Abstract
Proteoglycans, a class of carbohydrate-modified proteins, often modulate growth factor signaling on the cell surface. However, the molecular mechanism by which proteoglycans regulate signal transduction is largely unknown. In this study, using a recently-developed glycoproteomic method, we found that Windpipe (Wdp) is a novel chondroitin sulfate proteoglycan (CSPG) in Drosophila. Wdp is a single-pass transmembrane protein with leucin-rich repeat (LRR) motifs and bears three CS sugar chain attachment sites in the extracellular domain. Here we show that Wdp modulates the Hedgehog (Hh) pathway. Overexpression of wdp inhibits Hh signaling in the wing disc, which is dependent on its CS chains and the LRR motifs. Conversely, loss of wdp leads to the upregulation of Hh signaling. Furthermore, knockdown of wdp increase the cell surface accumulation of Smoothened (Smo), suggesting that Wdp inhibits Hh signaling by regulating Smo stability. Our study demonstrates a novel role of CSPG in regulating Hh signaling.
Introduction
Spatial and temporal regulation of growth factor signaling pathways is essential to proper development and disease prevention. Cell surface signaling events, such as ligand-receptor interactions, are often modulated by proteoglycans (D. Xu & Esko, 2014). Proteoglycans are carbohydrate-modified proteins that are found on the cell surface and in the extracellular matrix. They are composed of a core protein and one or more glycosaminoglycans (GAGs) covalently attached to specific serine residues on the core protein. GAGs are long, unbranched, and highly sulfated polysaccharide chains consisting of a repeating disaccharide unit. Based on the composition of the disaccharide units, proteoglycans are classified into several types, including heparan sulfate proteoglycans (HSPGs) and chondroitin sulfate proteoglycans (CSPGs).
HSPGs function as co-receptors by interacting with a wide variety of ligands and modulate signaling activities (Holt & Dickson, 2005; J.-S. Lee & Chien, 2004; Lindahl & Li, 2009; Poulain & Yost, 2015; D. Xu & Esko, 2014). Drosophila offers a powerful model system to study the functions of HSPGs in vivo because of its sophisticated molecular genetic tools and minimal genetic redundancy in genes encoding core proteins and HS synthesizing/modifying enzymes (Lander & Selleck, 2000; Nakato & Li, 2016; Perrimon & Bernfield, 2000; Takemura & Nakato, 2015). In vivo studies using the Drosophila model have shown that HSPGs orchestrate information from multiple ligands in a complex extracellular milieu and sculpt the signal response landscape in a tissue {Nakato & Li, 2016}. However, the molecular mechanisms of co-receptor activities of HSPGs still remain a fundamental question. Our previous studies predict that there are unidentified molecules involved in the molecular recognition events on the cell surface (Akiyama et al., 2008).
In addition to HS, Drosophila produces CS, another type of GAG (Toyoda, Kinoshita-Toyoda, & Selleck, 2000). CSPGs are well known as major structural components of the extracellular matrix. CSPGs have also been shown to modulate signaling pathways, including Hedgehog (Hh), Wnt, and fibroblast growth factor signaling (Cortes, Baria, & Schwartz, 2009; Townley & Bülow, 2018). Given the structural similarities between CS and HS, CSPGs may have modulatory, supportive and/or complementary functions to HSPGs. However, the mechanisms by which CSPGs function as a co-receptor are unknown. In contrast to a large number of studies on HSPGs, very few CSPGs have been identified and analyzed in Drosophila (Momota, Naito, Ninomiya, & Ohtsuka, 2011). Unlike HSPGs, CSPG core proteins are not well conserved between species (Olson, Bishop, Yates, Oegema, & Esko, 2006). Therefore, the identification of CSPGs cannot rely on the sequence homology to mammalian counterparts.
Recently, we have developed a glycoproteomic method to identify novel proteoglycans (Noborn et al., 2016; 2018; 2015). Briefly, this method includes trypsinization of protein samples, followed by enrichment of glycopeptides using strong anion exchange (SAX) chromatography. After enzymatic digestion of HS/CS chains, the glycopeptides bearing a linkage glycan structure common to HS and CS chains are identified using nano-liquid chromatography-tandem mass spectrometry (nLC-MS/MS). This method has successfully identified novel CSPGs in humans (Noborn et al., 2015) and Caenorhabditis elegans (Noborn et al., 2018).
To study the function of CSPGs in signaling, we applied the glycoproteomic method to identify previously unrecognized CSPGs in Drosophila. We found that Windpipe (Wdp) is a novel CSPG and affects Hh signaling. Overexpression of wdp inhibits Hh signaling in the wing disc. This inhibitory effect of Wdp on Hh signaling is dependent on its CS chains and LRR motifs. Consistent with the overexpression analysis, loss of wdp increases Hh signaling. Loss of wdp also increases cell surface accumulation of Smoothened (Smo), the Hh signaling transducer. Therefore, we propose that Wdp downregulates Hh signaling by disrupting cell surface accumulation of Smo.
