The N-end rule pathway is a proteolytic system in which single N-terminal residues of proteins act as N-degrons. These degrons are recognized by N-recognins, facilitating substrate degradation via the ubiquitin (Ub) proteasome system (UPS) or autophagy. We have previously identified a set of N-recognins [UBR1, UBR2, UBR4 (also known as p600) and UBR5 (also known as EDD)] that bind N-degrons through their UBR boxes to promote proteolysis by the proteasome. Here, we show that the 570 kDa N-recognin UBR4 is associated with maturing endosomes through an interaction with Ca2+-bound calmodulin. The endosomal recruitment of UBR4 is essential for the biogenesis of early endosomes (EEs) and endosome-related processes, such as the trafficking of endocytosed protein cargos and degradation of extracellular cargos by endosomal hydrolases. In mouse embryos, UBR4 marks and plays a role in the endosome-lysosome pathway that mediates the heterophagic proteolysis of endocytosed maternal proteins into amino acids. By screening 9591 drugs through the DrugBank database, we identify picolinic acid as a putative ligand for UBR4 that inhibits the biogenesis of EEs. Our results suggest that UBR4 is an essential modulator in the endosome-lysosome system.
In eukaryotes, cellular proteins are mainly degraded by the ubiquitin (Ub) proteasome system (UPS), autophagy, and/or the endosomal system. In the UPS, proteasomal proteases degrade Ub-tagged short-lived and misfolded/damaged substrates into short peptides (Braten et al., 2016; Ciechanover and Kwon, 2015, 2017; Ji and Kwon, 2017). Autophagy uses lysosomal hydrolases to degrade various long-lived and insoluble misfolded proteins into amino acids (Mizushima and Komatsu, 2011). By contrast, plasma membrane-associated and extracellular proteins are endocytosed, sequestered in endosomes and degraded by endosomal hydrolases into short peptides and truncated proteins (Authier et al., 1995; Blum et al., 1991; Huotari and Helenius, 2011; Pillay et al., 2002; Tjelle et al., 1996). Studies have shown that ∼50% of the plasma membrane and 5–30% of the cellular volume are internalized in every hour (Maxfield, 2014; Steinman et al., 1983). As such, the plasma membrane and associated proteins can be almost entirely replaced by new materials in just one hour (Maxfield, 2014; Steinman et al., 1983). Although the majority of cell surface proteins are short-lived substrates of endosomal proteolysis, the key regulators underlying endosomal processes remain incompletely understood.
Substrates of endosomal proteolysis are endocytosed and sequestered within endocytic vesicles (EVs) (Authier et al., 1996; Berg et al., 1995; Ciechanover et al., 1983a,b; Maxfield, 2014) (see Fig. S1). EVs can be recycled back to the membrane (Conner and Schmid, 2003) or fused with each other to form early endosomes (EEs; pH 5.8-6.3) (Cabrera et al., 2013; Huotari and Helenius, 2011) in which the cargos are digested by endosomal hydrolases into short peptides or truncated fragments (Abrahamson and Rodewald, 1981; Appelqvist et al., 2013; Besterman and Low, 1983; Huotari and Helenius, 2011; Steinman et al., 1983). The partially digested cargos are further degraded as EEs undergo maturation into late endosomes (LEs; pH 5.4–5.8) containing ∼50 hydrolases (see Fig. S1). The proteolytic products in LEs are released into the cytosol and further digested by the proteasome and aminopeptidases (Dasgupta et al., 2014; Pillay et al., 2002). Together, ∼80% of endosomal cargos destined for degradation are digested within EEs or LEs and thus occurs independently of lysosomes and via a process that is mechanistically distinct from the UPS and autophagy (Repnik et al., 2013; Tjelle et al., 1996). The undigested cargos in LEs are delivered to lysosomes (pH 5.0) and degraded into amino acids by lysosomal hydrolases (Pillay et al., 2002). Endosomes also mediate a lysosome-independent degradation of autophagic cargos sequestered in hydrolase-free autophagosomes. Specifically, a portion of EEs and LEs fuse with autophagosomes to form amphisomes and initiate the degradation of autophagic cargos via their endosomal hydrolases (Berg et al., 1998; Pillay et al., 2002) (Fig. S1). In addition, some LEs are co-deposited into multivesicular bodies (MVBs) along with autophagosomes and their cargos (Fig. S1), wherein autophagic cargos are degraded by endosomal hydrolases, independently of lysosomes (Fengsrud et al., 2000). The undigested and partially digested cargos in amphisomes and MVBs are eventually digested by endosome-derived hydrolases in lysosomes into amino acids (Fig. S1). Overall, the amounts of cargos degraded in EEs, LEs, MVBs and amphisomes, prior to lysosomal targeting, may rival or even exceed those degraded in lysosomes, especially in normally growing cells.
The EVs containing endosomal cargos mature into EEs and subsequently LEs through a series of homotypic fusions. During this process, GTP-bound RAB5 (which has RAB5A, RAB5B and RAB5C forms) is associated with EVs and induces the formation of phosphatidylinositol 3-phosphate (PI3P) on EVs (Huotari and Helenius, 2011) (Fig. S1). The PI3P moieties on EVs subsequently recruit early endosome antigen 1 (EEA1). PI3P-bound EEA1 is a maximum of ∼240 nm long and tethers two EVs through the interaction of its two ends with two RAB5 molecules, each of which are on an EV (Huotari and Helenius, 2011; Simonsen et al., 1998, 1999). The tethering of two EVs by EEA1 is facilitated by Ca2+-bound calmodulin, which binds EEA1 and induces its conformational change from 240 nm to 70 nm long (Colombo et al., 1997; Lawe et al., 2003; Zerial and McBride, 2001) (Fig. S1). The cooperation among RAB5, EEA1 and calmodulin is critical for homotypic fusion and maturation of EVs into EEs (Fig. S1). The resulting EEs can fuse with EVs or EEs in the course of maturation to an LE (Huotari and Helenius, 2011). During the conversion of EEs into LEs, RAB5 is dissociated from EEs and replaced with the G-protein RAB7 (which has RAB7A and RAB7B forms) (Poteryaev et al., 2010). To date, little is known about the peripheral components that modulate endosomal maturation and proteolysis.
