaches. As a result, we propose the cyclin D/Cdk4 complex as a downstream mediator of Hh-E2F1 signaling on TORC1. We further addressed that the Hh-E2F1-cyclin D/Cdk4TORC1 signaling axis is also conserved in the mammalian system. We found that activation of Hh signaling leads to a retained mTORC1 activity upon starvation, in which E2F1/2/3, CDK4/6, and cyclin D1/2/3 are responsible (Figures 7A–7E). Moreover, our mammalian cell-based study provides detailed molecular mechanisms of the signaling axis. Activation of Hh signaling leads to increased cyclin D1/2/3 transcript level, which is dependent on E2F1/2/3 (Figures 7F, S7E, and S7F, respectively), suggesting that E2F1/2/3 regulate cyclin D1/2/3 transcription. A previous study reported that cyclin D interacts with TSC2 and that the expression of cyclin D1 and CDK4/6 increases phosphorylation of TSC2 (Zacharek et al., 2005), suggesting that the cyclin D/Cdk4 complex could directly phosphorylate TSC2 to regulate mTORC1 activity. Interestingly, TSC2 subcellular localization is changed upon phosphorylation, and Akt phosphorylates TSC2 to inhibit its lysosomal localization in HeLa cells when insulin signaling is activated (Menon et al., 2014). Although whether insulin signaling affects the stability of TSC2 is controversial (Dong and Pan, 2004; Inoki et al., 2002; Manning et al., 2002; Potter et al., 2002), it has been reported that Akt-mediated phosphorylation results in degradation of TSC2 (Plas and Thompson, 2003). Therefore, it would be interesting to investigate whether the cyclin D and CDK4/6 complex regulates mTORC1 by modulating the subcellular localization and/or stability of the TSC complex through direct phosphorylation. Interestingly, an accompanying paper by the Teleman’s group in this issue of Developmental Cell dealt with the issue, providing evidence that the cyclin D/Cdk4 complex indeed phosphorylates TSC2 in Drosophila tissue (Romero-Pozuelo et al., 2017). It has previously been reported that N signaling promotes G1/S cell-cycle transition in the SMW downstream of Hh signaling (Baonza and Freeman, 2005; Firth and Baker, 2005). Here we

investigated the regulation between Hh, N, and TORC1 signaling using various genetic experiments. TORC1 activity was not reduced in N mutant or Dl and Serrate double-mutant cells, whereas TOR mutant cells showed normal protein level of Dl, which is downregulated in Smo mutant cells. These results indicated that N and TORC1 signaling pathways do not regulate each other and are independent downstream targets of Hh signaling. Moreover, we observed that cell-cycle progression into S phase did occur in some Smo mutant cells in contact with control clones (i.e., cells at the boundary of Smo mutant clone; arrows in Figure 6G), which was also previously reported by Baker’s group (Firth and Baker, 2005). Because Smo mutant cells adjacent to control cells can induce N signaling non-autonomously by neighboring control cells, this result suggests that Hh signaling promotes G1/S cell-cycle transition, not only through TORC1 signaling, but also through N signaling. Importantly, the S-phase-progressing Smo mutant cells in the clone boundary still showed no S6 phosphorylation, further supporting that N signaling does not mediate TORC1 activation by Hh signaling in the SMW (Figure 6G). Overall, these results indicate that Hh signaling promotes G1/S transition through two independent downstream signaling pathways, N and TORC1. During Drosophila eye development, precursor cells are synchronously arrested at the G1 phase, and R8, R2/5, and R3/4 cells begin to sequentially differentiate near the MF. In contrast, R1/6, R7, and cone cells differentiate after the SMW (reviewed in Baker, 2001). Previously, Bateman and McNeill reported that tsc2 and TOR mutant clones exhibited precocious and delayed differentiation of R1/6, R7, and cone cells, respectively, but that the differentiation of R8, R2/5, and R3/4 was not affected in these mutant clones; these results indicate that TORC1 specifically controls the timing of R1/6, R7, and cone cell differentiation in the Drosophila eye disc (Bateman and McNeill, 2004; McNeill et al., 2008). Here we observed that TOR mutant cells showed a delayed G1/S phase transition in the SMW. Based on our and previous results, we postulated that the delayed timing of photoreceptor neuron differentiation in TOR mutant cells was caused by a slowed G1/S transition in the SMW. In support of this idea, we observed that Rheb overexpression shortened the duration of the S phase of the SMW, suggesting that the precocious differentiation of tsc2 mutant cells was caused by a decrease in the duration of the S phase (data not shown). Notably, it was reported that the size of nucleolus in the eye disc varied, and that cells with cell-cycle progression have larger ones (Baker, 2013). Since ribosome biogenesis is one of the multiple pathways TORC1 is known to regulate (Iadevaia et al., 2014), there is a possibility that ribosome biogenesis is involved in the cell-cycle regulation by TORC1. Taken together, these results suggested that TORC1 regulates the differentiation timing of photoreceptor neurons by controlling cell-cycle progression in the SMW. Collectively, our data consistently suggest that TORC1 can be spatially controlled by delicate convergence of Hh signaling and previously known cell growth signaling cues, such as nutrient information and growth hormones during Drosophila tissue and organ development. Further studies focused on more precise molecular mechanism concerning the regulation of TORC1 during animal development would be meaningful and interesting to pursue. Developmental Cell 42, 363–375, August 21, 2017 373

