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Research Article
A cascade of ER exit site assembly that is regulated by p125A and lipid signals
David Klinkenberg, Kimberly R. Long, Kuntala Shome, Simon C. Watkins, Meir Aridor
Journal of Cell Science 2014 127: 1765-1778; doi: 10.1242/jcs.138784

ABSTRACT

The inner and outer layers of COPII mediate cargo sorting and vesicle biogenesis. Sec16A and p125A (officially known as SEC23IP) proteins interact with both layers to control coat activity, yet the steps directing functional assembly at ER exit sites (ERES) remain undefined. By using temperature blocks, we find that Sec16A is spatially segregated from p125A-COPII-coated ERES prior to ER exit at a step that required p125A. p125A used lipid signals to control ERES assembly. Within p125A, we defined a C-terminal DDHD domain found in phospholipases and PI transfer proteins that recognized PA and phosphatidylinositol phosphates in vitro and was targeted to PI4P-rich membranes in cells. A conserved central SAM domain promoted self-assembly and selective lipid recognition by the DDHD domain. A basic cluster and a hydrophobic interface in the DDHD and SAM domains, respectively, were required for p125A-mediated functional ERES assembly. Lipid recognition by the SAM–DDHD module was used to stabilize membrane association and regulate the spatial segregation of COPII from Sec16A, nucleating the coat at ERES for ER exit.

INTRODUCTION

The coat protein complex II (COPII) is composed of the small GTPase Sar1 and the cytosolic Sec23/24 and Sec13/31 protein complexes (Antonny and Schekman, 2001). These cytosolic proteins assemble on ER membranes at ER exit sites (ERES) to select cargo proteins destined for exit and to deform membranes into buds and vesicles released from the ER. The coat inner layer, Sar1–Sec23/24, binds acidic lipids and presents multiple binding sites for short peptide sequences, ER exit motifs, thus sorting cargo for incorporation into budded vesicles (Aridor et al., 2001; Aridor et al., 1998; Kuehn et al., 1998; Miller et al., 2003). The outer layer, which is composed of the Sec13/31 complex, forms an ancestral coat element 1 (ACE1) (Fath et al., 2007) that is recruited onto the inner layer and polymerizes to form a hedral cage (Stagg et al., 2008). Provision of a catalytic arginine residue by Sec23 and optimization of the position of catalytic residues within the Sar1 GTP-binding site by Sec31 both enhance GTP hydrolysis. A vesicle neck is constricted by the activity of Sar1, leading to GTPase-dependent vesicle release (Bielli et al., 2005; Lee et al., 2005; Long et al., 2010). The minimal set of COPII coat proteins recapitulates basic cargo selection and vesicle formation activities (Antonny et al., 2001; Matsuoka et al., 1998). However, COPII activities measured in minimal reactions are controlled by interacting proteins that couple sorting and budding activities with physiological biosynthetic demands (Zanetti et al., 2011). For example, the adaptation of ERES activities to changes in cargo load is mediated by the activities of mammalian Sec16 (Sec16A) and phosphoinositide 4-kinase (PI4K) IIIα (Farhan et al., 2008). PI4KIIIα generates phosphatidylinositol 4-phosphate (PI4P) on ER membranes where the dynamic generation of PI4P supports ERES assembly and ER export (Blumental-Perry et al., 2006; Farhan et al., 2008). In yeast, Sec16p interacts with all COPII subunits, enhances their assembly and potentiates COPII vesicle budding from the ER. The mechanisms by which these two activities, Sec16–COPII interactions and PI4P generation, control COPII budding at ERES remain to be defined. A link between Sec16 and lipid signals has been previously observed. Sec16p substitutes for acidic lipids in promoting COPII recruitment on synthetic liposomes (Supek et al., 2002). However, Sec16p hinders the functional linkage between the coat inner and outer layers, leading to inhibition of Sec31-enhanced GTPase activity of Sar1. The C-terminal domain of Sec16p, which binds Sar1–Sec23/24, inhibits the recruitment of the outer layer onto Sec23/24 (Yorimitsu and Sato, 2012; Kung et al., 2012; Whittle and Schwartz, 2010). A mechanism that allows for the reversal of Sec16-mediated inhibition of coat linkage to support ER exit remains to be defined.

p125A (officially known as SEC23IP) binds Sec31 in cytosol and is recruited to membranes where it also binds Sec23, thus uniquely linking the two coat layers (Ong et al., 2010). Knockdown and overexpression studies demonstrate that p125A is required for ERES organization (Iinuma et al., 2007; Shimoi et al., 2005) and cargo export from the ER (Ong et al., 2010). p125A belongs to a family of PA-preferring phospholipase A1 enzymes (Mizoguchi et al., 2000; Shimoi et al., 2005; Tani et al., 1999). We hypothesize that p125A uses lipid signals to reverse Sec16-inhibited layer coupling and to direct COPII assembly at ERES. We now identify a molecular cascade in which Sec16 is spatially segregated from p125A-COPII-coated ERES prior to exit. p125A functions to decode lipid signals such as PI4P and to support the displacement of Sec16A while directing COPII nucleation at ERES, which promotes ER exit.

RESULTS

p125A is recruited with COPII to PI4P-enriched liposomes

PI4P production at the ER is required for functional ERES organization (Blumental-Perry et al., 2006; Farhan et al., 2008). We examined whether similar dependency is observed when the recruitment of mammalian COPII from cytosol is measured on synthetic defined membranes. PI4P-containing or control liposomes were incubated with cytosol in the presence of constitutively active (Sar1H79G, termed Sar1-GTP) or inactive (Sar1T39N, Sar1-GDP) Sar1 proteins, and isolated by floatation in sucrose gradients. Effective COPII recruitment from cytosol to liposomes (measured by Sec23) was induced by Sar1 activation and required PI4P (fractions 1-3; Fig. 1A). The Sar1-dependent recruitment of the outer COPII cage (Sec31) was directed onto distinct (albeit not all) Sec23-containing fractions of PI4P-containing liposomes (Fig. 1B). The results are in agreement with previous studies monitoring the recruitment of purified yeast COPII proteins to synthetic membranes (Matsuoka et al., 1998) and support a role for PI4P in coat recruitment, yet they suggest that the link between inner and outer coat layers might be regulated. p125A regulates ERES assembly and ER export (Ong et al., 2010; Shimoi et al., 2005), associates with Sec31 in cytosol and binds Sec23 using separate segments on its N terminus (Ong et al., 2010). Immunodepletion of p125A from cytosol by immunoprecipitation, previously shown to co-deplete Sec31 (Ong et al., 2010), did not affect the Sar1-dependent recruitment of COPII inner layer Sec23/24 to ER membranes (Fig. 1C). The recruitment of p125A to PI4P-containing liposomes mirrored that of Sec31 (Fig. 1D). Thus, outer layer-associated p125A may link the two coat layers following initial recruitment of Sar1–Sec23/24 onto ER membranes to control the progression of ERES assembly.