The N-end rule pathway is a proteolytic system in which single N-terminal amino acids function as a determinant of degradation signals (degrons), called N-degrons (Bachmair et al., 1986; Sriram et al., 2011; Tasaki et al., 2012). The degradation determinants include positively charged (Arg, Lys and His; type 1) and bulky hydrophobic (Phe, Tyr, Trp, Leu and Ile; type 2) residues on the protein N-termini (Kwon et al., 1999; Lee et al., 2008). Among these, the Arg N-degron can be generated by the conjugation of the amino acid L-Arg by ATE1-encoded R-transferases (EC 2.3.2) (Kwon et al., 2000; Lee et al., 2005; Sriram and Kwon, 2010). The N-terminal Arg (Nt-Arg) of arginylated proteins can induce proteolysis by the N-end rule pathway via either the UPS or autophagy (Cha-Molstad et al., 2015, 2016, 2017). We have previously identified a family of N-recognins (UBR1, UBR2, UBR4 and UBR5) that recognize the Nt-Arg using their UBR boxes and induce substrate ubiquitylation and proteasomal degradation (Kwon et al.,1998, 2001, 2003; Tasaki et al., 2009). The E3 ligases UBR1 and UBR2 are functionally and structurally overlapping 200-kDa RING finger proteins that mediate ubiquitylation of short-lived and misfolded proteins in the cytosol and nucleus (Kwon et al., 2003). Compared with UBR1 and UBR2, the biochemical mechanisms and physiological functions of UBR4 and UBR5 as N-recognins remain poorly understood. More recently, we found that the Nt-Arg of arginylated proteins can induce autophagic proteolysis through the recognition by the N-recognin SQSTM1 (also known as p62) (Cha-Molstad et al., 2015, 2016, 2017). The substrates of autophagic proteolysis include endoplasmic reticulum (ER)-residing molecular chaperones, such as BiP (also known as GRP78 and HSPA5), calreticulin and protein disulfide isomerase (Cha-Molstad et al., 2015). To date, the N-end rule pathway has not been implicated in endosomal proteolysis.
Human UBR4 is a microtubule-associated protein with a size of 570 kDa (Kim et al., 2013; Shim et al., 2008; Tasaki et al., 2013). UBR4 has been implicated in various biological processes, ranging from ubiquitylation and proteasomal degradation of short-lived proteins (Rinschen et al., 2016) and the pathogenesis of neurodegeneration (Amorim et al., 2015; Wishart et al., 2012) to the secretion of microvesicles and ectosomes (Basso and Bonetto, 2016; Keerthikumar et al., 2015), cellular immortalization and transformation activities mediated by human papillomavirus (HPV) E7 (DeMasi et al., 2005, 2007; Huh et al., 2005; White et al., 2012), and cell adhesion and integrin-mediated apoptosis (Nakatani et al., 2005). Of note, UBR4 is a binding target of various viral proteins, such as influenza A virus M2 protein (Tripathi et al., 2015), the NS5 methyltransferase of dengue and Zika viruses (Grant et al., 2016; Morrison et al., 2013), and the E7 proteins of HPV, as well as bovine papillomavirus (BPV) (Szalmas et al., 2017; White et al., 2016). The mutations in human UBR4 has been linked to early-onset dementia as well as developmental regression, brain atrophy and ataxia (Monies et al., 2017). The Arabidopsis UBR4 homolog (known as BIG) is required for the vesicular transport of the master plant hormone auxin (Gil et al., 2001), whereas the Drosophila UBR4 homolog (known as Calossin or Pushover) plays a role in synaptic transmission in photoreceptor cells and perineurial glial growth (Sekelsky et al., 1999; Yager, 2001). Despite these isolated observations, the biochemical properties and physiological functions of UBR4 remain largely unknown.
We and others have constructed knockout mice lacking UBR4 (Nakaya et al., 2013; Tasaki et al., 2013). Our mutant animals, in which a DNA region encoding the UBR box was deleted, die during embryogenesis at midgestation [at embryonic day (E)9.5–10.5] because of vascular defects in the yolk sac (YS) (Kim et al., 2013; Tasaki et al., 2013). At this developmental stage, the YS is the major supplier of amino acids essential for protein synthesis in embryonic bodies, because the placenta is not functional until E15 (Malassine et al., 2003). The YS is divided into two layers: the visceral endoderm and extraembryonic mesoderm (Zohn and Sarkar, 2010). The outer endoderm massively endocytoses extraembryonic nutrients and maternal proteins (e.g. albumins and transferrins), which are subsequently degraded by endosomal hydrolases in EEs and LEs, leading to lysosomal degradation into amino acids (Huxham and Beck, 1985; Kawamura et al., 2012; Richardson et al., 2000). The resulting amino acids are transported to embryonic bodies via blood vessels within the YS mesoderm as an essential source for protein synthesis in embryonic bodies (Zohn and Sarkar, 2010). Thus, almost all proteins in embryos at midgestation are made of the proteolytic products of endocytosed extracellular proteins. To date, the mechanisms underlying the null phenotypes of Ubr4−/− embryos remain unknown.
Our earlier work has identified UBR4 as an N-recognin that can bind type-1 and type-2 N-degrons including Nt-Arg (Tasaki et al., 2005). In this study, we show that UBR4 modulates the biogenesis of endosomes and the formation and functionality of endosome-related cellular structures. UBR4 loss impacts the trafficking and proteolysis of endocytosed protein cargos. Our results suggest that UBR4 is a key regulator in the endosome-lysosome system, and thereby provide the mechanisms underlying the aforementioned observations on UBR4.
The EEA1-mediated endosomal pathway is impaired in the YS and embryos lacking mouse UBR4
We have previously found that Ubr4−/− mouse embryos die at E9.5–10.5 and show vascular defects in the YS (Tasaki et al., 2013). At this stage, embryos cannot synthesize amino acids de novo and, thus, obtain amino acids by breaking down endocytosed proteins in the YS (Fig. 1A) (Kawamura et al., 2012). These extracellular protein cargos (e.g. albumins), provided by the mothers, are endocytosed by the YS and sequestered and degraded by EEA1+ EEs (Pillay et al., 2002; Zohn and Sarkar, 2010). The cargos within EEs are further degraded upon the fusion with autophagosomes to generate amphisomes and, subsequently, by lysosomal hydrolases (Kawamura et al., 2012). To determine the role of UBR4 in endosomal pathways in animals, we examined the localization of UBR4 in cross sections of E9.5 +/+ and Ubr4−/− YSs and embryonic bodies. Immunostaining analysis showed that UBR4 is prominently expressed in the YS endoderm, a specialized tissue that has a high level of endosomal proteolysis (Fig. 1B,C). UBR4 signals formed cytosolic puncta that colocalized with EEA1+ EEs (Fig. 1C). UBR4+ EEA1+ punctate signals were more prominent in the YS endoderm than in the YS mesoderm (Fig. 1C). Within the YS endoderm, both UBR4 and EEA1 signals were enriched at the apical side of the cytosol and near lysosomal vacuoles. A significant portion of the UBR4+EEA1+ puncta were distributed along the axis from the plasma membrane to apical vacuoles (Fig. 1C). Moreover, the EEA1 staining was significantly reduced in the YS of Ubr4−/− embryos at E9.5 (Fig. 1D). These results suggest that UBR4 plays a role in endosomal proteolysis of extraembryonic proteins to generate the amino acids required for protein synthesis during embryogenesis.
To further validate these results in embryonic tissues, we immunostained for EEA1 on the cross sections of +/+ and Ubr4−/− mouse embryos at E9.5. Whereas EEA1 was detected as punctate structures in various tissues of +/+ embryos, such signals were markedly downregulated and disorganized in various tissues of Ubr4−/− embryos, including the neural tube and paraxial mesoderm of neuronal tissues (Fig. 1E). These results collectively suggest that UBR4 plays a role in the biogenesis of EEs in YS and various tissues of mouse embryos.