STAR+METHODS

Baker, N.E. (2001). Cell proliferation, survival, and death in the Drosophila eye. Semin. Cell Dev. Biol. 12, 499–507.

Detailed methods are provided in the online version of this paper and include the following:

Baker, N.E. (2007). Patterning signals and proliferation in Drosophila imaginal discs. Curr. Opin. Genet. Dev. 17, 287–293.

d d d

d

d

KEY RESOURCES TABLE CONTACT FOR REAGENT AND RESOURCE SHARING EXPERIMENTAL MODEL AND SUBJECT DETAILS B Drosophila Genetics B Human Cell Culture B Drosophila Cell Culture METHOD DETAILS B Generation of Anti-pS6 Antibody B Immunostaining B Generation of Tub-hS6 Fly B Clone Generation B BrdU Incorporation B Drosophila S2 Cell Culture and dsRNA Bathing B Immunoblotting B Mammalian Cell Culture and Transfection B Quantitative Real-Time PCR QUANTIFICATION AND STATISTICAL ANALYSIS

SUPPLEMENTAL INFORMATION Supplemental Information includes seven figures and three tables and can be found with this article online at http://dx.doi.org/10.1016/j.devcel.2017. 07.020. AUTHOR CONTRIBUTIONS Conceptualization, W.K. and J.C.; Methodology, W.K., Y.-G.J., and J.Y.; Investigation, W.K., Y.-G.J., and J.Y.; Writing – Original Draft, W.K., Y.-G.J., and J.Y.; Writing – Review & Editing, W.K., Y.-G.J., J.Y., and J.C.; Supervision, J.C.; Funding Acquisition, J.C. ACKNOWLEDGMENTS We would like to thank Drs. Clive Wilson, Shinya Yamamoto, Marco Milan, Ernst Hafen, Thomas Neufeld, Mary Lilly, Kwang-Wook Choi, Jin Jiang, Kenneth Irvine, and Bruce Edgar for fly stocks, Dr. Aurelio Teleman for anti-Drosophila S6K antibody and sharing the unpublished results, and Dr. Suk-Chul Bae for human cyclin D1 plasmid. We truly appreciate Drs. Kwang-Wook Choi and Eunjoo Cho for helpful discussion. We are grateful to the Bloomington Stock Center, National Institute of Genetics, and Vienna Drosophila Resource Center. We also would like to thank all members of the Chung lab for helpful insights and discussion. This research was supported by National Creative Research Initiatives Program Grant 2010-0018291 from the Korean Ministry of Science, ICT and Future Planning. Y-G.J., J.Y., and J.C. were supported by the BK21 Plus Program from Korean Ministry of Education. Received: August 29, 2016 Revised: June 17, 2017 Accepted: July 23, 2017 Published: August 21, 2017 REFERENCES Anjum, R., and Blenis, J. (2008). The RSK family of kinases: emerging roles in cellular signalling. Nat. Rev. Mol. Cell Biol. 9, 747–758. Aza-Blanc, P., Ramirez-Weber, F.A., Laget, M.P., Schwartz, C., and Kornberg, T.B. (1997). Proteolysis that is inhibited by hedgehog targets Cubitus interruptus protein to the nucleus and converts it to a repressor. Cell 89, 1043–1053.