Fig. 1.
Fig. 1.

Sar1-dependent COPII and p125A recruitment requires PI4P. (A,B) Sar1-dependent COPII recruitment to floated liposomes is dependent on PI4P. Active (Sar1-GTP) or inactive (Sar1-GDP, both tested at 1 µg, 50 µl final volume) were incubated with rat liver cytosol and synthetic large unilamellar vesicles (LUV, 400 µM) composed of 45% phosphatidylcholine (PC), 35% phosphatidylethanolamine (PE), 10% phosphatidylserine (PS) and 10% cholesterol or 35% PC, 35% PE, 10% PS, 10% cholesterol and 10% PI4P at 26°C for 1 hour and floated onto a sucrose gradient (Bielli et al., 2005). Fractions were analyzed by western blot with antibodies against Sec23 (at 1:10,000 dilution) (A) or hSec31a (1:500) (B) (floated fractions as labeled). (C) Recruitment of Sec23 to ER membranes is not affected by depletion of p125A. p125A or SNX9 (control) was depleted from rat liver cytosol by immunoprecipitation and p125A depletion was verified using western blot, as indicated. Sar1-GTP-dependent Sec23 recruitment from control or p125A-depleted cytosol to ER microsomes was monitored. Sec23 was recruited by Sar1-GTP (50 ng, 500 ng and 1 µg, final volume 60 µL) in a dose-dependent manner in the absence (lanes 1–4) or presence (lanes 5–8) of p125A. (D) Floated PI4P-containing liposomes (fractions 1–3) from a COPII recruitment reaction (described in A) were probed for p125A (1:5000), Sec23 and Sec31. (E) Transiently expressed EGFP-tagged p125A colocalizes predominantly with hSec31a (as observed with the endogenous p125A protein) at ERES and was juxtaposed to ERGIC (ERGIC53) or cis-Golgi compartments (GPP130). Arrowheads indicate EGFP-p125A localizing with Sec31 (antibody dilution 1:100), adjacent to membranes containing ERGIC53 (1:100) and gpp130 (1:500). Scale bars: 5µm.

The DDHD and SAM domains cooperate to support lipid recognition in vitro and binding of PI4P-rich membranes in cells

We hypothesized that p125A recognizes selective lipid signals to regulate ERES assembly. To test this, we first examined the role and specificity of selected p125A domains in lipid-signal binding. p125A binds membranes using its C terminus, which contains a DDHD domain and a sterile α motif (SAM). DDHD domains are ∼180 residues long (residues 779–989 in p125A) and contain four conserved residues (DDHD) that can form a putative metal binding site typically found in phosphoesterase domains. DDHD domains are found in retinal degeneration B proteins, in the N-terminal domain-interacting receptor (Nir1-3), where Nir2 functions as a phosphatidyl inositol (PI)-transfer protein (Litvak et al., 2005), and in the p125A-containing PLA1 phospholipase protein family (Sato et al., 2010; Shimoi et al., 2005; Yamashita et al., 2010). In the context of many multidomain proteins, SAM domains are common protein-protein interaction motifs that modulate function through their ability to homo- or hetero-associate. Several forms of SAM domains have also been shown to polymerize into larger functional structures (Qiao and Bowie, 2005). We analyzed the role of p125A SAM (residues 643–704) and DDHD domains in vivo and in vitro. In vivo, we examined the cellular localization of the selected EGFP-tagged domains in transiently transfected HeLa cells. In vitro, we analyzed the role of the domains in lipid recognition and oligomerization using lipid-blot overlays and sedimentation assays. As previously shown for endogenous p125A (Shimoi et al., 2005), EGFP-tagged p125A localized with COPII at ERES (marked by the outer layer Sec31 subunit) that normally distributed in the cell periphery or clustered in the perinuclear region (Fig. 1E). At these latter sites, ERES were adjacent to, but did not localize with, the ER to Golgi intermediate compartment (ERGIC53, Fig. 1E), the cis-Golgi (gpp130, Fig. 1E) or the trans-Golgi network (TGN46, not shown) compartments. At higher expression levels, EGFP-p125A led to the enlargement of COPII-coated ERES membrane structures (Mizoguchi et al., 2000) (Fig. 6 and supplementary material Fig. S3). By contrast, the isolated EGFP-tagged DDHD domain did not colocalize with Sec31 but, strikingly, was distributed in a cytosolic pool and also showed robust association with PI4P-rich Golgi membranes (Fig. 2A). Membrane bound EGFP–DDHD decorated the rims of ERGIC (ERGIC53), cis-Golgi (gpp130 not shown) and TGN (TGN46, Fig. 2A) compartments, behaving like typical PI4P reporters such as the PH domain of Fapp1 (Weixel et al., 2005). We prepared a His6-tagged fragment of the C terminus encompassing the DDHD domain (residues 701–989) for analysis. Shorter fragments were difficult to produce because of low yields, indicating poor folding. In contrast to its targeting to PI4P-rich membranes in cells, when measured using lipid blot overlays the DDHD domain displayed only weak binding (observed with prolonged exposures) with somewhat broad specificities toward acidic lipids (Fig. 2B). Given this ineffective lipid recognition, we produced a larger fragment that included both SAM and DDHD domains (643–989) for analysis. Importantly, the combined domain exerted highly defined specificity to PI3P, PI5P and, in agreement with our cellular observations, PI4P (Fig. 2A,C). The domain also recognized PA and phosphatidylserine (PS), and weakly recognized phosphatidylinositol (3,4) bisphosphate [PI(3,4)P2], yet for the most part did not recognize more acidic polyphosphate PIs. The results suggest that inclusion of the SAM domain enhanced selective phospholipid recognition. It is possible that the SAM domain binds selective lipids. Alternatively, it may assemble DDHD domains to optimize lipid-binding avidity. To evaluate these possibilities, we generated a GST-tagged SAM domain. Analysis using lipid-blot overlay showed no lipid binding (not shown). Furthermore, the transiently expressed EGFP-tagged SAM domain remained cytosolic (Fig. 2F). Structural studies have demonstrated that the homologous SAM domain of diacyl glycerol kinase (DAGK) δ, a protein that regulates COPII assembly at ERES (Nagaya et al., 2002), dimerizes and generates oligomeric sheet structures that fall out of solution upon Zn2+ binding. Zn2+-induced sheet formation is dependent on the ability of the domain to dimerize, thus providing us with an easy test to monitor SAM oligomerization (Knight et al., 2010). As observed with the SAM domain of DAGKδ, addition of Zn2+ to GST-p125A-SAM domain led to robust polymerization of the protein, which quantitatively fell out of solution under these conditions (Fig. 2H). Addition of Ca2+ or Mn2+ had minimal effects on SAM solubility (not shown). The solubility of the control GST protein was also variably affected by the addition of Zn2+. We thus cleaved the SAM domain from the GST. The non-tagged SAM domain effectively precipitated in the presence of Zn2+, whereas a dimer mutant remained soluble (Fig. 2I, see below). The lipid blot overlay analysis suggested that DDHD-mediated lipid recognition is assisted by the p125A-SAM domain, which may optimize avidity-based membrane binding. EGFP–SAM–DDHD also localized to PI4P-enriched Golgi yet occasionally caused Golgi disassembly (not shown), as observed with other PI4P-binding domains (Weixel et al., 2005). A GFP-tagged fragment encompassing the linker region between the SAM and DDHD domains (701–778) remained cytosolic (not shown). Collectively, these results suggest a model in which cooperative activities of assembled SAM and DDHD domains promote selective lipid recognition and cellular membrane binding.