UBR4 is dispensible for the internalization of extracellular ligands
To characterize the role of UBR4 in endosomal pathways, we compared the internalization of extracellular ligands in +/+ and Ubr4−/− mouse embryonic fibroblasts (MEFs) by measuring the uptake of horseradish peroxidase (HRP), a fluid-phase marker for bulk endocytosis (Li et al., 1995; Li and Stahl, 1993; Stoorvogel, 1998). Quantification analyses showed that the cytosol of Ubr4−/− MEFs contained a comparable amount of HRP to that in +/+ MEFs (Fig. 2A). As an alternative way to monitor the endocytosis and visualize intracellular migration of EEs, cells were treated with FITC-conjugated TAT peptide derived from the transactivator of transcription (TAT) of human immunodeficiency virus (HIV). The cell-penetrating TAT peptide binds various cell surface receptors and is internalized through the endosomal pathways (Lönn et al., 2016; Vivès et al., 1997; Yezid et al., 2009). Fluorescence assays were employed to assess the number and sizes of initially generated TAT+ punctate signals in +/+ and Ubr4−/− MEFs (Fig. 2B). Quantification of fluorescence signals showed that the mean diameter of TAT–FITC+ vesicles in the Ubr4−/− MEF cell was half of that in +/+ (Fig. 2C). Instead, Ubr4−/− MEFs accumulated an excessive number (∼6-fold increase) of small-sized TAT-positive puncta as compared with +/+ MEFs (Fig. 2D). Nonetheless, the total amounts of internalized TAT+ signals in Ubr4−/− MEFs were largely similar to those in +/+ cells (Fig. 2B–D). These results suggest that UBR4 is largely dispensable for the internalization of extracellular ligands but plays a role in EE maturation.
UBR4 is associated with endosomes through the interaction with Ca2+-calmodulin
We examined the intracellular localization of UBR4 in comparison with endosomal markers. Immunostaining analysis using super-resolution confocal microscope (N-SIM) with a resolution of 100 nm showed that UBR4 staining was present as punctate structures with sizes of 150×300 nm (Fig. 3A; Fig. S2A). A significant portion of UBR4+ punctate signals colocalized in the proximity of RAB5+ endosomes in a fashion that shows that they tether two RAB5+ endosomes (Fig. 3B,C). To further characterize the localization of UBR4 on endosomes, we performed transmission electron microscopy using HEK293 cells stably expressing UBR4–V5. Immunogold-labeled UBR4 molecules were also associated with two EVs (Fig. 3D). Given the morphology and sizes, these RAB5+ endosomes associated with UBR4 appeared to be EVs in the process of maturation into EEs. As an alternative way to monitor the association of UBR4 with endosomes, we separated +/+ and Ubr4−/− MEFs into soluble cytosol (S150), cell membrane (P15S) and cytosolic vesicle (P150) fractions using differential centrifugation (Fig. 3E). Both UBR4 and EEA1 were mainly retrieved from cytosolic vesicles in addition to the soluble cytosol. These results suggest that UBR4 is associated with endosomes (Fig. 3F).
Next, we searched for the proteins that link UBR4 to endosomes. During the maturation of EVs, Ca2+-bound calmodulin is associated with EVs and induces the conformational change of EEA1, facilitating the homotypic fusion of two EVs into an EE (Dumas et al., 2001; Lawe et al., 2000). To date, little is known about how calmodulin activity is regulated during endosome biogenesis. Given the association of UBR4 with endosomes, we paid attention to a previous study in which UBR4 was identified as a calmodulin-binding protein, although its meaning remains unknown (Nakatani et al., 2005). To validate this finding, we performed in vitro interaction assays using UBR4–GFP stably expressed in HEK293 cells. The results showed that UBR4 indeed bound calmodulin and that their interaction was disrupted when Ca2+ was chelated (Fig. 3G). We therefore examined the colocalization of UBR4 and calmodulin on endosomes in HeLa cells transiently expressing calmodulin–EGFP and EEA1–TagRFP. Co-immunostaining analyses showed that virtually all of the calmodulin+ EEA1+ puncta were also positive for UBR4 (Fig. 3H). When cells were treated with the calmodulin inhibitor W7 for 15 min, UBR4 as well as calmodulin were dissociated from EEA1+ puncta (Fig. 3H,I). Next, to obtain more decisive evidence that UBR4 is recruited to EEA1+ endosomes, we depleted EEA1 with siRNA. Immunostaining analyses showed that knockdown of EEA1 in HeLa cells induced the delocalization of UBR4 from RAB5+ endosomes (Fig. S2B,C). These results suggest that UBR4 is associated with maturing endosomes in part through its interaction with Ca2+-calmodulin.
UBR4 is required for the biogenesis of EEs
To determine the role of UBR4 in endosomal pathways, we compared the biogenesis of EEs in +/+ and Ubr4−/− MEFs via immunostaining analysis of EEA1. Wild-type MEFs contained ∼245 EEA1+ puncta per cell throughout the cytosol (Fig. 4A,B). By contrast, Ubr4−/− MEFs contained a significantly reduced number of mature EEs associated with excessively accumulated EV-like small-sized EEA1+ puncta with a diameter of less than 200 nm (Fig. 4C,D). Such a downregulation of EEA1+ EEs was readily reproduced by transiently depleting UBR4 using siRNA (Fig. 4D,E). Despite the downregulation of EEA1+ puncta maturation, Ubr4−/− MEFs contained a normal level of EEA1 as determined by a triplicate set of immunoblotting analyses (Fig. 4F,G). Consistent with this, UBR4-knockdown HeLa cells showed no differences in the levels of EEA1 compared with controls (Fig. S3A,B). Moreover, the overexpression of EEA1 in Ubr4−/− MEFs did not restore the defect (Fig. 4I). These results suggest that the biogenesis of EEs is impaired in the absence of UBR4.
To address the direct role of UBR4 in EE biogenesis, we asked whether exogenous expression of UBR4 would rescue the null phenotype of Ubr4−/− MEFs. Immunostaining analyses of EEA1 showed that the overexpression of UBR4 significantly restored the ability of Ubr4−/− MEFs to form EEA1+ puncta (Fig. 4J,K; Fig. S3D). This suggests that the failure to form EEA1+ endosomes is not caused by EEA1 insufficiency but the misregulation of EEA1 functionality. To validate this result, we used another marker for the early endosome, RAB5, which is associated with EVs and EEs (Rink et al., 2005). Consistent with the results with EEA1, Ubr4−/− MEFs were notably impaired in the formation of RAB5+ EEs (Fig. 4L). Moreover, transiently overexpressed UBR4 readily rescued the biogenesis of RAB5+ EEs in Ubr4−/− MEFs (Fig. 4M,N). Whereas there were fewer RAB5+ EEs in Ubr4−/− MEFs, immunoblotting analysis showed that Ubr4−/− MEFs contained a comparable level of RAB5 (Fig. 4F,H). Consistent with this, UBR4-knockdown HeLa cells also showed no difference in the level of RAB5 compared with controls (Fig. S3A,C). Next, we also monitored the formation of EEs positive for APPL1, a marker of signaling endosomes. During EE biogenesis, RAB5+ endosomes undergo maturation into either EEA1+ or APPL1+ endosomes, which are mutually exclusive with each other (Kalaidzidis et al., 2015). Consistent with our findings that UBR4-deficient cells are impaired in the formation of not only RAB5+ endosomes but also EEA1+ EEs, a markedly reduced level of APPL1+ EEs were generated when UBR4 was depleted (Fig. S4A–C). These results suggest that UBR4 modulates the biogenesis of EEs.