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Developmental Cell 42, 363–375, August 21, 2017 375

STAR+METHODS KEY RESOURCES TABLE

REAGENT or RESOURCE

SOURCE

IDENTIFIER

Rabbit anti-phosphorylated ribosomal protein S6 antibody

This study

N/A

Rabbit anti-Phospho-Akt (Ser473) antibody

Cell Signaling Technology

Cat# 4060; RRID: AB_2315049

Mouse anti-BrdU antibody

BD Biosciences

Cat# 555627; RRID: AB_395993

Rabbit anti-S6 ribosomal protein antibody


Cat# 2217; RRID: AB_331355

Rabbit anti-Phospho-S6 Ribosomal Protein (Ser235/236) Antibody


Cat# 2211; RRID: AB_331679

Antibodies

Rat anti-cubitus interruptus antibody

DSHB

Cat# 2A1; RRID: AB_2109711

Mouse anti-Delta antibody

DSHB

Cat# c594.9b; RRID: AB_528194

Mouse anti-Bruchpilot antibody

DSHB

Cat# nc82; RRID: AB_2314865

Mouse anti-cyclin A antibody

DSHB

Cat# a12; RRID: AB_2090494

Rabbit anti-Drosophila Cyclin E (d-300)

Santa Cruz Biotechnology

Cat# sc-33748; RRID: AB_638863

Rabbit anti-Phospho-Drosophila p70 S6 Kinase (Thr398) Antibody


Cat# 9209S; RRID: AB_2269804

Guinea pig anti-S6 Kinase antibody

Hahn et al., 2010

N/A

Mouse anti-S6 Ribosomal Protein (54D2) antibody


Cat# 2317; RRID: AB_2238583

Rabbit anti-Phospho-S6 Ribosomal Protein (Ser240/244) (D68F8) XP antibody


Cat# 5364; RRID: AB_10694233

Mouse anti-HA-tag antibody

MBL International

Cat# M180-3; RRID: AB_10951811

Mouse anti-b-tubulin antibody

DSHB

Cat# E7; RRID: AB_528499

5-Bromo-2’-deoxyuridine

Roche

10 280879 001

RQ1 RNase-Free DNase

Promega

M6101

Schneider’s Insect Medium w/L-Glutamine w/o Calcium Chloride, Sodium Bicarbonate, Amino Acids, Yeast

US Biological

S0100-01

Chemicals, Peptides, and Recombinant Proteins

Schneider’s Drosophila Medium

Gibco

21720-024

DMEM

Welgene

LM 001-05

Fetal bovine serum

Gibco

16000-044

Lipofectamine LTX Reagent

Invitrogen

15338-100

RNAiMAX reagent

Invitrogen

13778150

TOPreal qPCR 2X PreMIX

Enzynomics

RT500

Insulin

Roche

11376497001

Purmorphamine

Sigma

SML0868

Recombinant human sonic hedgehog

R&D SYSTEMS

1845-SH-025

Shields and Sang M3 media

Sigma

S8398

D. melanogaster: Cell line S2-DRSC

DGRC

Cat# 181, RRID:CVCL_Z992

HeLa cell line

Laboratory of Dr. Chin Ha Chung

N/A

D. melanogaster: tub-hS6

This study

N/A

D. melanogaster: TORDP FRT40A

Bloomington Drosophila Stock Center

BDSC: 7014; FlyBase: FBst0007014

D. melanogaster: DE-GAL4



D. melanogaster: 4E-BPk13517



D. melanogaster: UAS-Raptor RNAi



Experimental Models: Cell Lines

Experimental Models: Organisms/Strains

(Continued on next page)