Fig. 2.
Fig. 2.

Cooperative lipid recognition by the SAM–DDHD module of p125A. (A) The DDHD domain targets PI4P-rich Golgi membrane in isolation. Transiently expressed EGFP–DDHD domain (779–989) dissociated from Sec31-containing ERES (arrowhead shows lack of colocalization between Sec31-stained ERES and localized EGFP–DDHD). The DDHD domain targets to the periphery of PI4P-enriched membranes and can be seen coating Golgi [right image, TGN46) (antibody dilution 1:500)] and ERGIC (ERGIC 53, middle image). Scale bars: 10 µm. (B-F) Selective lipid recognition is dependent on a module consisting of the DDHD and SAM domains. (B) 1 µg/ml of purified His-tagged p125A fragment (701–989) containing the DDHD domain, but not the SAM domain, was probed on lipid blot overlay using HRP-conjugated antibody against His6 (at 1:500 dilution). The fragment bound weakly to acidic lipids. (C) As in B: extending the fragment to contain the upstream SAM domain (643–989) conferred lipid selectivity to monophosphorylated PIs, PA, PS and PI(3,4)P2. (D) As in C: replacing a basic stretch of residues in the DDHD domain of the SAM–DDHD module with glutamic acid residues (850-KGRKR-854→EGEEE, termed PI-X) abolished lipid recognition. (E) The EGFP–DDHD domain is targeted to Golgi membranes (left), whereas EGFP–DDHDPI-X lost Golgi targeting (right). Scale bar: 5µm. (F) The EGFP-SAM domain transiently expressed in HeLa cells shows diffuse cytosolic distribution. Scale bar: 5µm. (G) The predicted structure of p125A SAM domain (643–704) (generated in Phyre) with red arrowhead indicating the position of a conserved leucine (690) within a hydrophobic dimer interface. (H) GST-SAM oligomerization is promoted by addition of Zn2+ and is sensitive to the L690E mutation. Buffer with or without the addition of 20 µM Zn(AOc)2 was added to GST-SAM or GST-SAML690E (10 µM of each), as indicated. Oligomerization was followed by the precipitation of the proteins from supernatant (S) to pellet (P) fractions (as indicated), using centrifugation and analysis on Coomassie-stained gels. (I) SAM and SAML690E domains (0.455 mM each) were incubated with 0.455 mM of Zn(AOc)2 and analyzed as in H.

Charge and hydrophobic interactions are used by the SAM and DDHD domains to support lipid recognition and assembly

Our in vivo and in vitro analyses (Fig. 2) suggest that lipid recognition resides within the DDHD domain and is assisted by SAM domain-mediated assembly. To test this hypothesis, we generated mutations in these domains and examined their functionality. Within the DDHD domain we focused on a group of basic residues (851-KGRKR-855) and replaced them with glutamic acid (851-EGEEE-855, termed PI-X). When tested in lipid blot overlay assays, the His6-tagged SAM–DDHDPI-X domain showed no lipid recognition (Fig. 2D), and transiently expressed EGFP-tagged DDHDPI-X lost its Golgi localization and presented a diffuse cytoplasmic distribution (Fig. 2E). This localization was also observed when EGFP–SAM–DDHDPI-X was analyzed (not shown). The results suggest that the DDHD domain is required for lipid recognition.

We further hypothesized that the SAM domain, which does not display lipid recognition or cellular targeting in isolation (Fig. 2F and not shown), supports protein assembly. To test this hypothesis, we used structural information available for the DAGK δ-SAM domain to guide mutagenesis aimed at abolishing protein assembly (Knight et al., 2010; Qiao and Bowie, 2005). In the DAGK δ-SAM domain, hydrophobic interactions between valine and leucine residues mediate dimerization required for oligomerization of the domain (Fig. 2G), whereas introduction of a charged residue at this position prevented both dimerization and Zn2+-induced high-order oligomerization. The hydrophobic dimer interface is fully conserved in p125A, thus we could generate a single residue replacement in the p125A SAM-domain (L690E) for analysis. Unlike the wild-type protein, GST-SAM(L690E) did not precipitate in response to Zn2+ (Fig. 2H,I). Thus, in common with DAGK δ, this single point mutation abolished dimerization and therefore high order assembly of the domain. Similarly, while the untagged SAM domain robustly precipitated with Zn2+ addition, SAM(L690E) remained completely soluble. The results suggest that the hydrophobic assembly interface within the SAM domain of p125A is functional.

Traffic inhibition by low temperatures reveals an exclusive localization of p125A at ERES

p125A may recognize monophosphorylated PIs, including PI4P at ERES, or on ERGIC or Golgi membranes adjacent to COPII bud sites. To define the site of p125A-membrane binding, we analyzed the localization of p125A in relation to COPII, ERGIC and Golgi markers in cells incubated at reduced temperatures that preferentially slow anterograde and retrograde traffic between the ER and the Golgi. When cells were incubated at 15°C under conditions that arrest biosynthetic anterograde and retrograde cargo traffic at ERGIC, ERGIC53 strongly accumulated in perinuclear ERGIC compartments and, as previously reported, ERGIC53 also scattered in defined puncta throughout the cytoplasm (Saraste and Svensson, 1991) (supplementary material Fig. S2A,C). This distribution was rapidly reversed when returned to 37°C (not shown). Golgi morphology remained largely unperturbed under these conditions (supplementary material Fig. S2B). Importantly, at 15°C the number of ERES (marked by Sec31) was reduced, while individual sites were enlarged and cytosolic COPII was effectively concentrated at these sites (Fig. 3A). Incubation of cells at 10°C leads to the selective accumulation of folded biosynthetic cargo in COPII-coated ERES membranes that maintain continuity with the bulk ER membrane and are functional in ER export (Mezzacasa and Helenius, 2002). Under these conditions, ERES further coalesced, collecting COPII layers in enlarged puncta (Fig. 3A–C). Importantly, p125A exclusively partitioned with COPII at both 15°C and 10°C [Fig. 3A, and Fig. 7A,B (endogenous p125A)]. p125A-COPII sites segregated from perinuclear sites where both ERGIC and Golgi compartments largely clustered (supplementary material Fig. S2A–B). To determine whether enlarged COPII-p125A sites coat ERES membranes, we analyzed the membrane protein ERGIC53, which cycles between ER and ERGIC. Similar to endogenous protein, overexpressed YFP-p58 (rat ERGIC53) accumulated in the perinuclear region at 15°C or 10°C, and also in distinct scattered puncta that were particularly evident at 15°C. YFP-p58 also backed up in the ER, most likely due to overexpression (supplementary material Fig. S2C). As a result, YFP-p58 was also arrested at enlarged ERES at 15°C and 10°C to demonstrate that ERES membranes are coated with p125A and COPII under these conditions [supplementary material Fig. S2C (Mezzacasa and Helenius, 2002)]. Assembly of ERES at 10°C required functional p125A (see below, Fig. 7A,B and not shown). Overall, the analysis suggests that p125A exclusively co-segregates with COPII and coats ERES membranes. Lipid recognition mediated by the cooperative activity of SAM and DDHD domains supports ERES localization.