Next, we examined the possible role of UBR4 in the formation of LEs and lysosomes by immunostaining for RAB7+ LEs and LAMP1+ lysosomes. The results showed that Ubr4−/− MEFs contained a similar level of RAB7+ LEs to +/+ MEFs (Fig. S4D,E), consistent with the facts that LEs can be generated from diverse sources such as autophagic membranes from the ER and mitochondria, and that RAB7 is associated with various types of vesicular structures, such as autophagosomes and MVBs, in addition to LEs (Yamano et al., 2018; Zhang et al., 2009). Likewise, Ubr4−/− MEF cells contained a comparable level of LAMP1+ lysosomes, indicating that UBR4 is not required for biogenesis of lysosomes (Fig. S4F,G). Finally, we asked whether knockdown of EEA1 would impair the formation of RAB5+ EEs. As expected, upon transient knockdown of EEA1, the number of RAB5+ EEs with a diameter of ∼500 nm was reduced to ∼30% relative to control (Fig. S4H,I). Collectively, these results emphasize the role of UBR4 in the biogenesis of EEs.
We have previous identified a set of N-recognins (UBR1, UBR2, UBR4 and UBR5) characterized by the UBR box (Tasaki et al., 2005). Among these, UBR1 and UBR2 have been implicated in the UPS-linked N-end rule pathway (Lee et al., 2008; Sriram et al., 2011). To determine whether the aforementioned role in endosomal pathways is confined to UBR4 among these N-recognins, we performed immunostaining analysis. UBR1−/−UBR2−/− MEFs contained a normal number of EEA1+ as well as RAB5+ EEs as compared with +/+ MEFs (Fig. S4J). These results suggest that UBR4 can be a designated N-recognin that controls endosomal maturation.
UBR4 regulates endosomal trafficking
To determine the role of UBR4 in endosomal trafficking, we treated +/+ and Ubr4−/− MEFs with Alexa Fluor 555-conjugated epidermal growth factor (EGF) and monitored the intracellular trafficking of the EGF–EGFR complex using co-immunofluorescence analyses. First, we checked the level of total EGFR (Fig. S5A). Even though Ubr4−/− MEFs showed a ∼2-fold reduction of EGFR level relative to +/+ MEFs, the amount of EGF internalized was comparable with that in wild type (Fig. S5B). By 10 min, EGFR+ endosomes in +/+ MEFs were distributed across the cytosol, from the plasma membrane to the peri-nuclear region (Fig. 5A–C). By 20 min, most EGF+ EGFR+ endosomes migrated to the peri-nuclear region, indicative of endosomal trafficking (Fig. 5B,C). Most of the EGF+ EGFR+ signals observed in +/+ MEFs appeared to represent EEs as they were positive for the EV/EE marker RAB5 (Fig. S5C) but negative for the LE marker RAB7 (Fig. S5D). By sharp contrast, Ubr4−/− MEFs contained a markedly decreased number of EGF+ EGFR+ signals, which appeared to be confined to EVs based on their morphology and sizes (Fig. 5C). As an alternative assay, we quantified the intracellular localization of EGF+ EGFR+ endosomes along the plasma membrane–nucleus axis at 10 min and 20 min. Whereas most of the EGF+ EGFR+ endosomes in +/+ MEFs migrated to the peri-nuclear region, those in Ubr4−/− MEFs stayed near the plasma membrane (Fig. 5D–F). The generation and intracellular trafficking of EGF+ RAB5+ endosomes were also impaired in Ubr4−/− MEFs in a pattern similar to that of EGF+ EGFR+ endosomes (Fig. S5C). Despite impaired endosomal trafficking, Ubr4−/− MEFs retained cytoskeleton structures, as determined by immunostaining analyses of actin filaments and microtubules (Fig. S5E). These results suggest that UBR4 is required for endosomal trafficking of ligand–receptor complexes internalized from the plasma membrane.
Defective endosomal proteolysis in UBR4-deficient MEFs
One important function of endosomes is the turnover of plasma membrane-associated and extracellular proteins. To characterize the role of UBR4 in endosomal proteolysis, we treated +/+ and Ubr4−/− MEFs with the endocytic cargo DQ-BSA (Marwaha et al., 2017), which generates fluorescence upon endosomal proteolysis. Fluorescence analysis showed that in +/+ MEFs, DQ-BSA was normally endocytosed and degraded by endosomal hydrolases (Fig. 6A,B). In sharp contrast, the degradation of DQ-BSA was markedly impaired in Ubr4−/− MEFs as compared with +/+ cells (Fig. 6A,B). Such an inhibition of DQ-BSA degradation was reproduced when UBR4 was transiently silenced using siRNA (Fig. 6A,B). These results show that the defect in EE biogenesis causes the failure to degrade endosomal protein cargos, implicating UBR4 in endosomal proteolysis.
Identification of a UBR4 ligand and its use to reversibly inhibit EE biogenesis
To develop a pharmaceutical reagent targeting UBR4, we screened 9591 drugs through DrugBank 5.0 database, a bioinformatics and cheminformatics resource containing drugs and their (putative) targets (Law et al., 2014). This screening identified picolinic acid as a putative ligand for the zinc finger of UBR4. Picolinic acid is a pyridine carboxylate metabolite of the type-2 amino acid tryptophan of the N-end rule pathway (Law et al., 2014) (Fig. 7A). To determine whether picolinic acid binds UBR4, we assessed the ability of picolinic acid to pulldown full-length 570 kDa UBR4 from HEK293 cell extracts. In comparison, we performed an analogous assay using biotin-conjugated 11-mer peptides bearing Nt-Arg (type 1), Nt-Phe (type 2) and Nt-Gly (stabilizing). UBR4 was pulled down by picolinic acid, the Arg-containing peptide and the Phe-containing peptide but not by the Gly-containing peptide (Fig. 7B,C; Fig. S6A,B). Although we do not exclude the possibility of off-target effects, these results suggest that picolinic acid binds UBR4.
Next, to assess the efficacy of picolinic acid, we treated HeLa cells transiently expressing EEA1–EGFP and RAB5–mRFP with this compound. This treatment markedly downregulated the formation of EEA1+ as well as RAB5+ EEs (Fig. 7D,E). The inhibitory effect of picolinic acid on endosomal maturation was reversible (Fig. 7F–I). Specifically, although transfected EGFP signals were not affected by picolinic acid (Fig. 7F), EEA1+ and RAB5+ puncta became smaller in a time-dependent manner and were readily restored when picolinic acid was removed (Fig. 7G–I). The disrupted endosomal maturation observed in picolinic acid-treated cells was not caused by destabilization of actin filaments and microtubules (Fig. S6C). These results provide a means to modulate endosomal processes by targeting UBR4.