e1 Developmental Cell 42, 363–375.e1–e4, August 21, 2017

Continued REAGENT or RESOURCE

SOURCE

IDENTIFIER

D. melanogaster: RhebAV4



D. melanogaster: UAS-InR



D. melanogaster: UAS-PI3KD954A



D. melanogaster: Smo3



D. melanogaster: RagCEY11726



D. melanogaster: FRT82B DlRevF10, SerRX82



D. melanogaster: cyclin AC8LR1



D. melanogaster: UAS-Ci RNAi



D. melanogaster: UAS-E2F1 RNAi



D. melanogaster: UAS-Rbf1 RNAi



D. melanogaster: UAS-cycD RNAi



D. melanogaster: UAS-Cdk4 RNAi



D. melanogaster: y w hs-FLP tub-GAL4 UAS-GFP; RpS174 tub-GAL80 FRT80B



D. melanogaster: UAS-4E-BP



D. melanogaster: dimm-GAL4



D. melanogaster: Pdf-GAL4



D. melanogaster: PCNA-GFP



D. melanogaster: UAS-tsc2 RNAi

Vienna Drosophila Resource Center

VDRC: 6313; FlyBase: FBst0470275

D. melanogaster: UAS-p18 RNAi



D. melanogaster: UAS-HBXIP RNAi



D. melanogaster: UAS-mio RNAi


VDRC: 108609; FlyBase: Fbst0480419

D. melanogaster: UAS-Dicer2


VDRC: 60009; FlyBase: N/A

D. melanogaster: UAS-Akt RNAi

National Institute of Genetics

NIG: 4006R-3; FlyBase: N/A

K1409A

D. melanogaster: tsc2192 FRT80B

(Ito and Rubin, 1999)

N/A

D. melanogaster: PTEN3 FRT40A

(Goberdhan et al., 1999)

N/A

D. melanogaster: N54/9 FRT19A

(Yamamoto et al., 2012)

N/A

D. melanogaster: UAS-Nintra

(Milan and Cohen, 2003)

N/A

D. melanogaster: S6Kl-1 FRT80B

(Montagne et al., 1999)

N/A

D. melanogaster: UAS-RagAQ61L

(Kim et al., 2008)

N/A

D. melanogaster: UAS-RagAT16N

(Kim et al., 2008)

N/A

D. melanogaster: seh1D15

(Senger et al., 2011)

N/A

D. melanogaster: UAS-CiR

(Wang and Jiang, 2004)

N/A

D. melanogaster: UAS-cyclin D UAS-Cdk4

(Datar et al., 2000)

N/A

dsRNA synthesis primer

This paper

Table S1

qRT-PCR primer

This paper

Table S2

Oligonucleotides

Recombinant DNA pHA

(Lee et al., 2013)

N/A

pHA-cyclin D1

(Lee et al., 2013)

N/A

pCaSpeR4 Tubp-Gal80

Addgene

17748

Graphpad

N/A

Software and Algorithms Prism 7

CONTACT FOR REAGENT AND RESOURCE SHARING Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Jongkyeong Chung ([email protected]). Developmental Cell 42, 363–375.e1–e4, August 21, 2017 e2