Fig. 3.
Fig. 3.

p125A associated with COPII-coated ERES during temperature-induced traffic inhibition. (A) HeLa cells transiently expressing mRFP-p125A (8–12 hours) were maintained at 37°C, or incubated at 15°C or 10°C, as indicated, for 4 hours, then fixed and analyzed for localization with ERES (hSec31A). (B) Enlarged images of boxed areas in A, arrowheads indicate the extensive colocalization of p125A to ERES. (C) Colocalization of endogenous hSec31a and hSec24c (antibody at 1:100 dilution) in cells incubated at 10°C, as in A. Scale bars: 5µm.

COPII-p125A-containing ERES segregate from Sec16A during ER exit

Sec16p substitutes for the requirement of acidic lipids during COPII assembly on synthetic membranes, whereas Sec16A and PI4KIIIα are both required during adjustment of ERES assembly with cargo load (Farhan et al., 2008; Supek et al., 2002). We hypothesized that lipid recognition by p125A may support COPII assembly at stages that precede or follow Sec16 regulation. To explore this hypothesis, we localized Sec31, Sec16A and p125A using the temperature blocks described above. We analyzed the localization of both endogenous Sec16A (KIAA0310, using specific antibody), as well as a EGFP-tagged Sec16A with relation to the localization of Sec31 and p125A with similar results (Fig. 4 and Fig. 7A). At 37°C, endogenous Sec16A and transiently expressed EGFP-Sec16A presented ERES localization and diffused cytosolic distribution. Importantly, slowing traffic at the ER (10°C) or at ERGIC (15°C), led to robust collection of Sec16A at perinuclear sites adjacent and closely associated with ERGIC53/p58-containing membranes (Fig. 4 and inset in Fig. 4C). The results support a dynamic distribution of Sec16A between cytosol and membranes, which is reduced at low temperatures. Surprisingly, Sec31, Sec23, mRFP-p125A and endogenous p125A exhibited different behavior to the perinuclear clustering of Sec16A under these conditions, assembling at enlarged scattered ERES and segregating from Sec16A (Figs 3, 4 and not shown). The enhanced segregation of Sec16 from COPII-p125A-coated ERES under conditions that either delay (10°C) or enable (15°C) slow cargo exit suggest that p125A may regulate a late stage that follows the initial nucleation of COPII by Sec16.

Fig. 4.
Fig. 4.

ERES coated p125A segregates from Sec16A at low temperatures. (A) HeLa cells transiently expressing EGFP-Sec16A (8–12 hours) were maintained at 37°C, or incubated at 15°C or 10°C for 4 hours, and the localization of ERES (marked by hSec31a) and EGFP-Sec16A was determined. (B,C) Control HeLa cells (B) or cells transiently expressing mRFP-p125A (C) were incubated at 10°C and the localization of endogenous Sec16A (antibody at 1:1000 dilution), hSec31a and mRFP-p125A was determined, as indicated. Inset in C shows HeLa cells transiently expressing YFP-p58 analyzed for the localization of endogenous Sec16A and YFP-p58 during 10°C incubations. Arrowheads indicate hSec31 (A,B) or mRFP-p125A (C) sites, segregated from Sec16A. Scale bar: 5 µm.

Assembly controlled lipid recognition is required to regulate COPII organization at ERES

We hypothesized that p125A uses lipid recognition to direct COPII assembly at ERES. Tagged p125A in which specific residues required for SAM-domain assembly and DDHD-supported lipid recognition (Fig. 2) were mutated (p125API-X, p125AL690E, and SAM and DDHD double mutant p125API-X, L690E) were generated to test this hypothesis. Individual mutations in the SAM (L690E) or DDHD domain (PI-X) may not be sufficient to generate a dominant phenotype because PI-X mutants may assemble with the endogenous protein and L690E mutants may retain lipid recognition (Fig. 2B-I). Importantly, the interactions of mutated and wild-type p125A with both COPII layers are maintained by the unperturbed N terminus (Ong et al., 2010). All mutants were expressed at similar levels when analyzed by western blots (Fig. 7E and not shown). EGFP-p125A localized to ERES (Fig. 5). EGFP-p125API-X was also targeted to ERES. However, a diffused cytosolic component was clearly evident. For the most part, overexpression of EGFP-p125API-X did not lead to clustering, as observed with the wild-type protein (not shown and supplementary material Fig. S3A). Because this mutant retains the ability to interact with endogenous p125A using the dimerization interface, we analyzed both a SAM oligomerization mutant p125AL690E (Fig. 2H,I) and a mutant (p125API-X, L690E), in which both the dimerization and lipid recognition abilities were disabled. EGFP-p125AL690E exhibited diffuse labeling and association with ERES, similar to the PI-X mutant. Overexpression of EGFP-p125AL690E led to augmented diffuse cytosolic distribution with occasional targeting to both ERES and the perinuclear Golgi region (Fig. 5). Importantly, in marked contrast to wild type, p125API-X, L690E, in which the presumed cooperative lipid-binding module is disabled, showed no ERES localization and exhibited diffuse cytosolic distribution, which was maintained even at very high expression levels (Fig. 5 and supplementary material Fig. S3A). These results suggest that selective lipid recognition mediated by the SAM–DDHD module is required for p125A-membrane binding at ERES. We thus tested whether selective membrane binding by p125A regulates COPII assembly at ERES. Indeed, p125API-X, L690E became a trans-dominant negative inhibitor of ERES assembly. Sec31 lost ERES localization in p125API-X, L690E-expressing cells and remained diffusely localized in the cytoplasm (Fig. 5). Moreover, p125API-X, L690E also inhibited the assembly of ERES when measured at 10°C, although it did not affect Sec16A localization (not shown). These results suggest that cooperative lipid recognition derived from SAM–DDHD activities is required to regulate COPII assembly at ERES. The DDHD domain recognized PI4P-rich membranes in cells and on lipid blot overlays. Similar to p125API-X, L690E, deletion of the DDHD domain (Δ778–989) in the background of SAM domain inactivation (using the L690E mutation, p125AL690E, ΔDDHD) led to cytosolic dispersion of the protein and inhibited ERES assembly, as analyzed by Sec31 staining (Fig. 6A). We used this truncation to further test the role of p125A SAM–DDHD as a selective lipid recognition module directing ERES assembly, by artificially replacing the domain with a bona fide PI4P-binding domain, the pleckstrin-homology (PH) domain of Fapp1 (Blumental-Perry et al., 2006). Fapp1-PH confers selective recognition of PI4P and, as also observed with the isolated DDHD domain, is targeted in isolation to PI4P-rich Golgi membranes (Weixel et al., 2005). When expressed in cells, the chimera (p125AL690E, ΔDDHD, +Fapp1-PH) displayed both cytosolic and punctate distribution, with some preferential targeting to the Golgi (Fig. 6A and Fig. 5). Importantly, in contrast to expression of p125API-X, L690E or p125AL690E, ΔDDHD, which disrupted ERES assembly, the chimera colocalized with and maintained assembly of endogenous Sec31 at both perinuclear Golgi adjacent sites and peripheral ERES (Fig. 6A and Fig. 5). The ability to replace the SAM–DDHD lipid recognition module artificially with a PI4P-binding domain and partially restore ERES assembly supports the role of PI4P in p125A-mediated assembly of ERES.