We have previously identified two classes of N-recognins that can recognize the Nt-Arg residues of arginylated proteins via the UBR box (UBR1, UBR1, UBR4 and UBR5) or the ZZ domain (SQSTM1) (Cha-Molstad et al., 2015; Tasaki et al., 2005). Studies by us and others have shown that UBR1 and UBR2 mediate the ubiquitylation of short-lived regulators carrying N-degrons as well as soluble misfolded proteins, leading to their proteasomal degradation into short peptides (Eisele and Wolf, 2008; Lee et al., 2005; Tasaki et al., 2005). More recently, we have shown that the autophagic receptor SQSTM1 binds the Nt-Arg of arginylated ER-derived proteins and mediates their lysosomal degradation along with misfolded proteins (Cha-Molstad et al., 2015). Compared with other N-recognins, little is known about UBR4 and UBR5. In this study, we characterized the functions and mechanisms of action of UBR4. We show that UBR4 is associated with the EVs in part through the binding to calmodulin (Figs 3H, 8A). Functionally, UBR4 is not critical for endocytosis (i.e. the internalization of extracellular materials such as HRP and TAT) (Fig. 2). However, the majority of the EVs are arrested at a premature stage, possibly in the course of homotypic fusion with each other (Fig. 8B). Overall, our results indicate that UBR4 is required for the biosynthesis of EEs (Fig. 8B). This conclusion is supported by several lines of evidence: (1) Ubr4−/− MEFs contained a significantly reduced number of mature EEs associated with the excessive accumulation of arrested EVs with aberrant morphology (Fig. 4A–D), (2) the treatment of an UBR4 inhibitor markedly downregulated the formation of EEA1+ as well as RAB5+ EEs (Fig. 7D–I), and (3) UBR4 is required for endosomal trafficking of ligand–receptor complexes internalized from the plasma membrane (Fig. 5C,F). Additionally, the disrupted endosomal maturation and trafficking observed in UBR4-deficient MEF cells were not caused by destabilization of actin filaments and microtubules (Fig. S5E). Both picolinic acid and W7 also did not damage these cytoskeletal components (Fig. S6C). Given that UBR4 is an N-recognin, it is tantalizing to speculate that the function of UBR4 could be modulated by the Nt-Arg residues of arginylated proteins. In this model, ATE1 R-transferase conjugates the amino acid arginine to the N-termini of proteins. The resulting Nt-Arg residue acts as an activating ligand that binds the UBR box of UBR4, enabling UBR4 to bind Ca2+-calmodulin and to be associated with endosomes. Consistent with this model, we found that endosomal processes are impaired in ATE1-deficient cells (data not shown), although the direct link between ATE1 and UBR4 needs further investigation.
The majority of cell surface proteins and various extracellular materials are endocytosed and degraded by endosomal hydrolases in the EEs, LEs, MVBs and amphisomes (Pillay et al., 2002). Several isolated studies suggest that the overall amounts of cargos that are digested by endosomal hydrolases that might match or even exceed those by lysosomal hydrolases in normal conditions (Abrahamson and Rodewald, 1981; Authier et al., 1996; Berg et al., 1995; Besterman and Low, 1983; Maxfield, 2014; Pillay et al., 2002; Steinman et al., 1983). Despite the importance of the endosomal proteolytic system, little attention has been paid to its functions and mechanisms, for example, in the turnover of cell surface proteins. In principle, internalized cell surface proteins carried in the EVs are either recycled back to the plasma membrane or move forward to the degradative endosomal pathway. Although the relative portion and rate depend on cell types and physiological states, it is estimated that ∼50% of the plasma membrane-associated proteins and 5–30% of cellular volumes are internalized in every hour (Maxfield, 2014; Steinman et al., 1983). In this study, we show that UBR4 plays a role in endosomal proteolytic flux. This conclusion is supported by the findings that (1) UBR4 marks and is required for endosomal proteolysis in the YS of mouse embryos (Fig. 1), (2) the degradation of the model substrate DQ-BSA is markedly impaired in Ubr4−/− MEFs as compared with +/+ cells (Fig. 6A,B), and (3) UBR4 is required for the biogenesis of EEs (Fig. 4).
Our earlier work has shown that UBR4-knockout mice die at E9.5–10.5 with vascular defects in the visceral YS that are associated with autophagic induction in embryotic tissues as well as cultured cells (Tasaki et al., 2013). The vascular development in UBR4-lacking YSs is arrested during angiogenic remodeling of primary capillary plexus. To date, the mechanisms underlying the YS phenotypes remain unknown. In this study, we show that UBR4 is prominently expressed in the YS endoderm, a specialized tissue that has a high level of endosomal proteolysis (Fig. 1B,C). In the YS, UBR4 marks EEA1+ EEs that carry internalized extracellular proteins provided by the mother. The cargos are partially degraded within EEs and amphisomes, and eventually converted into amino acids (Kim et al., 2013). Notably, our immunostaining analyses on the sections of embryonic tissues show that EEA1+ EEs are markedly downregulated in the YS of Ubr4−/− embryos (Fig. 1D). Given that the resulting amino acids are the major source for protein translation during embryogenesis, defective endosomal proteolysis in the YS may contribute to developmental abnormalities and arrest. One question that remains to be addressed is the functional relationship between endosomal defects (this study) and autophagic induction observed in UBR4-deficient cells and animal tissues (Kim et al., 2013). Because autophagy is induced by glucose and/or amino acid starvation (Mizushima and Komatsu, 2011), we speculate that endosomal defects in UBR4-deficient cells cause insufficient proteolysis of extracellular cargos for nutrient supply, leading to starvation. Further investigation is needed to validate this model.
Human UBR4 is a 570 kDa microtubule-binding protein that is prominently expressed in neuronal cells (Parsons et al., 2015; Tasaki et al., 2005). Studies have implicated UBR4 in various biological processes, including neurodegeneration (Parsons et al., 2015; Tasaki et al., 2005), auxin transport, trafficking via microvesicles and ectosomes (Gil et al., 2001), synaptic transmission (Sekelsky et al., 1999; Yager, 2001), the interaction with various viral proteins across diverse strains (Tripathi et al., 2015), proteasomal degradation of short-lived regulators (Szalmas et al., 2017; White et al., 2016) and E7-mediated cellular immortalization and transformation activities (DeMasi et al., 2005, 2007; Huh et al., 2005; White et al., 2012). Despite rather extensive studies, however, little is known about the cellular functions of UBR4 that explain these seemingly unrelated observations. Given our findings showing that UBR4 is a key modulator in the biogenesis of EEs and endosomal proteolysis, the aforementioned functions of UBR4 in various biological systems may need to be re-evaluated in the light of endosomal processes.
MATERIALS AND METHODS
Rabbit polyclonal anti-human UBR4 antibody against residues 3755–4160 (1:400) was used for immunohistochemical analysis of YS and embryos (Shim et al., 2008). Rabbit polyclonal anti-UBR4 antibody (Bethyl Laboratories, IHC-00640, 1:300) was used for immunostaining of cultured cells. Other antibodies are as follows: mouse monoclonal anti-APPL1 (Santa Cruz Biotechnology, SC-271901, 1:100), rabbit polyclonal anti-EEA1 (Cell Signaling, 2411, 1:100), mouse monoclonal anti-EGFR (Santa Cruz Biotechnology, SC-374607, 1:200), rabbit polyclonal anti-LAMP1 (Sigma, L1418, 1:100), rabbit polyclonal anti-RAB5 (Cell Signaling, 3547, 1:400), mouse monoclonal anti-RAB7 (Sigma, R8779, 1:1300), rabbit polyclonal anti-V5 (Sigma, V8137, 1:500). For immunoblotting analysis, mouse monoclonal anti-β actin (Sigma, A1978, 1:10,000), rabbit polyclonal anti-UBR4 (Abcam, AB86738, 1:100) and mouse monoclonal anti-GAPDH (Sigma, G8795, 1:20,000) antibodies were used. The following secondary antibodies are from Invitrogen: Alexa Fluor 488-conjugated goat anti-rabbit IgG (A11034, 1:200), and Alexa Fluor 555-conjugated goat anti-mouse IgG (A21424, 1:200).