EXPERIMENTAL MODEL AND SUBJECT DETAILS Drosophila Genetics Species: Drosophila melanogaster. All flies were grown on food containing approximately 35 g cornmeal, 70 g dextrose, 5 g agar, 50 g dry active yeast, 4.6 ml propionic acid, and 7.3 ml Tegosept (100 g/l in ethanol) per liter at 25 C. w1118 strain was used as wild type control. The following fly lines were obtained from the Bloomington Stock Center (Bloomington, IN): TORDP FRT40A (BL7014), DEGAL4 (BL29650), 4E-BPk13517 (BL9558), UAS-Raptor RNAi (BL31529), RhebAV4 (BL9690), UAS-InRK1409A (BL8253), UAS-PI3KD954A (BL25918), Smo3 (BL3277), RagCEY11726 (BL21364), FRT82B DlRevF10, SerRX82/TM6B (BL6300), cyclin AC8LR1 (BL6627), UAS-Ci RNAi (BL28984), UAS-E2F1 RNAi (BL27564), UAS-Rbf1 RNAi (BL36744), UAS-cycD RNAi (BL33653), UAS-Cdk4 RNAi (BL27714), y w hs-FLP tub-GAL4 UAS-GFP; RpS174 tub-GAL80 FRT80B/TM6B (BL42732), UAS-4E-BP (BL9147), dimm-GAL4 (BL25373), PdfGAL4 (BL6900) and PCNA-GFP (BL25749). UAS-tsc2 RNAi (v6313), UAS-p18 RNAi (v35614), UAS-HBXIP RNAi (v31814), UASmio RNAi (v108609) and UAS-Dicer2 (v60009) were obtained from Vienna Drosophila Resource Center (Vienna, Austria). UAS-Akt RNAi (4006R-3) was obtained from National Institute of Genetics (Kyoto, Japan). tsc2192 FRT80B was obtained from Dr. Gerald Rubin. PTEN3 FRT40A was obtained from Dr. Clive Wilson. N54/9 FRT19A was provided from Dr. Shinya Yamamoto. UAS-Nintra was provided from Dr. Marco Milan. S6Kl-1 FRT80B was a gift from Dr. Ernst Hafen. UAS-RagAQ61L and UAS-RagAT16N were gifted by Dr. Thomas Neufeld. seh1D15 was provided by Dr. Mary Lilly. UAS-TSC1 and UAS-TSC2 were obtained from Dr. Kwang-Wook Choi. UAS-CiR was kindly provided by Dr. Jin Jiang. y w hs-FLP tub-GAL4 UAS-GFP/FM7; tub-GAL80 FRT40A/CyO was provided by Dr. Kenneth Irvine. UAS-cyclin D and UAS-Cdk4 were provided by Dr. Bruce Edgar. For immunostaining of the eye disc and larval brain, wandering larvae were dissected. For immunostaining of fat body, third instar larvae were dissected. For immunostaining of adult brain or ovary, age-matched male or female flies were dissected, respectively. Third instar larvae or age-matched adult male flies were incubated in 1% agarose in phosphate-buffered saline (PBS) for nutrient starvation. Human Cell Culture HeLa cells were cultured in DMEM (Welgene) supplemented with 10% fetal bovine serum (FBS) (Invitrogen) at 37 C in a humidified atmosphere of 5% CO2. Drosophila Cell Culture Drosophila melanogaster S2 cells were maintained in Shields and Sang M3 media (Sigma, S8398) supplemented with 10% FBS and Penicillin-Streptomycin. METHOD DETAILS Generation of Anti-pS6 Antibody A phosphospecific antibody against Drosophila S6 was generated by immunizing rabbits with the phospho-peptide, EAKRRR[pS]A[pS]IRE. Immunostaining The eye discs and brains of wandering larvae, the fat body of third instar larvae, and the ovaries of adult flies were dissected in PBS and then fixed with 4% paraformaldehyde for 20 minutes at room temperature. For immunostaining of the adult brains, flies were fixed with 4% paraformaldehyde in PBS containing 0.1% Triton X-100 (PBST) for 4 hours at room temperature, and the adult brains were dissected in PBS. The samples were washed twice with PBST for 10 minutes and permeabilized in PBS with 0.5% Triton X-100 for 5 minutes at room temperature. The samples were washed once with PBST for 10 minutes and blocked with PBST containing 3% BSA. Primary antibodies were applied, and the samples were incubated at 4 C for overnight on a nutator. The samples were washed three times for 10 minutes with PBST and incubated at room temperature for 2 hours with secondary antibodies and Hoechst 33538 dye. The samples were washed three times for 10 minutes with PBST and mounted in 80% glycerol-PBS solution. The following primary antibodies were used in this study: anti-pS6 polyclonal rabbit antibody (1:400), anti-phosphospecific Akt rabbit antibody (1:100, Cell Signaling Technology (CST), 4060), anti-BrdU mouse antibody (1:100, BD Biosciences, 555627), antihuman RpS6 rabbit antibody (1:100, CST, 2217), anti-phosphospecific human RpS6 rabbit antibody (1:100, CST, 2211), anti-Ci mouse antibody (1:10, Developmental Studies Hybridoma Bank (DSHB), 2A1), anti-Delta mouse antibody (1:2,000, DSHB, C594.9B), anti-Bruchpilot mouse antibody (1:100, DSHB, nc82), anti-cyclin A mouse antibody (1:10, DSHB, A12), and anti-cyclin E rabbit antibody (1:100, Santa Cruz, sc-33748). Generation of Tub-hS6 Fly To generate tub-hS6 flies, GAL80 cDNA was replaced by the human RpS6 ORF in pCasper4 tub-GAL80 (Addgene, 17748). The pCasper4 tub-hS6 plasmid was microinjected into w1118 embryos.