Results
A glycoproteomic approach identified Wdp as a novel Drosophila CSPG
We investigated the potential presence of CSPGs in Drosophila using our recently-developed glycoproteomic approach that identifies core proteins and its CS attachment sites. A general workflow for the sample preparation, CS-glycopeptide enrichment, LC-MS/MS analysis and the subsequent data analysis is shown in Fig. 1A. Brifely, Drosophila third-instar larvae were collected from two different genotypes (wild type [Oregon-R] and a loss-of-function mutant for tout-velu [ttv524]) and the material was homogenized in ice-cold acetone. ttv encodes a Drosophila HS polymerase, and ttv mutants lack HS chains (Toyoda et al., 2000). The samples were incubated with trypsin and then passed over an anion exchange column equilibrated with a low-salt buffer. This procedure enriches for CS-attached glycopeptides as the matrix retains anionic polysaccharides and their attached peptides, whereas neutral or positively charged peptides flow through the column. The bound structures were eluted stepwise with three buffers of increasing sodium chloride concentrations. The resulting fractions were treated with chondroitinase ABC. This procedure reduces the lengths of the CS chains and generates a residual hexasaccharide structure still attached to the core protein. The chondroitinase-treated fractions were analyzed with positive mode nLC-MS/MS and an automated search strategy was used to identify CS modified peptides in the generated data set (Noborn et al., 2015).
The analysis revealed the Windpipe (Wdp) protein as a novel CSPG, which was modified with three CS-polysaccharides on two unique peptides (Fig. 1B and 1C). We detected Wdp glycopeptides from both wild-type and ttv mutant samples, further supporting that Wdp bears CS chains, not HS. One of the identified precursor ions (m/z 983.38; 3+) equated to the mass of a peptide with a SDQVEGSGDLSETNMELK sequence, derived from the middle part of the protein (amino acids 276– 293) (Fig. 1B). The peptide was modified with one hexasaccharide structure and one methionine oxidation. The measured mass (2947.1186 Da) deviated - 3.27 ppm from the theoretical value. The other identified precursor ion (m/z 1276.76; 4+) equated to the mass of a peptide with a EEHIVKDEDEDDEGSGSGGGLLIIPDPSK sequence, located in proximity to the previous peptide (amino acids 320–348) (Fig. 1C). The peptide was found to be modified with two hexasaccharide structures and where one of the hexasaccharides were modified with one phosphate modification. The measured mass (5102.9389 Da) deviated +3.05 ppm from the theoretical value. Detailed inspection of the spectra revealed several b- and y-ions as well as the prominent diagnostic oxonium ion at m/z 362.1, corresponding to the disaccharide [GlcAGalNAc-H2O+H]+ (Fig. 1B and 1C). Furthermore, one of the glycans in Fig 1C was found modified with one phosphate group at a xylose residue (peptide + xylose + phosphate, m/z 1625.70; 2+).
Wdp is a single-pass transmembrane protein containing four leucine rich repeat (LRR) motifs in the extracellular domain (Huff, Kingsley, Miller, & Hoshizaki, 2002). The three CS attachment sites (S282, S334, and S336) revealed by our glycoproteomic analysis are located in the extracellular domain (Fig. 3A). Interestingly, a recent study reported that Wdp negatively regulates JAK–STAT signaling by promoting internalization and lysosomal degradation of the Domeless (Dome) receptor (W. Ren et al., 2015). We further investigated the role of Wdp, a novel CSPG, in signal transduction.
Overexpression of wdp inhibits Hh signaling
The growth and patterning of the Drosophila wing are controlled by multiple signaling pathways, including Decapentaplegic (Dpp; the Drosophila BMP), Wingless (Wg; the Drosophila Wnt), and Hedgehog (Hh) signaling (Baena-Lopez, Nojima, & Vincent, 2012; Tabata & Takei, 2004). To determine the role of wdp in these developmental signaling pathways, we first asked whether overexpression of wdp affects adult wing morphology. When wdp was overexpressed in the wing pouch using BxMS1096-GAL4 (Capdevila & Guerrero, 1994) (BxMS1096>wdp), the wing size was reduced compared to that of control flies (BxMS1096>) (Fig. 3C; compared to Fig. 3B). In addition, the distance between longitudinal wing veins 3 and 4 (L3 and L4) was aberrantly narrower. This decreased distance between L3 and L4 is indicative of reduced Hh signaling during wing development (Mullor, Calleja, Capdevila, & Guerrero, 1997; Strigini & Cohen, 1997).