Plasmids and other reagents
mRFP–RAB5 and EGFP–calmodulin were Addgene plasmid #14437 (Vonderheit and Helenius, 2005) and Addgene plasmid #47602 (deposited by Ari Helenius and Emanuel Strehler, respectively). EGFP–EEA1 and TagRFP–EEA1 were Addgene plasmid #42307 (Lawe et al., 2000) and #42635 (Navaroli et al., 2012), respectively (deposited by Silvia Corvera). GFP- and V5-tagged UBR4 plasmids were prepared as described in the previous studies (Tasaki et al., 2013, 2005). These plasmids were transiently transfected with Lipofectamine 3000 reagent following the manufacturer's instructions (Invitrogen). DQ-Red BSA was purchased from Invitrogen (D12051). Normal goat serum (ab7481) was obtained from Abcam. Hoechst 33342 (H21492) was obtained from Invitrogen. 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI; D8417) and nocodazole (M1404) were obtained from Sigma. W7 (681629) and latrunculin A (428021) were obtained from Calbiochem. Synthetic TAT–FITC peptide was purchased from Anygen (Kwangju, South Korea). The sequence was FITC–GGGGYGRKKRRQRRR-NH2. Vectashield antifade mounting medium (H1000) was from Vector laboratories. All other chemicals were reagent grades from Sigma or Merck.
We have previously constructed UBR4-knockout mice, in which the UBR box, a substrate recognition domain for destabilizing N-terminal residues, was replaced with internal ribosome entry site (IRES)-translated tau-lacZ (Tasaki et al., 2013). Animal studies were conducted according to protocols (SNU130604-2-10) approved by the Institutional Animal Care and Use Committee at Seoul National University.
Primary MEFs were established from +/+ and Ubr4−/− embryos at E8.5. The embryos were minced by pipetting in Iscove's modified Dulbecco's medium (IMDM; Gibco, 31980-022), 15% fetal bovine serum (FBS; Hyclone), 0.1 mM non-essential amino acids (Invitrogen), 0.1 mM β-mercaptoethanol and 100 U/ml penicillin-streptomycin (Gibco, 15140-148). The cells were seeded on the gelatinized 35 mm culture dish. Immortalized cell lines were established from primary MEFs through crisis-mediated immortalization over 10 passages (Tasaki et al., 2013). CCL2 HeLa and HEK293 cell were purchased from the American Type Culture Collection (ATCC). All the cell lines were determined to be negative in a mycoplasma test using a MycoAlert detection kit (Lonza, LT07-118). HeLa and HEK293 cells were cultured in Dulbecco's modified Eagle's medium (DMEM; Gibco, 10566016). The medium was supplemented with 10% FBS and 100 units/ml penicillin-streptomycin. All the culture plates and the cell lines were maintained at 37°C and 5% CO2 in a humidified incubator.
RNA interference assay
Reagents for siRNA silencing were purchased from Life Technologies. Transfection was performed according to the manufacturer's protocol. Briefly, cells were transfected with either siControl (cat. #4390843), EEA1 siRNA (Origene, cat. #SR30003), or UBR4 siRNA (cat. #4392420, ID #23628) at a final concentration of 10 nM using Lipofectamine RNAiMAX reagent (Invitrogen, 13778150). At ∼48 h after siRNA silencing, cells were harvested for immunoblotting and immunocytochemical analyses. The sequences of UBR4 siRNAs are 5′-GCCUGUUCGAAAGCGCAAA-3′ (sense) and 5′-UUUGCGCUUUCGAACAGGC-3′ (antisense).
Cells were washed with cold phosphate-buffered saline (PBS) and lysed using RIPA buffer (50 mM Tris-HCl, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate and 0.1% SDS) containing freshly prepared protease inhibitor cocktail (Sigma, P8340). Lysates were centrifuged at 12,000 g for 20 min at 4°C, and the supernatants were used for immunoblotting. Protein concentrations were measured using the BCA protein assay kit (Pierce, 23225). The samples were diluted with 2× Laemmli sample buffer [65.8 mM Tris-HCl, pH 6.8, 26.3% (w/v) glycerol, 2.1% SDS, 0.01% bromophenol blue, Bio-Rad, 161-0737] or in lithium dodecyl sulfate (LDS) sample buffer (Invitrogen, NP0007) with a reducing reagent, followed by heating for 10 min at 70°C. Whole-cell lysates were separated by SDS-PAGE and transferred onto polyvinylidene difluoride membranes (Millipore, IPVH00010). Blocking was performed using TBS-T [20 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 0.05% (v/v) Tween 20] containing 1% BSA for 1 h at room temperature, and the membrane was incubated with antibodies diluted with the blocking solution overnight at 4°C.
Histology and immunohistochemistry
For histological analysis, embryos were fixed overnight at 4°C in 4% paraformaldehyde (PFA) in PBS, pH 7.4. Fixed embryos were gradually dehydrated with 70%, 90% and 100% ethanol, followed by immersion in Neo-Clear (Millipore, 65351). Tissues were embedded in paraffin wax at 58°C and sectioned transversely or sagittally with 7 μm thickness. Immunostaining of paraffin sections and whole-mount immunohistochemical staining of embryos were performed as described previously (Tasaki et al., 2013). Paraffin-embedded slides were freshly treated with Neo-Clear twice for 10 min each, followed by gradual rehydration in ethanol (100%, 90%, 80% and 70%; 6 min each) and water for 20 min. For immunohistochemistry, the slides were treated with blocking solution (5% normal goat serum and 0.2% Triton X-100 in PBS) for 1 h and incubated with primary antibodies and subsequently with secondary antibodies. Confocal images were taken with a Zeiss LSM 700 laser-scanning confocal microscope equipped with Zeiss C-Apochromat 60× (1.2 NA) and 40× (1.2 NA) water immersion lens and analyzed using ZEN (black edition) 2012 SP5 software (Zeiss). Using the ZEN software, z-stacks of images covering the entire cell thickness were acquired and projected maximally. Image processing and annotation was performed with Adobe Photoshop, Adobe Illustrator and Fiji software (Schindelin et al., 2012).