e3 Developmental Cell 42, 363–375.e1–e4, August 21, 2017

Clone Generation In the eye disc, mitotic clones were generated via FRT recombination by driving FLP expression using the eyeless (ey) or heat shock protein (hs) promoter. hs-FLP was also used in the MARCM analysis. To induce mitotic clones by heat shock, embryos were collected for 24 hours, reared for 2 days, and then incubated for 1 hour at 37 C. After 3 days, wandering larvae were dissected and analyzed. BrdU Incorporation Wandering larvae were dissected on Shields and Sang M3 media (Sigma, S8398) and then incubated in M3 medium containing 0.1 mg/ml BrdU (Roche, 10 280879 001) for 30 minutes at room temperature on a nutator. After briefly washing the larvae three times with PBS, they were fixed in 4% paraformaldehyde for 20 minutes. The samples were washed three times in PBST and then incubated with 5 units of DNase I (Promega, M6101) in 200 ml of DNase I buffer and PBS for 1.5 hours at 37 C. The larvae were washed three times with PBST for 10 minutes each and then incubated with an anti-BrdU antibody overnight in blocking solution at 4 C. For an experiment when we wanted to fix the larva right after the dissection (Figure 6G), BrdU was fed for 2 hours before dissection. Forty ml of 10 mg/ml BrdU stock solution was mixed to 2 ml of melted Kankel-White medium for the food preparation. Drosophila S2 Cell Culture and dsRNA Bathing Drosophila melanogaster S2 cells were maintained in Shields and Sang M3 media (Sigma, S8398) supplemented with 10% FBS and Penicillin-Streptomycin. Double-stranded RNA (dsRNA) was synthesized and treated as previously described (Kim et al., 2015). For amino acid-starvation, S2 cells were washed once in PBS, and then incubated in Schneider media without amino acids (US Biological, S0100-01) for 1 hour. Amino acids were replenished by incubating cells in Schneider media (Gibco, 21720-024) for 20 minutes. For dsRNA synthesis, we used the primers in Table S1. Immunoblotting S2 cells were collected and lysed with lysis buffer (20 mM Tris-HCl pH 7.5, 100 mM sodium chloride, 1 mM ethylenediaminetetraacetic acid, 2 mM ethyleneglycoltetraacetic acid, 1 mM sodium vanadate, 50 mM b-glycerophosphate, 50 mM sodium fluoride, 0.1% Triton X-100, 1 mM dithiothreitol, 1 mg/ml leupeptin, and 1 mM phenylmethylsulfonyl fluoride). HeLa cells were lysed with lysis buffer (Kim et al., 2016). The lysates were clarified by centrifugation at 13,200 rpm for 15 minutes at 4 C. The protein concentration in the lysates was measured by BCA assay, and lysate samples of equivalent concentration and volume were prepared. Sample buffer was added to the lysates and the solution was boiled for 5 minutes at 95 C to run SDS-PAGE. The following primary antibodies were used for immunoblot analyses: anti-pS6 rabbit antibody (1:5,000), anti-phosphospecific S6K (pS6K) rabbit antibody (1:1,000, CST, 9209), anti-S6K guinea pig antibody (1:1,000, a gift from Dr. Aurelio Teleman), anti-human RpS6 rabbit antibody for fly lysate (1:1,000, CST, 2217), anti-human RpS6 mouse antibody for HeLa cell lysate (1:1,000, CST, 2317), anti-phosphospecific human RpS6 rabbit antibody for fly lysate (1:1,000, CST, 2211), anti-phosphospecific human RpS6 rabbit antibody for HeLa cell lysate (1:1,000, CST, 5364), anti-HA antibody for HeLa cell lysate (1:1,000, MBL, M180-3), and anti-b-tubulin antibody for fly and HeLa cell lysate (1:500, DSHB, E7). Mammalian Cell Culture and Transfection HeLa cells were cultured in DMEM (Welgene) supplemented with 10% FBS (Invitrogen) at 37 C in a humidified atmosphere of 5% CO2. HeLa cells were seeded in 6-well plates at a density of 1 3 106 cells per well. pHA vector or pHA-cyclin D1 (a gift from Dr. Suk-Chul Bae) was transfected using Lipofectamine LTX Reagent (Invitrogen) according to the manufacture’s protocol. siRNAs for control (Bioneer, #SN-1003), human E2F1 (Bioneer, #1045134), human E2F2 (Bioneer, #1045142), human E2F3 (Bioneer, #1045154), human Cdk4 (Bioneer, #1029179), human Cdk6 (Bioneer, #1029264), human cyclin D1 (Bioneer, #1027294), human cyclin D2 (Bioneer, #1027300) or human cyclin D3 (Bioneer, #1027310) were transfected to HeLa cells using the RNAiMAX reagent (Invitrogen) according to the manufacture’s protocol. Quantitative Real-Time PCR Quantitative real-time PCR was performed using TOPreal qPCR 23 Premix (Enzynomics) on CFX96 (Bio-rad). Human actin levels were measured for internal control of HeLa samples. The results were expressed as fold changes relative to the control. The average mRNA level with standard deviation was obtained from three independent experiments. For quantitative real-time PCR, we used the primers in Table S2. QUANTIFICATION AND STATISTICAL ANALYSIS All experiments were repeated at least three times, and all results were expressed as mean ± SD. Student’s two-tailed t-test was used to determine statistical significance. Prism 7 (Graphpad) was used for the statistical analyses.