Hh is produced in the posterior compartment of the wing disc and spreads towards the anterior compartment where Hh signaling induces target genes expression in a concentration-dependent manner (Briscoe & Thérond, 2013; Gradilla & Guerrero, 2013; Hartl & Scott, 2014). Expression of high-threshold target genes, such as Patched (Ptc; the Hh receptor) (Capdevila, Pariente, Sampedro, Alonso, & Guerrero, 1994) and Engrailed (En) (Patel et al., 1989) are induced in anterior cells near the anteroposterior compartment boundary by high levels of Hh signaling (Jia, Tong, Wang, Luo, & Jiang, 2004) (Fig. 2A and 2E). Low levels of Hh signaling induce the expression of dpp and the accumulation of full-length Cubitus interruptus (Ci; the transcriptional factor of Hh signaling) in a broader region (more distant away from the anteroposterior boundary) (Fig. 2A and 2C). To determine if Hh signaling is indeed affected by wdp, we overexpressed wdp in the dorsal compartment of the wing disc using ap-GAL4 (Calleja, Moreno, Pelaz, & Morata, 1996; O’Keefe, Thor, & Thomas, 1998). We found that wdp overexpression in the dorsal compartment reduced the expression domains of both “high-threshold” targets (Ptc and En) and “low-threshold” targets (dpp-lacZ10638, a reporter for dpp expression, and full-length Ci) compared to those in the ventral compartment (Fig. 2B, 2D, and 2F). Notably, overexpression of wdp did not affect the pattern of a hh transcriptional reporter hh-lacZP30 (J. J. Lee, Kessler, Parks, & Beachy, 1992) (Fig. 2F). Together, wdp acts as a negative regulator of Hh signaling without affecting hh transcription
On the other hand, Wdp does not appear to affect Dpp and Wg pathways. When wdp is overexpressed using ap-GAL4 or hh-GAL4 (a posterior compartment-specific GAL4 driver) (Tanimoto, Itoh, Dijke, & Tabata, 2000), we did not observe apparent defects in Dpp signaling activity, which was monitored by the expression of phosphorylated Mad (pMad) and Spalt major (Salm) (readouts of Dpp signaling). Similarly, no changes in expression of Senseless (Sens) and Distal-less (Dll) (readouts of Wg signaling) were detected (Fig. S1). These results are consistent with a previous report (W. Ren et al., 2015).
We also found that overexpression of wdp induces massive apoptosis, as detected with anti-cleaved Caspase-3 antibody (Fig. S2B). This likely contributed to the smaller adult wing phenotype observed in BxMS1096>wdp flies. It was recently reported that Hh signaling is required for cell survival in wing disc cells (Lu, Wang, & Shen, 2017). To determine whether reduced Hh signaling is responsible for the observed apoptosis, we first asked if reduced Hh signaling results in apoptosis. We inhibited Hh signaling either by expressing an RNAi construct targeting smo (TRiP.HMC03577) (Fig. S2E), or by overexpressing ptc in the dorsal compartment using ap-GAL4. We found that neither treatment caused massive apoptosis (Fig. S2F and S2G), indicating that reduced Hh signaling is not sufficient to induce massive apoptosis in the wing disc. Furthermore, coexpression of a constitutively active form of Smo with Wdp did not suppress apoptosis in the wing disc (Fig. S2H). Thus, these results suggest that overexpression of wdp induces apoptosis, independent of reduced Hh signaling.
CS and LRR motifs are necessary for Wdp to inhibit Hh signaling
Next, we asked whether the CS chains of Wdp are required for its function. In a CSPG core-protein, CS is attached to specific serine residues in the consensus serine-glycine dipeptide surrounded by acidic amino acids (Esko & Zhang, 1996). We generated a UAS-wdpΔGAG construct in which all three serine residues (S282, S334, and S336) are substituted with alanine residues so that CS cannot be attached to the core protein (Fig. 3A). The UAS-wdpΔGAG construct was inserted in the same genomic location (ZH-86Fb; (Bischof, Maeda, Hediger, Karch, & Basler, 2007)) as UAS-wdp using the phiC31 site-specific integration system (Groth, Fish, Nusse, & Calos, 2004) in order to ensure the same expression level of the UAS transgenes.
We found that BxMS1096>wdpΔGAG adult wings did not display the reduction in the distance between L3 and L4 (Fig. 3B). Consistent with this, the expression of Ptc, En, Ci, and dpp-lacZ in the wing disc were not affected by wdpΔGAG overexpression in the dorsal compartment of the wing disc (Fig. 3E–G). These results indicate that CS chains are required for Wdp’s activity to downregulate Hh signaling.
To determine whether the LRR motifs and/or the intracellular domain of Wdp are necessary for inhibiting Hh signaling, we generated several Myc-tagged mutant constructs (Fig. S3) and examined their activities. Consistent with the earlier result (Fig. 2B), expression of a Myc-tagged Wdp (Myc:Wdp) led to the narrower Ptc expression domain (Fig. 3G). A mutant wdp construct lacking LRR motifs (Myc:WdpΔLRRs) failed to inhibit Hh signaling (Fig. 3H). On the other hand, a truncated construct lacking the intracellular domain (Myc:WdpΔICD) retained the ability to inhibit Hh signaling (Fig. 3I). Thus, in addition to CS chains, the LRR motifs of Wdp are required for inhibiting Hh signaling.