Fig. 5.
Fig. 5.

PI4P binding by the SAM–DDHD module of p125A controls ERES assembly. The localization of transiently expressed (12–14 hours) full-length EGFP-p125A wild type, PI-X, L690E or the double mutant (p125AL690E,PI-X) and ERES (hSec31a) were analyzed in HeLa cells as indicated. The bottom panel shows the expression of a EGFP-p125A chimera where the DDHD domain has been substituted with the PH domain from Fapp1 in the backbone of an L690E mutant. EGFP-p125API-X or EGFP-p125AL690E mutants become partly cytosolic, whereas the double mutant p125API-X, L690E lost membrane localization and had completely disrupted ERES assembly. Membrane targeting and ERES assembly was partially restored in the Fapp1-PH-containing p125A chimera-expressing cells. Arrowheads indicate a non-transfected cell surrounded with cells expressing p125API-X, L690E (*). Scale bar: 10 µm.

Lipid recognition controls p125A residency at ERES

How could lipid recognition by p125A control ERES assembly? One possibility is that lipid recognition is required to target p125A and associated Sec31 proteins to ERES. However, p125A is targeted to ERES through association of its N terminus with Sec23 and Sec31. Alternatively, lipid recognition may regulate the residency time of p125A and its associated coat proteins on ERES membranes. To test this possibility, we analyzed the dynamics of p125A proteins and COPII at ERES using fluorescence recovery after photobleaching (FRAP). HeLa cells were transiently transfected with constructs expressing YFP-Sec23, EGFP-p125A (supplementary material Fig. S1) or mRFP-p125A. An average of 27–35 measured events for each time-point collected in three independent experiments is shown. COPII marked by both YFP-Sec23 (supplementary material Fig. S1B) or CFP-Sec31 (not shown) exhibited typical recovery kinetics (Forster et al., 2006), whereas EGFP-p125A (supplementary material Fig. S1C) or mRFP-p125A (not shown) both exhibited similar or slightly slower dynamics. Importantly, faster recovery kinetics were observed for EGFP-p125AL690E (supplementary material Fig. S1D) or EGFP-p125API-X (supplementary material Fig. S1E), providing an explanation for the observed diffused cytosolic population of these mutants (Fig. 5). The initial 20 seconds of the recovery phases were fitted into single exponentials (Forster et al., 2006) with overall averaged T½ for YFP-Sec23 (3.14 seconds) or EGFP-p125A (3.36 seconds). EGFP-p125AL690E (T½ = 2.98 seconds.) and EGFP-p125API-X (T½ = 2.56 seconds) showed faster kinetics. Because EGFP- p125API-X, L690E did not assemble at defined sites, it could not be measured. However, limited association with ERES was observed when EGFP- p125API-X, L690E was analyzed at reduced temperatures (not shown), suggesting that enhanced turnover at ERES prevented stable localization. The results suggest that selective lipid recognition by the SAM–DDHD module controls p125A residency at ERES membranes to regulate COPII assembly.

p125A functions at a late stage in ERES nucleation

p125A-Sec23-Sec31-coated ERES segregated from Sec16A during temperature-induced traffic blocks at ERES or ERGIC (Figs 3,4). We hypothesized that p125A actively displaces Sec16A from ERES. We therefore prolonged the expression time of p125A from 12 to 16 hours, used in experiments presented so far, to 24 hours to examine if such overexpression would lead to Sec16 displacement from ERES. Overexpression of p125A led to a marked uniform enlargement of ERES, as previously demonstrated (mRFP-p125A, Fig. 6B and supplementary material Fig. S3A and EGFP-p125A, not shown) (Mizoguchi et al., 2000; Shimoi et al., 2005; Tani et al., 1999). The large sites were marked by ER membranes containing the cargo protein Venus-tsVSV-G (vesicular stomatitis virus glycoprotein). Bud structures containing tsVSV-G were evident on these structures upon a temperature shift that leads to exit of tsVSV-G from the ER (super resolution SIM images, supplementary material Fig. S3B and Movies S3-S5). Analysis of tsVSV-G arrival at the cis/medial Golgi (using Endo-H digestion), under these extreme p125A expression conditions, showed that although traffic was somewhat inhibited it remained functional (supplementary material Fig. S3B). Indeed, the enlarged ERES collected both layers of COPII, as analyzed by the colocalization of endogenous Sec31 or co-expressed YFP-Sec23 (Fig. 6B and not shown). We analyzed whether EGFP-Sec16A is displaced from these sites. When expressed at low levels, mRFP-125A largely colocalized with Sec16A, Sec31 and Sec23 at ERES (Figs 3,4). By contrast, p125A-induced large ERES were clearly lacking EGFP-Sec16A (compare YFP-Sec23-mRFP-p125A with EGFP-Sec16A-mRFP-p125A; Fig. 6B, middle versus top row). Occasionally, Sec16A was found adjacent to large rounded sites that varied from 500 to 1500 nm in size, as shown by SIM (supplementary material Fig. S3C). Similar localization was observed when endogenous Sec16A was analyzed (not shown).

Fig. 6.
Fig. 6.