Developmental Cell 42, 363–375.e1–e4, August 21, 2017 e4

ERK signaling through E2F1 and promotes melanoma cell growth.

Hedgehog-regulated atypical PKC promotes phosphorylation and activation of Smoothened and Cubitus interruptus in Drosophila.

Cilia-mediated hedgehog signaling in Drosophila.

Spatial expression of the Drosophila segment polarity gene armadillo is posttranscriptionally regulated by wingless.

dachshund Potentiates Hedgehog Signaling during Drosophila Retinogenesis.

Gli Activity as Regulated by Sufu in Drosophila and Mammalian Hedgehog Response.

Cdk4 and Discontinuities in Dpp Signaling Activate TORC1 in the Drosophila Wing Disc.

The vitamin D receptor is required for activation of cWnt and hedgehog signaling in keratinocytes.

Brahma is essential for Drosophila intestinal stem cell proliferation and regulated by Hippo signaling.

Genes targeted by the Hedgehog-signaling pathway can be regulated by Estrogen related receptor β.

Glucose induced activation of canonical Wnt signaling pathway in hepatocellular carcinoma is regulated by DKK4.

Hedgehog and extramacrochaetae in the Drosophila eye: an irresistible force meets an immovable object.

SUMO regulates the activity of Smoothened and Costal-2 in Drosophila Hedgehog signaling.

Insulin-Like Growth Factor and Ecdysone Signaling in Drosophila.

The Histone Acetyltransferase GCN5 Expression Is Elevated and Regulated by c-Myc and E2F1 Transcription Factors in Human Colon Cancer.

Ca2+ Signaling Pathways.

The small GTPase Rap1 is a modulator of Hedgehog signaling.

SPOP-Crn axis in Drosophila.

Asterless Reduction during Spermiogenesis Is Regulated by Plk4 and Is Essential for Zygote Development in Drosophila.

Hedgehog Signaling Strength Is Orchestrated by the mir-310 Cluster of MicroRNAs in Response to Diet.

Microglial phagocytosis and activation underlying photoreceptor degeneration is regulated by CX3CL1-CX3CR1 signaling in a mouse model of retinitis pigmentosa.

Habenular Neurogenesis in Zebrafish Is Regulated by a Hedgehog, Pax6 Proneural Gene Cascade.

Hedgehog signaling pathway regulated the target genes for adipogenesis in silkworm Bombyx mori.

Hedgehog signaling and steroidogenesis.