Wdp expression in the wing disc
To monitor Wdp expression, we generated transgenic flies (wdpKI.HA and wdpKI.OLLAS) expressing epitope-tagged Wdp protein from its endogenous locus. We inserted a spaghetti monster GFP with 10 copies of HA or OLLAS tags (Nern, Pfeiffer, & Rubin, 2015; Viswanathan et al., 2015) near the C-terminus of Wdp (after Q652; Fig. 4A) using CRISPR–Cas9-mediated homology-directed repair (Gratz et al., 2014; X. Ren et al., 2014). The Wdp:HA expression was detected in the eye disc, adult midgut, and tracheal system (Fig. S3), consistent with previous reports (Huff et al., 2002; W. Ren et al., 2015).
In the wing disc, Wdp:HA is expressed in most of the wing disc cells with enrichment in the basal side, as detected by anti-HA antibody (Fig. 4B and 4C). This result was confirmed by anti-OLLAS antibody staining of the wdpKI.OLLAS wing discs (Fig. S3A and S3B). In the wing disc epithelium, mitotic nuclei apically translocate, but the cells maintain contact with the basal lamina via actin-rich basal projection (Ragkousi & Gibson, 2014). Interestingly, Wdp:HA is strongly enriched in such basal projections (Fig. 4A and 4D). However, physiological significance of this localization of Wdp in the basal projections of mitotic cells is unknown.
Loss of wdp leads to higher levels of Hh signaling
To determine whether loss of wdp affects Hh signaling activity, we examined the effect of wdp RNAi knockdown in the wing disc. Expression of a wdpRNAi construct (TRiP.HMC06302) using ap-GAL4 in wdpKI.HA/+ flies led to the loss of Wdp:HA staining specifically in the dorsal compartment (Fig. 4E), validating the efficacy of RNAi-mediated knockdown of wdp. We then examined the effect of wdp knockdown on Hh signaling using the Ptc expression level as a readout of the Hh signaling activity. In control wing discs (ap>FLP), the signal intensity of Ptc staining in the dorsal compartment is comparable to that in the ventral compartment (Fig. 5B). On the other hand, wdpRNAi expression using ap-GAL4 increased the signal intensity of Ptc staining only in the dorsal compartment (Fig. 5A and 5C). In addition, we observed that the dpp-lacZ expression domain was expanded anteriorly by wdp knockdown (Fig. 5D). In the adult wing, knockdown of wdp slightly expanded the distance between wing vein L3 and L4 near the distal tip (Fig. 5J, compared to Fig. 5I). Thus, wdp RNAi knockdown results in a moderate increase in Hh signaling.
To confirm the wdp knockdown phenotypes, we generated a loss-of-function allele of wdp (wdpKO.ΔCDS), in which most of the wdp coding sequence was removed using CRISPR–Cas9-mediated defined deletion (Gratz et al., 2013) (Fig. 5E–G). wdpKO.ΔCDS homozygous mutant clones were induced in the wing pouch using the FLP–FRT system with nubbin (nub)-GAL4 UAS-FLP and their effect on Hh signaling was examined using anti-Ptc antibody. Consistent with the RNAi knockdown results, we observed a modest increase of Ptc expression in cells mutant for wdp (Fig. 5H). Taken together, we conclude that wdp negatively regulates Hh signaling in the Drosophila wing.
Wdp inhibits Smo cell surface accumulation
The seven-pass transmembrane protein Smo is a key transducer of Hh signaling. In the absence of Hh, Ptc inhibits the phosphorylation of Smo, which is internalized and degraded (Zhu, Zheng, Suyama, & Scott, 2003). In the presence of Hh, restriction of Ptc on Smo is relieved, allowing Smo to accumulate on the cell surface and activate Hh signaling. Although smo transcription is ubiquitous, Smo protein expression levels are high in the posterior compartment of the wing disc where Ptc is not expressed (Fig. 6A) (Denef, Neubüser, Perez, & Cohen, 2000). We found that knockdown of wdp increases the cell surface accumulation of Smo (Fig. 6B). This result suggests that Wdp downregulates Hh signaling either by disrupting Smo translocation to the cell membrane or the stability of Smo on the cell surface.