Lipid recognition by p125A regulates Sec16A displacement from ERES. (A) HeLa cells transiently expressing (12–14 hours) mRFP-p125AL690E, ΔDDHD or mRFP-p125AL690E, ΔDDHD + Fapp1-PH were analyzed for hSec31 localization, as indicated. ERES [hSec31a (antibody dilution 1:200)] are disassembled in cells expressing mRFP p125AL690E, ΔDDHD (asterisks mark transfected cells), and mRFP-p125AL690E, ΔDDHD + Fapp1-PH and hSec31a are colocalized (arrows). (B) HeLa cells transiently expressing mRFP-p125A, mRFP-p125AL690E, ΔDDHD, EGFP-Sec16A, YFP-Sec23A, as indicated, for 24 hours were fixed and analyzed for the localization of transfected proteins. Note the segregation of EGFP-Sec16A from mRFP-p125A as opposed to the co-assembly of mRFP-p125A with YFP-Sec23a (arrow), and the collection of EGFP-Sec16A in mRFP-p125AL690E, ΔDDHD sites (arrows). Scale bar: 5 µm.

As Sec16p negates the requirements for acidic lipids in COPII binding to liposomes (Supek et al., 2002), we hypothesized that p125A might use PI4P binding to direct Sec16A displacement, thus stabilizing COPII-membrane binding. We therefore analyzed whether overexpressed p125A proteins in which the lipid-binding module was inactivated (mRFP- p125API-X, L690E) or deleted (p125AL690E, ΔDDHD) are also defective in Sec16 displacement. In marked contrast to mRFP-p125A, mRFP- p125API-X, L690E remained dispersed even at very high expression levels (supplementary material Fig. S3A). Overexpressed p125AL690E, ΔDDHD also remained largely cytosolic, but enlarged sites were now occasionally evident (Fig. 6B; supplementary material Fig. S3A). Importantly, these sites effectively collected EGFP-Sec16A (Fig. 6B). Analysis using SIM demonstrated that p125AL690E, ΔDDHD-induced sites were engulfed within EGFP-Sec16A (supplementary material Fig. S3C; Movies S1,S2). Wild type and p125AL690E, ΔDDHD interact similarly with both COPII layers. Therefore, the presence of a lipid-binding module in p125A controls the displacement of Sec16A from COPII-ERES.

The results suggest that PI4P (or lack of) can regulate p125A to control Sec16A localization. Depletion of PI4KIIIα is known to inhibit ERES assembly. We asked how such depletion would affect Sec16A localization. siRNA-mediated PI4KIIIα depletion led to redistribution of hSec31a, which now clustered at perinuclear sites, whereas peripheral ERES numbers were reduced, as previously demonstrated (Farhan et al., 2008). Importantly, PI4KIIIα depletion led to clustering of Sec16A at perinuclear sites, where colocalization with hSec31a increased (supplementary material Fig. S4). These results support a model (Fig. 8) in which PI4P signals regulate the displacement of Sec16A from ERES during COPII assembly.

Functional contributions of the SAM–DDHD membrane-binding module

Depletion of p125A using RNAi (Fig. 7E) leads to disruption of ERES (Shimoi et al., 2005) as further evidenced by the pronounced loss of ERES in p125A-depleted cells incubated at 10°C, under conditions where Sec16A is segregated (Fig. 7A and inset). ERES assembly under these conditions was restored by the expression of an RNAi-resistant form of mRFP-p125A (Fig. 7B). p125A depletion causes kinetic inhibition of membrane and soluble cargo secretion from the ER, leading to the disruption of Golgi morphology (Ong et al., 2010). To examine the functional contribution of the SAM–DDHD lipid-binding module to p125A activity in ER exit, we replaced endogenous p125A with EGFP-p125API-X, L690E. We analyzed Golgi morphology as a reporter for overall steady-state traffic activities. Golgi morphology was analyzed using GalNAcT2-GFP localization (Fig. 7C) and quantified using gpp130 morphology (Fig. 7D). Morphology was heterogeneous with four distinct phenotypes (Fig. 7D): (1) intact Golgi localized to the perinuclear region; (2) loosely packed or dispersed Golgi located in the perinuclear region or around the nuclei; (3) completely shattered (vesiculated) Golgi dispersed throughout the cell; and (4) cells missing a detectable Golgi compartment, perhaps suggestive of a mitotic cell population. Effective depletion of endogenous p125A was achieved as previously reported (Ong et al., 2010) (Fig. 7E). Depletion of p125A led to dramatic reduction in the intact Golgi population and a concomitant increase in shattered morphology when compared with control RNAi-treated cells (Fig. 7F,G). The effect was specific as expression of an RNAi resistant form of EGFP-p125A (Fig. 7E-G) reversed these effects, namely eliminating the shattered Golgi morphology and restoring intact Golgi populations (Fig. 7F,G). By contrast, expression of EGFP-p125API-X, L690E was ineffective, leading to only a partial restoration of intact Golgi morphology and to a reduction in shattered Golgi compartments (Fig. 7F,G). Overall, these results suggest that defects in the activity of the SAM–DDHD lipid-binding module of p125A, which led to robust morphological defects in ERES assembly (Fig. 5), to reduced association of p125A with ERES (supplementary material Fig. S1) and to inhibition of Sec16 segregation from ERES (Figs 4,6 and supplementary material Fig. S3), also translated into functional defects in steady-state traffic activities required to maintain Golgi morphology.

Fig. 7.
Fig. 7.

Functional SAM–DDHD module is required for steady-state ER-to-Golgi traffic. (A) Merge images showing the localization of endogenous p125A (green, antibody dilution 1:1000) and hSec31a (red) at ERES arrested at 10°C in control and p125A-depleted cells, as indicated (dsRNAi, supplementary material Table S2). Inset shows the localization of endogenous Sec16A (green) and hSec31a (red) under similar conditions. (B) Control or p125A-depleted cells were transfected with RNAi-resistant mRFP-p125Ar and traffic was arrested by incubation at 10°C. The localization of hSec31a was analyzed [merge images are shown, arrowhead indicates depleted cell next to rescued cells (asterisk)]. (C) HeLa cells stably expressing GFP-tagged N-acetylgalactosaminyltransferase-2 (GalNAcT2-GFP) were treated with control or p125A directed dsRNAi (supplementary material Table S2), as indicated, and visualized for GFP. Note the dramatic change in Golgi morphology (arrowheads) with loss of intact Golgi populations and the concomitant increase in shattered Golgi morphology. Scale bar: 5 µm. (D) Golgi morphology was quantified in HeLa cells by analyzing gpp130 localization. Typical observed Golgi morphologies are shown and color-coded, including intact (blue), dispersed (pink) and shattered (vesiculated, green). In some cells, Golgi morphology was not recognizable (labeled as missing, purple). (E) Analysis of p125A knockdown efficiency by western blots. Endogenous expression of p125A (middle panel, 1:2500) and actin (lower panel, 1:10,000) are shown. The upper panel shows the comparable expression of EGFP-p125A-resistant clones (EGFP-p125Ar; GFP antibody dilution 1:10,000) in control and KD cells. EGFP ran below the analyzed area and is not shown. The expression of EGFP-p125Ar (wild type) and EGFP-p125AL690E, PI-X were also detected by the p125A-specific antibody but required prolonged exposures due to partial transfection efficiency of KD cells. (F) The fractional distribution of Golgi morphologies in control, p125A-depleted and rescued cell populations with EGFP-p125Ar and EGFP-p125API-X, L690Er, as indicated. The percentage of cells with either intact Golgi (blue), dispersed Golgi (pink), shattered Golgi (green) or missing Golgi (purple) under each treatment condition is shown. (G) Statistical analysis of intact Golgi morphology in control and KD cells (mean±s.d.). Three experiments were performed for each condition. Ten images were collected from each experiment and Golgi phenotypes were determined for all cells expressing EGFP-tagged proteins (93–220 cells per group). Unpaired Student's t-test was used to test for significant differences between groups.