Immunocytochemistry of cultured cells
Three 22 mm2 coverslips per well were placed in six-well plates, followed by incubation of diluted poly-L-lysine (1:10 in sterile deionized water; Sigma, P8920) for 30 min at room temperature. After washing once, the plate was exposed under an UV radiation lamp overnight in a fume hood. Cells were cultured in the plate for further experiments. Cells were fixed in 4% PFA in PBS, for 30 min at room temperature. After washing twice with PBS, the cells were incubated for 1 h in blocking solution (5% normal goat serum and 0.3% Triton X-100 in 0.1 M PBS). After blocking, immunohistochemical processes were conducted as described above. N-SIM images were taken with a Nikon Eclipse Ti inverted microscope (Nikon, Tokyo, Japan) equipped with a Nikon Apo TIRF 100× (1.49 NA) oil-immersion lens. The SIM images were reconstructed from the raw images through the Nikon NIS-element (ver. 3.22.10).
TAT–FITC peptide internalization assay
TAT peptide incubation was performed as previously described with minor modification (Vivès et al., 1997). Cells were washed twice with PBS and preincubated in Opti-MEM (Gibco, 31985070) for 30 min, followed by TAT–FITC peptide treatment for the indicated times. Cells were fixed with 4% PFA for further experiments. Immunocytochemical processes were conducted as described above.
Horseradish peroxidase uptake analysis
HRP uptake analysis was performed as previously described with minor modifications (Li et al., 1995; Li and Stahl, 1993; Stoorvogel, 1998). After washing cells with pre-warmed culture medium, cells were incubated in IMDM containing 5 mg/ml HRP and 1% bovine serum albumin (BSA) for 30 min. The uptake was stopped by washing cells five times with ice-cold PBS-containing 0.1% BSA. After the final wash, cells were scraped into 1 ml of ice-cold PBS and pelleted at 800 g for 3 min at 4°C. The cell pellet was lysed in 400 μl of ice-cold PBS containing 0.1% Triton X-100. The lysate was assayed for horseradish peroxidase activity. The enzyme assay was conducted in a 96-well microplate using 1-Step™ Ultra TMB-ELISA substrate solution as the chromogenic substrate. The reaction was started by adding 1 μl of the lysate to 100 μl of 1-Step™ Ultra TMB-ELISA Substrate Solution (Thermo Scientific 34028). The reaction was conducted at room temperature for 20 min and stopped by adding 100 μl of 2 M H2SO4. Quantified by measuring the optical density at 425 nm (OD425nm) in a Tecan microplate reader (Tecan, Sunrise™). Protein content was determined by the Thermo Scientific Pierce BCA assay according to the manufacturer's instructions.
UBR4-V5 stable HEK293 cells were fixed in 2% PFA in PBS, pH 7.4, for 1 h at room temperature and washed with PBS. Fixed cells were collected by scraping and resuspended in 3% gelatin. Gelatin was solidified on ice, and cells embedded in gelatin were fixed again with 2% PFA for 15 min. After cryoprotection with 2.3 M sucrose with PVP solution overnight, samples were frozen in liquid nitrogen and were trimmed into 0.5 mm cubes. The frozen cell blocks were sectioned with a cryo-microtome (Leica EM Crion) in 70 nm sections and collected on 200 mesh grids. The primary antibody against V5 was diluted in 0.1 M PBS with 0.5% BSA, 0.15% glycine according to the established condition for immunohistochemistry. A donkey anti-rabbit IgG antibody labeled with 12 nm gold beads (Jackson ImmunoResearch) was diluted to 1:25 before use. After immunostaining, sections were incubated with 2.5% glutaraldehyde for 10 min and with 2% neutral uranyl acetate for 7 min. Sections were further processed with 4% uranyl acetate and methyl cellulose for contrasting and drying, respectively. After drying, samples were recorded using a JEOL JEM1011 TEM with a high-resolution AMT digital camera (Peabody, MA).
Fractionation of proteins
Crude extracts were prepared by homogenizing +/+ and Ubr4−/− MEFs in a hypotonic solution. Cellular proteins were separated into cytosolic (S150) and microsomal (P150) fractions using differential centrifugations and solubilization (Tasaki et al., 2013). Cells were harvested from 5 of 15 cm dishes by trypsin-EDTA solution, followed by three washes with PBS. Cell pellets were washed once with 5 packed cell volumes (pcv) of hypotonic buffer (10 mM HEPES pH 7.9, 1.5 mM MgCl2 and 10 mM KCl) supplemented with protease inhibitor cocktail (Sigma), 20 mM NaF, 10 mM sodium orthovanadate and 1 mM DTT. Cells were suspended with 2 pcv of hypotonic buffer with supplements, and cells were incubated for 10 min on ice. The expanded cells were homogenated with a dounce homogenizer, and a soluble fraction was collected after a centrifugation at 15,000 g for 5 min. The pelleted fraction including the nucleus, mitochondria and plasma membranes, was solubilized with RIPA buffer (10 mM Tris-HCl, pH 7.4, 1% NP-40, 0.1% sodium deoxycholate, 0.1% SDS, 0.15 M NaCl and 1 mM EDTA) to yield a soluble fraction (P15S) and an insoluble fraction (P15I). The soluble fraction after dounce homogenization and a centrifugation was further fractionated by an ultra-centrifugation at 150,000 g for 1 h. The supernatant represents a cytosol (S150) and the pelleted membranes represent microsomes including the ER (P150). The protein concentration was quantified, using the BCA protein assay and each fraction was kept frozen at −80°C until used for SDS-PAGE.
EGFP–EEA1 and mRFP–RAB5 puncta analysis
HeLa cells were transiently transfected with either 1 µg/ml of EGFP–EEA1 or mRFP–RAB5 plasmids using Lipofectamine 3000 reagent (Invitrogen). Following the transfection, cells were cultured for 24 h and trypsinized for replating on coated slides. At 24 h after replating, cells were treated with picolinic acid and fixed with 4% PFA for further experiments at the indicated times. After PBS washing, immunocytochemical processes were conducted as described above.
Pulldown assay of UBR4 and calmodulin
The pulldown assay was performed as previously described with minor modifications (Tasaki et al., 2005). HEK293 cells were transfected with UBR4–GFP and GUS–GFP plasmids using Lipofectamine LTX and Plus reagent (Invitrogen) and harvested 48 h after transfection. Cells were lysed in 1% Nonidet-P40 (NP-40), 0.15 M NaCl, 1 mM EDTA and 10 mM Tris-HCl (pH 7.5). The lysates were incubated with biotin-conjugated goat polyclonal anti-GFP antibody (Abcam, ab6658; 1:100) in binding buffer A (0.1% Nonidet P-40, 10% glycerol, 0.15 M KCl, and 20 mM HEPES, pH 7.9) and mixed with streptavidin–Sepharose beads (Amersham Bioscience). The beads were pelleted by centrifugation at 2400 g for 30 s, washed three times with binding buffer A and resuspended in SDS-PAGE sample buffer, and heated and analyzed by immunoblotting with rabbit monoclonal anti-calmodulin antibody (Abcam, ab45689; 1:1000).