Discussion
The molecular mechanism by which HSPG co-receptors regulate growth factor signaling remains a central question in cell biology. Dally, a Drosophila HSPG of the glypican type, potentiates Dpp signaling by stabilizing the ligand-receptor complex on the cell surface (Akiyama et al., 2008), suggesting that controlling the rate of receptor-mediated internalization of the signaling complex is the basis for co-receptor activity. However, it is still unknown how HSPGs affect endocytosis and internalization. Since glypicans do not have an intracellular domain, it is likely that these molecules cooperate with other factors (e.g. membrane proteins) to exert co-receptor activity. Thus, it is clear that there are many more unknown factors involved in molecular recognition events on the cell surface. To understand the molecular basis for cell communications, it is critical to identify novel cell surface players.
We found that in the wing disc, Wdp negatively regulates Hh signaling in a CS- and LRR motif-dependent manner. It has also been reported that Wdp negatively regulates JAK–STAT signaling and controls adult midgut homeostasis and regeneration (W. Ren et al., 2015). The authors showed that Wdp interacts with the Dome receptor and promotes its endocytosis and lysosomal degradation. Thus, it is interesting to test if Wdp interacts with Dome via CS chains to modulate JAK–STAT signaling. We observed that Wdp affects cell surface accumulation of Smo, suggesting its role in regulating the stability of Smo protein. Thus, it is possible that Wdp modulates these pathways via a similar mechanism: controlling the internalization of Dome and Smo on the cell membrane.
It is worth noting that both JAK–STAT and Hh signaling, the two pathways negatively controlled by Wdp, are also regulated by HSPGs. Dally-like, a glypican family of HSPGs, positively regulates Hh signaling by interacting with Hh and Ptc (Desbordes & Sanson, 2003; M.-S. Kim, Saunders, Hamaoka, Beachy, & Leahy, 2011; Lum, Yao, et al., 2003a; Williams et al., 2010; Yan et al., 2010). In the developing ovary, Dally upregulates the JAK–STAT pathway (Y. Hayashi et al., 2012). Given the importance of precise dosage control of oncogenic pathways, such as JAK–STAT and Hh signaling, this dual proteoglycan system could play an important role in fine-tuning of the signaling output in order to prevent cancer formation. In vertebrates, HSPGs and CSPGs show opposing effects in neural systems. For example, axon growth is typically promoted by HSPGs but inhibited by CSPGs (Bandtlow & Zimmermann, 2000; Coles et al., 2011; Kantor et al., 2004; Matsumoto, Irie, Inatani, Tessier-Lavigne, & Yamaguchi, 2007; Silver & Miller, 2004; Van Vactor, Wall, & Johnson, 2006). Our findings suggest that such competing effects of HSPGs and CSPGs may be a general mechanism for the precise control of signaling cascades and pattern formation.
In addition to the functions in signaling, Wdp may play other roles. We found that overexpression of wdp results in massive apoptosis in the wing disc, independent of Hh signaling inhibition (Fig. S2). Since CSPGs are well known for structural functions, an excess amount of Wdp may affect the epithelial integrity of the wing disc, leading to subsequent apoptosis. Our observation that Wdp is enriched on the basal side of the wing disc and adult midgut cells (Fig. 4B and S3F) suggests that Wdp may interact with components of the basement membrane, which surrounds these organs.
In mice, sulfated CS is necessary for Indian hedgehog (Ihh) signaling in the developing growth plate (Cortes et al., 2009). Ihh and Sonic hedgehog (Shh) bind to CS (Cortes et al., 2009; Whalen, Malinauskas, Gilbert, & Siebold, 2013; F. Zhang, McLellan, Ayala, Leahy, & Linhardt, 2007). Thus, it will be interesting to check if Wdp interacts with Hh via its CS chains.
Previous studies also reported that wdp is associated with aggressive behaviors in Drosophila species. wdp is upregulated in the head of socially isolated male flies, which exhibit more aggressive behaviors than males raised in groups (L. Wang, Dankert, Perona, & Anderson, 2008). Also, wdp expression is slightly higher in the brain of Drosophila prolongata, which is more aggressive compared to its closely-related species (Kudo et al., 2017). Since CSPGs are important in neuronal patterning (Saied-Santiago & Bülow, 2018), it is interesting to study the molecular mechanisms behind Wdp’s effect on Drosophila behavior.
In mammals, there are a class of CSPG molecules with LRR motifs (small leucine-rich proteoglycans, or SLRPs). A number of SLRP members are known as causative genes of human genetic disorders (Bech-Hansen et al., 2000; Pusch et al., 2000; Schaefer & Iozzo, 2008). Although Wdp does not have cysteine-rich regions that are commonly found in mammalian SLRPs, MARRVEL (ver 1.1) (J. Wang et al., 2017) reports that wdp is a potential Drosophila ortholog of the human NYX gene (nyctalopin), a member of SLRPs (DIPOT score 1 (Hu et al., 2011)). Mutations in NYX cause X-linked congenital stationary night blindness (Bech-Hansen et al., 2000; Pusch et al., 2000). Further studies on Wdp will provide a novel insight into the function of these disease-related human counterparts.