DISCUSSION

Although COPII core subunits are sufficient in mediating vesicle biogenesis, COPII-interacting proteins control budding activities at ERES. We identified a cascade in which p125A, a protein that links both COPII layers, uses a lipid recognition module (Figs 1, 2) to stabilize membrane binding (supplementary material Fig. S1) and promotes the spatial segregation of COPII from Sec16A (Figs 3,4,6 and supplementary material Fig. S2–S4), leading to functional ERES assembly (Figs 5,7). The results support a model in which regulatory activities such as lipid signaling (Blumental-Perry et al., 2006; Farhan et al., 2008; Nagaya et al., 2002; Pathre et al., 2003) control the progression of a molecular cascade that directs COPII nucleation and activity at ERES (Fig. 8).

Fig. 8.
Fig. 8.

The COPII budding cascade at ERES. (A–C) Following initial association of COPII layers with Sec16 on ER membranes (A), the binding of PI4P by p125A promotes the displacement of Sec16 from COPII inner and outer layers to allow for effective linking between coat layers. This drives coat retention (B) while enhancing GTP hydrolysis to support later steps, including Sar1 exchange by Bet3 on Sec23, and Sar1-induced vesicle neck constriction and fission (C).

The SAM–DDHD lipid-binding module

Previous studies suggested that the C terminus of p125A, which contains SAM and DDHD domains, is involved in membrane binding. A defined function of SAM domains is protein oligomerization (Qiao and Bowie, 2005), which is used to increase avidity between assembled complexes and substrates. Our findings suggest the basic assembly activity of SAM is functional in p125A. First, we showed that, in common with its close homolog, the SAM domain of DAGKδ (Knight et al., 2010), the p125A-SAM domain oligomerized when bound to Zn2+. Second, we demonstrated that the basic assembly interface is conserved and introduction of a single point mutation within the site abolished oligomerization (Fig. 2). Third, we demonstrated that the SAM assembly interface within p125A is required to support ERES organization and activity (Figs 5-⇑7; supplementary material Figs S1 and S3).

Because p125A-SAM lacked membrane targeting in vivo or lipid-binding activities in vitro, we favor a model in which this domain enhances the avidity for lipid recognition by the DDHD domain, as suggested by lipid blot overlay analysis (Fig. 2). This activity may be shared in other lipid processing enzymes that function in vesicular transport. In DAGKδ, a module containing SAM and PH domains is required for inhibition of ERES assembly although lipid-binding specificities of this PH domain are undefined (Nagaya et al., 2002). The SAM domain of the inositol 5-phosphatase Ship2, which regulates clathrin-mediated endocytosis, may be similarly required for membrane recognition (Nakatsu et al., 2010). The Ship2-SAM domain further supports heterodimerization with Arap3 (Raaijmakers et al., 2007). By analogy, heterodimeric interactions between SAM-DAGKδ and SAM-p125A may function to form lipid-recognition and processing hubs at ERES.

Lipid recognition most likely resides in the p125A-DDHD domain. When expressed in cells in isolation, EGFP–DDHD was targeted to PI4P-rich Golgi membranes (Fig. 2). Mutations in the domain (DDHDPI-X), which prevented Golgi binding in cells (and further abolished the targeting of EGFP–SAM–DDHDPI-X domain, not shown), also abolished lipid recognition by the SAM–DDHD module in vitro (Fig. 2). These mutations probably did not destabilize DDHDPI-X because the protein did not aggregate in cells and was produced in bacteria at yields higher than its wild-type version. Moreover, deletion of the DDHD domain (p125AL690E, ΔDDHD) abolished membrane binding and dispersed ERES similarly to proteins carrying the DDHDPI-X mutations (p125API-X, L690E, Figs 5,6 and supplementary material Fig. S3). Future structural studies are needed to define the contributions of the basic PI-X cluster in lipid recognition. Together, the SAM–DDHD module provides a lipid recognition unit for p125A that regulates COPII assembly (Fig. 5).

In vitro analysis suggests that the SAM–DDHD module recognizes monophosphorylated PIs, PA and PS (Fig. 2). However, several lines of evidence suggest that PI4P might be a primary target for this module: (1) full-length p125A shows specificity for PIP recognition (Inoue et al., 2012; Shimoi et al., 2005); (2) p125B, a family member of p125A that localizes to the Golgi, recognizes PI4P using its SAM and DDHD domains, although individual contributions of these domains were not defined (Inoue et al., 2012). p125B–SAM–DDHD can replace the p125A lipid-recognition module and the resulting chimera localizes at ERES (Shimoi et al., 2005); (3) EGFP–DDHD domain is targeted to PI4P-rich Golgi membranes (Fig. 2); (4) p125A mediated ERES targeting and organizing activity, which is lost when the SAM dimer interface is disabled (L690E mutant) and the lipid-binding DDHD domain is mutated (PI-X) or deleted (Figs 5,6 and supplementary material Fig. S3), was partially restored by simple replacement of the DDHD domain with a well-characterized PI4P-binding domain (Fapp1-PH, Fig. 6); (5) deletion of the DDHD domain in p125A or siRNA-mediated PI4KIIIα depletion, which reduces PI4P levels, both inhibit the displacement of Sec16A from ERES (Fig. 6 and supplementary material Fig. S3,S4); (6) DDHD domains are found in Golgi (PI4P-rich)-targeted proteins, including p125B (Nakajima et al., 2002; Yamashita et al., 2010) and Nir-2 (Litvak et al., 2005). Collectively, the results suggest that p125A can use PI4P to localize and regulate COPII activities at ERES.

Role of p125A in ERES regulation

A dependency on acidic lipids and in particular PI4P in yeast COPII assembly, which is abrogated by Sec16p, provided the first link between the two activities. However, such dependency is observed only on synthetic membranes (Supek et al., 2002). Depletion of the yeast major PI4 kinases PIK1 and Stt4 failed to affect ER-to-Golgi traffic (Audhya et al., 2000). Subsequent studies have shown that sequestration of Golgi PI4P in vitro or prolonged PIK1 inactivation in vivo do inhibit ER to Golgi traffic (Lorente-Rodríguez and Barlowe, 2011), yet inhibition is exerted on fusion of COPII vesicles with Golgi membranes. Unlike yeast, mammalian COPII vesicles do not fuse with the Golgi but rather fuse homotypically in the vicinity of ER bud sites and PI4P formation is initiated at these sites (Blumental-Perry et al., 2006; Farhan et al., 2008).