Synthesis of biotinylation of picolinic acid
Picolinic acid was biotin-conjugated using EZ-link pentylamine-Biotin (Thermo, 21345). 0.18 mM picolinic acid was added to the mixture including 0.22 mM N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDCI.HCl), 6.2 mM hydroxybenzotriazole (HOBt), 0.54 mM N,N-diisopropylethylamine (DIPEA), and 2 ml DMF. After 10 min, 0.15 mM 5-(biotinamido)pentylamine was added to the mixture with continued stirring. The reaction was quenched by the addition of water and then ethyl acetate was added. It was washed with saturated citric acid to eliminate unreacted amine derivatives followed by sequential washing with sodium bicarbonate to remove unreacted acid derivatives followed by washing with saturated brine (26% NaCl in water). The combined organic layers were dried over anhydrous Na2SO4 and concentrated under vacuum. The crude residue was purified using silica gel column chromatography (methanol:dichloro methane ratio of 1:9) to yield the biotinylated picolinic acid as a white solid. The structure of the product biotinylated picolinic acid was confirmed with liquid chromatography-mass spectrometry (LC-MS) (Fig. S5A,B). Its mass spectrum revealed a molecular ion peak at 434.32 [M+H]+ corresponding to the molecular formula C21H31N5O3S, confirming the structure as biotinylated picolinic acid.
X-peptide and biotinylated picolinic acid pulldown assay
The X-peptide and biotinylated picolinic acid pulldown assays were carried out as previously described with minor modifications (Kwon et al., 2003). We used a set of 12-mer X-nsP4 peptides (X-IFSTIEGRTYK–biotin) bearing N-terminal arginine (type 1), phenylalanine (type 2) and glycine (stabilizing control) residues, which is known as a N-end rule substrate. For cross-linking with resin, the biotin-conjugated picolinic acid and peptides were mixed with high-capacity streptavidin agarose resin (Thermo Scientific, 20361) with a ratio of 0.5 mg peptide per 1 ml settled resin and incubated on a rotator at 4°C overnight. After washing five times with PBS, the peptide–bead conjugates were diluted with PBS at 1:1 ratio. To prepare protein extracts, cells were collected by centrifugation (1000 g for 5 min) and lysed by freezing and thawing at least ten times in hypotonic buffer (10 mM KCl, 1.5 mM MgCl2, and 10 mM HEPES, pH 7.9) with a protease inhibitor mix (Sigma, P8340). After spinning down with centrifugation in 14,300 g at 4°C for 15 min, the amount of protein was quantified using a BCA protein assay kit (23227, Thermo Scientific). Total proteins (300 μg) diluted in 250 μl binding buffer (0.05% Tween-20, 10% glycerol, 0.2 M KCl, and 20 mM HEPES pH 7.9) were mixed with 50 μl peptide–bead resin and incubated at 4°C overnight on a rotator. The protein-bound beads were collected by centrifugation at 2400 g for 1 min and washed five times with binding buffer. The beads were resuspended in 25 μl SDS sample buffer, heated at 70°C for 10 min, and subjected to SDS-PAGE and immunoblotting.
Fluorescence labeling of EGF
Labeling with EGF was performed as previously described with minor modification (Mesaki et al., 2011). Cells were serum-starved for 1 h and stimulated with 1 μg/ml EGF conjugated to Alexa Fluor 555 (Invitrogen, E35350) in DMEM-HEPES buffer containing 1% BSA on ice for 1 h and washed with cold PBS. The samples were incubated in pre-warmed medium for the indicated times, followed by 4% PFA fixation at the appropriate time point. After fixation, we followed the same procedures as we described above (immunocytochemistry). A quantitative analysis was performed as previously described but with a manual calculation of the distance from the plasma membrane (Pike et al., 2017). Confocal images were manually segmented based on the distance from the plasma membrane. Each band has a width of 1 μm.
EGF–biotin dot blot assay
Labeling with EGF was performed as previously described with minor modifications (Mesaki et al., 2011). Cells were serum-starved for 1 h and stimulated with 1 μg/ml EGF conjugated with biotin (Invitrogen, E3477) in DMEM-HEPES buffer containing 1% BSA on ice for 1 h and washed with cold PBS. The samples were incubated in pre-warmed medium for the indicated times, followed by lysis. The internalized EGF-biotin was analyzed by dot blotting. For dot blot experiments, isolated EGF-biotin samples were spotted onto a nitrocellulose membrane and incubated with HRP-conjugated goat anti-biotin secondary antibody (Cell Signaling, cat#7075, 1:1000).
All numerical data are presented as means±s.e.m. or mean percentages±s.e.m. Western blots shown are representative of three or more independent experiments. Comparisons among treatment groups were performed with one-way analysis of variance (ANOVA) and Tukey test or Dunnet's multiple comparison as a post hoc comparison. Instances involving only two comparisons were evaluated with a two-tailed t-test. Statistical significance was accepted if the null hypothesis was rejected with P<0.05. All statistical analyses were determined using Prism 7.0a software (GraphPad, La Jolla, CA).
We thank the laboratory members of Y.T.K. and B.Y.K. for critical discussions.
Conceptualization: S.T.K., Y.J.L., T.T., Y.T.K.; Methodology: S.T.K., Y.J.L., T.T., S.R.M., J.H., M.J.K., S.G., E.C.Y.; Software: S.T.K., Y.J.L., M.J.K.; Validation: S.T.K., Y.J.L., T.T., S.R.M., J.H., M.J.K., S.G., E.C.Y., B.Y.K., Y.T.K.; Formal analysis: S.T.K., Y.J.L., T.T., S.R.M., J.H., M.J.K., Y.T.K.; Investigation: S.T.K., Y.J.L., T.T., S.R.M., M.J.K., Y.T.K.; Resources: S.G., E.C.Y.; Data curation: S.T.K., Y.J.L., T.T., S.R.M., J.H., M.J.K., S.G.; Writing - original draft: S.T.K., T.T., B.Y.K., Y.T.K.; Writing - review & editing: S.T.K., Y.J.L., Y.T.K.; Visualization: E.C.Y., B.Y.K.; Supervision: S.T.K., Y.J.L., T.T., E.C.Y., B.Y.K., Y.T.K.; Project administration: S.T.K., Y.J.L., B.Y.K., Y.T.K.; Funding acquisition: Y.J.L., T.T., E.C.Y., B.Y.K., Y.T.K.
This work was supported by grants from the Seoul National University (SNU) Nobel Laureates Invitation Program, the National Research Foundation of Korea (NRF) funded by the Ministry of Science and ICT (MSIT) (NRF-2016R1A2B3011389 to Y.T.K., NRF-2015M3A9B6073835 to E.C.Y., NRF-2014M39Ab5073938 to B.Y.K., and NRF-2015-Global PhD Fellowship to Y.J.L.), the Brain Korea 21 PLUS (to SNU), Seoul National University Hospital (to Y.T.K.), the Bio and Medical Technology Development Program (project no. 2012M3A9B6055305 to B.Y.K.) through the Ministry of Science, ICT and Future Planning, the R&D Convergence Program (CAP-16-03-KRIBB) of National Research Council of Science and Technology (NST), Korea Research Institute of Bioscience and Biotechnology (KRIBB) Research Initiative Program (NRF-2014M39Ab5073938 to B.Y.K.), and a grant for Promoted Research from Kanazawa Medical University (S2016-13, S2017-6 to T.T.) and by Japan Society for the Promotion of Science (JSPS) KAKENHI grant numbers JP25430119 and JP18K06119 (to T.T.).
The authors declare no competing or financial interests.