Materials and Methods
Preparation of glycosaminoglycan-glycopeptides and LC-MS/MS analysis
Glycosaminoglycan-glycopeptide samples were prepared from wild-type (Oregon-R) and ttv mutant (ttv524) third-instar larvae as previously described (Noborn et al., 2015; 2018). Briefly, 200-400 third instar larvae (wet weight; 200-400 mg) were lyophilized and homogenized using a motor pestle in 1 ml of ice-cold acetone. After extensive washes with acetone, the insoluble fraction was recovered by centrifugation. After overnight desiccation, the pellet was dissolved in 1.5 ml 1% CHAPS lysis buffer and boiled for 10 min at 96°C. The sample was adjusted to 2 mM MgCl2 and incubated with 3 μl Benzonase (MilliporeSigma, Burlington, MA) at 37°C for three hours. After heat-inactivation of Benzonase, the sample was centrifuged and the supernatant was collected in a new tube.
An aliquot of the preparation (1 mg of protein) was further used. The sample was reduced and alkylated in 1 ml 50 mM NH4HCO3, and trypsinized at 37°C overnight with 20 μg trypsin (Promega, Madison, WI). The digested samples were applied onto DEAE (GE Healthcare, Chicago, IL) columns (600 μl) at 4°C. The columns were washed with three different low-salt washing solutions at 4°C: 50 mM Tris-HCl, 100 mM NaCl, pH 8.0; 50 mM NaAc, 100 mM NaCl, pH 4.0; and 100 mM NaCl. The glycopeptides that were bound to DEAE were eluted stepwise with three buffers with increasing sodium chloride concentrations at 4°C: 4 ml 250 mM NaCl, 400 mM NaCl, 800 mM NaCl, and 3 ml 1500 mM NaCl. Each fraction was desalted using PD10-columns (GE Healthcare).
All fractions were lyophilized and the salt-free samples were then individually treated with 1 mU of chondroitinase ABC (Sigma-Aldrich, St. Louis, MO) for 3 h at 37°C. Prior to MS-analysis, the samples were desalted using a C18 spin column (8 mg resin) according to the manufacturer’s protocol (Thermo Fisher Scientific, Waltham, MA). LC-MS/MS analysis was performed as previously described (Noborn et al., 2015; 2018). In brief, the samples were analyzed on a Q Exactive mass spectrometer coupled to an Easy-nLC 1000 system (Thermo Fisher Scientific). Briefly, glycopeptides (2-μl injection volume) were separated using an analytical column with Reprosil-Pur C18-AQ particles (Dr. Maisch GmbH, Ammerbuch, Germany). The following gradient was run at 300 nl/min; from 7-35 % B-solvent (acetonitrile in 0.2% formic acid) over 75 min, to 100 % B-solvent over 5 min, with a final hold at 100% B-solvent for 10 min. The A-solvent was 0.2% formic acid. Spectra were recorded in positive ion mode and MS scans were performed at 70,000 resolution with a mass range of m/z 600–1800. The MS/MS analysis was performed in a data-dependent mode, with the top ten most abundant charged precursor ions in each MS scan selected for fragmentation (MS2) by higher energy collision dissociation with normalized collision energy values of 30. The MS2 scans were performed at a resolution of 35,000 (at m/z 200). The data analyses were performed as previously described (Noborn et al., 2015) with some small adjustments. In brief, the HCD.raw spectra were converted to Mascot .mgf format using Mascot distiller (version 2.3.2.0, Matrix Science, London, UK). The ions were presented as singly protonated in the output Mascot file. Searches were performed using an in-house Mascot server (version 2.3.02) with the enzyme specificity set to Trypsin, and then to Semitrypsin, allowing for one or two missed cleavages, in subsequent searches on Drosophila sequences of the UniprotKB (42, 507, sequences, 2018-06-18). The peptide tolerance was set to 10 parts per million (ppm) and fragment tolerance was set to 0.01 Da. The searches were allowed to include variable modifications at serine residues of the residual hexasaccharide structure [GlcA(-H2O)GalNAcGlcAGalGalXyl-O-] with 0 (C37H55NO30, 993.2809 Da), 1 (C37H55NO33S, 1073.2377 Da), or 2 (C37H55NO36S2, 1153.1945 Da) sulfate groups attached.
Fly husbandry and fly strains, and transgenic flies
The following fly strains were used in this study:
Oregon-R, w1118 (Bloomington Drosophila Stock Center [BDSC] #5905), ttv524 (Takei, 2004), ap-GAL4 (O’Keefe et al., 1998), hh-GAL4 (Tanimoto et al., 2000), BxMS1096-GAL4 (BDSC #8860) (Capdevila & Guerrero, 1994), AB1-GAL4 (BDSC #1824) (Tavsanli et al., 2004), elavC155>mCD8:GFP (BDSC #5146) (Lin & Goodman, 1994), UAS-GFP (BDSC #1521), UAS-tdTomato (BDSC #36327 and #36328), UAS-FLP (BDSC #4539 and #4540), UAS-ptc (BDSC #44614), nub-GAL4 (BDSC #25754), FRT42D 2xUbi-GFP, UAS-smo:GFP (BDSC #44624), UAS-FLAG:smoAct (BDSC #44621), UAS-wdpRNAi (TRiP.HMC06302, BDSC #66004), UAS-wdpRNAi (TRiP.HM05118, BDSC #28907), UAS-smoRNAi (TRiP.HMC03577, BDSC #53348), hh-lacZP30 (a gift from Gary Struhl) (J. J. Lee et al., 1992), dpp-lacZ10638 (BDSC #12379) (Zecca, Basler, & Struhl, 1995), vas-Cas9 (BDSC #55821), esg-GAL4 (DGRC #113886) (S. Hayashi et al., 2002). The UAS-wdp, UAS-wdpΔGAG, UAS-Myc:wdp, UAS-Myc:wdpΔLRRs, UAS-Myc:wdpΔICD, wdpKO.ΔCDS, wdpKI.HA, wdpKI.OLLAS flies were generated in this study. A full list of genotypes used in this study can be found in Table S1.
For constructing UAS-wdp, wdp CDS (corresponding to wdp-RA–E in FlyBase) was inserted into the XhoI- and XbaI-digested pJFRC7 vector (a gift from Gerald Rubin; Addgene # 26220) (Pfeiffer et al., 2010) using NEBuilder HiFi DNA Assembly Master Mix (New England Biolabs [NEB], Ipswich, MA, E2621S). Similarly, wdpΔGAG (S282A, S334A, and S336A), Myc:wdp, Myc:wdpΔLRRs, and Myc:wdpΔICD were inserted into the pJFRC7 vector. The UAS transgenic flies were generated using phiC31 integrase-mediated transgenesis at the ZH-86Fb attP (FBti0076525) integration site. Embryonic injection was performed by BestGene Inc (Chino Hills, CA). Primers used in this study will be available upon request.
To generate the wdpKO.ΔCDS allele, two sgRNAs (pU6-sgRNA-wdp-1 and pU6-sgRNA-wdp-2) were introduced to delete the wdp CDS. To construct sgRNA plasmids, 5’ - CTTCGACAGGGCCAACCAGGCGGTC - 3’ and 5’ - AAACGACCGCCTGGTTGGCCCTGTC - 3’ were annealed (pU6-sgRNA-wdp-1); and 5’ - CTTCGAGTGGCCATTGATCACCTGG - 3’ and 5’ - AAACCCAGGTGATCAATGGCCACTC - 3’ (pU6-sgRNA-wdp-2) were annealed and ligated in the BbsI-digested pU6-BbsI-chiRNA plasmid (a gift from Melissa Harrison, Kate O’Connor-Giles, and Jill Wildonger; Addgene #45946) (Gratz et al., 2013). A mixture of 50 ng/µl of pU6-sgRNA-wdp-1 and pU6-sgRNA-wdp-2 was injected into the embryos of the vas-Cas9 flies, which express Cas9 under the control of the germline vasa regulatory elements (Gratz et al., 2014), by BestGene Inc. The wdpKO.ΔCDS allele was screened by PCR and verified by Sanger sequencing.
To generate the wdpKI.HA allele, we constructed a donor plasmid, which contained a Gly-Gly-Ser linker, smGFP-HA, and approximately 1-kb homology arms to wdp flanking the linker and smGFP-HA, for homology-directed repair. smGFP-HA and the wdp homology sequences on either side of the targeted DSB were PCR-amplified from pJFRC201-10XUAS-FRT>STOP>FRT-myr:smGFP-HA (a gift from Gerald Rubin; Addgene plasmid #63166) (Nern et al., 2015) and genomic DNA extracted from the vas-Cas9 flies, respectively. These fragments were cloned into the pHD-DsRed-attP backbone (a gift from Melissa Harrison, Kate O’Connor-Giles and Jill Wildonger; Addgene #51019) (Gratz et al., 2014) using NEBuilder HiFi DNA Assembly Master Mix (NEB, E2621S). Similarly, we generated a donor plasmid with OLLAS tags amplified from pJFRC210-10XUAS-FRT>STOP>FRT-myr:smGFP-OLLAS (a gift from Gerald Rubin; Addgene plasmid #63170) (Nern et al., 2015). A mixture of 50 ng/µl of pU6-sgRNA-wdp-2 and 125 ng/µl of each donor plasmid was injected into the vas-Cas9 embryos by BestGene Inc. The wdpKI.HA and wdpKI.OLLAS alleles were screened by PCR and verified by Sanger sequencing.
Flies were raised on a standard cornmeal fly medium at 25°C unless otherwise indicated.
Mosaic analysis
The wdpKO.ΔCDS homozygous clones were generated by FLP/FRT-mediated mitotic recombination (T. Xu & Rubin, 1993). The FLP expression was induced by nub-GAL4 UAS-FLP.
Immunohistochemstry
Third-instar larval imaginal discs were stained as described previously (Takemura & Adachi-Yamada, 2011) with some modifications. Wing discs were dissected from third-instar wandering larvae in phosphate-buffered saline (PBS, pH 7.4) and subsequently fixed in 3.7% formaldehyde in PBS for 15 min at room temperature. After three 10-min washes with PBST (PBS containing 0.1% (vol/vol) Triton X-100 [Sigma, T8532]), the samples were incubated in primary antibodies overnight at 4°C. After three 10-min washes with PBST, the samples were incubated with Alexa Fluor-conjugated secondary antibodies (1:500, Thermo Fisher Scientific) overnight at 4°C or 2 hours at room temperature. After three 10-min washes with PBST, the samples were stained with 1 µg/ml DAPI (Thermo Fisher Scientific, 62248) and subsequently mounted in VECTASHIELD Antifade Mounting Medium (Vector Laboratories, Burlingame, CA, H-1000). F-actin was stained with Alexa Fluor 568 phalloidin (Thermo Fisher Scientific, A12380). Adult midguts were dissected and immunostained as previously described (Takemura & Nakato, 2017). Images were acquired on a LSM710 confocal microscope (Carl Zeiss, Oberkochen, Germany). For quantification of Ptc staining, images were acquired with the same condition, and fluorescence intensity was measured in a set area with Fiji (Schindelin et al., 2012).
Antibodies
The primary antibodies used were as follows: mouse anti-Ptc Apa 1 (1:20, Developmental Studies Hybridoma Bank [DSHB], Iowa City, IA, deposited by Isabel Guerrero) (Capdevila et al., 1994), rat anti-Ci 2A1 (1:20, DSHB, deposited by Robert Holmgren) (Motzny & Holmgren, 1995), chicken anti-βGalactosidase (1:2000, Abcam), mouse anti-En 4D9 (1:20, DSHB, deposited by Corey Goodman) (Riggleman, Schedl, & Wieschaus, 1990), rabbit anti-pH3 (1:1000, Millipore, 06-570), rat anti-HA 3F10 (1:200, Roche, 11867423001), rabbit anti-HA C29F4 (1:1000, Cell Signaling, 3724), mouse anti-Smo 20C6 (1:50, DSHB, deposited by Philip Beachy) (Lum, Zhang, et al., 2003b), rabbit anti-pSmad3 (1:1000, Epitomics, 1880-1) (Smith, Machamer, Kim, Hays, & Marques, 2012), rabbit anti-Salm (1:30, a gift from Scott Selleck), mouse anti-Dll 1:500 (1:500, a gift from Dianne Duncan) (D. M. Duncan, Burgess, & Duncan, 1998), guinea pig anti-Sens (1:1000, a gift from Hugo Bellen) (Nolo, Abbott, & Bellen, 2000), rabbit anti-cleaved Caspase-3 (1:200, Cell Signaling, 9661), rat anti-OLLAS L2 (1:500, Novus Biologicals, NBP1-06713), mouse anti-Arm N2 7A1 (1:50, DSHB, deposited by Eric Wieschaus) (Riggleman et al., 1990), mouse anti-Pros MR1A (1:50, DSHB, deposited by C.Q. Doe) (Campbell et al., 1994), and mouse anti-Fas3 7G10 (1:50, DSHB, deposited by Corey Goodman) (Patel, Snow, & Goodman, 1987). Alexa488, Alexa548, Alexa564 and Alexa633-conjugated secondary antibodies (Thermo Fisher Scientific) were used at a dilution of 1:500.
Adult wing preparation
The left wings from female flies were dissected and mounted on slides using Canada balsam (Benz Microscope, BB0020) as previously described (Takemura & Adachi-Yamada, 2011). Images were taken using a ZEISS Stemi SV 11 microscope equipped with a Jenoptic ProgRes C3 digital camera.
Competing interests
The authors declare no competing or financial interests.
Acknowledgements
We thank Melissa Harrison, Kate O’Connor-Giles, Jill Wildonger, Gary Struhl, Scott Selleck, Hugo Bellen, the Bloomington Drosophila Stock Center (NIH P40OD018537), the Transgenic RNAi Project at Harvard Medical School [NIH/NIGMS R01-GM08947], and the Drosophila Genomics Resource Center (NIH 2P40OD010949) for sharing fly strains and plasmids. We also thank the Proteomics Core Facility at the Sahlgrenska Academy, University of Gothenburg, Sweden for running all the MS analyses. This work was supported by the National Institutes of Health (R01 GM115099) to H.N. M.T. held postdoctoral fellowships from the Japan Society for the Promotion of Science and the Uehara Memorial Foundation.