Sec16A is required to nucleate new ERES, and both Sec16A and the ER-localized PI4KIIIα are required to maintain these sites (Farhan et al., 2008). Sec16p-COPII interactions hinder functional linkage between COPII layers, thus inhibiting Sec31-stimulated GAP activity (Kung et al., 2012; Supek et al., 2002; Yorimitsu and Sato, 2012). However, functional linkage is required for the completion of vesicle budding (Fromme et al., 2007). Whereas Sec16p is dispensable for in vitro budding and only enhances vesicle release, it is a vital component in vivo. Our results provide a plausible working model to explain the seemingly contradictory inhibitory and supportive roles of Sec16. In this model, COPII interactions with Sec16 and with PI4P-p125A each represent sequential steps in a budding cascade (Fig. 8). Sec16 initiates COPII nucleation on ER membranes yet is physically segregated from the coat as it establishes proper linkage between its layers. Indeed, Sec16A is slightly removed from bud sites at steady state (Hughes et al., 2009). At low temperatures that inhibit ER exit, it is displaced from arrested p125A-controlled ERES. This enhanced displacement is maintained when ER exit is active, yet traffic is slowed subsequently at ERGIC (Fig. 4). Sec16A displacement is also enhanced by overexpression of p125A and conversely is inhibited by inactivation of the lipid-binding module of p125A (Fig. 6; supplementary material Fig. S3) or by PI4KIIIα depletion (supplementary material Fig. S4). p125A, which interacts with both of the COPII layers, may provide a mechanism to facilitate the progression of COPII budding, promoting Sec16 dissociation while linking both COPII layers. p125A is required to stabilize ERES following segregation from Sec16A (Fig. 7A,B) and uses its SAM–DDHD lipid binding module in these reactions (Fig. 6 and supplementary material Fig. S3), thus employing lipid signals to control the assembly cascade. It remains to be determined how p125A-membrane binding regulates the segregation of COPII from Sec16A. p125A is a late addition in the evolution of COPII, which is lacking in yeast; thus, a mechanism by which Sec16p inhibition of functional coupling between COPII layers is reversed has yet to be determined.

The physiological role of p125A is unknown. Acute depletion or overexpression of p125A leads to general traffic defects (Fig. 7 and supplementary material Fig. S3). p125A depletion interferes with neural crest cell migration in Xenopus and was predicted to be a causative gene in the development of Waardenburg syndrome (McGary et al., 2010). However, p125A-KO mice are relatively unaffected (Arimitsu et al., 2011), mainly presenting defects in spermiogenesis. Sec16 may provide sufficient support for COPII activities in these animals, similar to the documented functionality of COPII on liposomes that contain Sec16p yet lack acidic lipids (Supek et al., 2002). The identified steps in ERES assembly are probably subjected to physiological regulation, which can now be defined.

MATERIALS AND METHODS

HeLa cells were maintained at sub-confluence in Dulbecco's modified Eagle's media (DMEM) (HyClone Fisher-Scientific) supplemented with up to 10% fetal bovine serum (FBS) (Serum Source International) and 5% penicillin-streptomycin (Cellgro) under a standard incubation environment (37°C, 5% CO2). Antibodies against p125A (MSTP053, Bethyl Lab; AP114511b, Abgent), GFP (24240, Polysciences), ERGIC53 (G1/93, ALX-804-602, Enzo Life Sciences), anti-Sec31A (612350, BD Transduction Laboratories), Flag (M2, F1804, Sigma-Aldrich), β-Actin (ab6276, Abcam), GFP (332600, Invitrogen), PI4KIIIα (Cell Signaling), HRP-conjugated anti-GST antibodies (ab3416-250, Abcam) and HRP-conjugated anti-His (040905270001, Roche) were used. Mouse monoclonal anti p125A and rabbit anti KIAA03100 (Sec16L) provided by Katsuko Tani (Tokyo University of Pharmacy and Life Science, Hachioji, Tokyo, Japan) were used. All Golgi-specific antibodies were provided by Adam Linstedt (Carnegie Mellon University, Pittsburgh, PA). Alexa-conjugated secondary antibodies were from Invitrogen. HRP- conjugated secondary antibodies were from Pierce.

EGFP-Sec16A was kindly provided by Vivek Malhotra (GRC, Barcelona, Spain). YFP-p58 was kindly provided by Jennifer Lippincot-Schwartz (NIH). p125A was excised from pFlag-CMW-6c-p125A, kindly provided by Katsuko Tani, (Tokyo University of Pharmacy and Life Science, Tokyo, Japan) using unique HindIII and SmaI restriction sites and ligated into a modified pEGFP-C1 where a HindIII site was added in-frame. The construction of EGFP- and mRFP-tagged constructs expressing p125A, p125A mutants, selected p125A domains and p125A-Fapp1-PH chimera is detailed in supplementary material Table S1.

Transfection was carried out using Effectene (Qiagen) or Lipofectamine 2000 (Life Sciences) reagents, according to manufacturers' protocols, with optimized DNA concentrations. NRK-derived ER microsomes and rat liver cytosol were prepared as described previously (Rowe et al., 1996). His6-tagged Sar1 H79G and T39N proteins were purified as described previously (Aridor et al., 1995; Rowe and Balch, 1995). His6-DDHD and SAM–DDHD expression in BL 21DE3 (Invitrogen) was induced with IPTG (0.1 mM) for 4 hours at 37°C and cells were lysed as described previously (Aridor et al., 1995). 10% N-Lauroyl Sarcosine (MP Biomedicals) was added to lysates that were further homogenized by sonication. Cell debris were removed by centrifugation and supernatants were supplemented with 2% n-Octyl-β-D-glucopyranoside (OG) (Gold Biotechnology, USA). Solubilized proteins were loaded on Ni-NTA-Agarose (Qiagen). Protein-bound beads were washed three times with 50 mM Tris-HCl (pH 8.0), 100 mM NaCl, 1 mM EDTA, 1 mM PMSF and 2% OG, followed by three washes with HNE buffer [50 mM HEPES (pH = 7.4), 300 mM NaCl, 1 mM MgCl2, 0.5 mM EGTA, 2% OG] and three washes with HNE buffer supplemented with 25 mM imidazole (pH 7.4). Bound proteins were eluted with HNE buffer containing 500 mM imidazole (Frangioni and Neel, 1993). GST-tagged SAM and SAML690E proteins were purified on GS-Sepharose 4B (GE Healthcare Life Science) and thrombin cleaved using a standard bulk GST purification protocol.

p125A knockdown-replacement analysis

Knockdown of p125A is described in supplementary material Table S2. For analysis of Golgi morphology, 10 individual fields positive for EGFP expression were collected from three independent experiments (30 images in total for each condition). All cells in the field were visually scored for Golgi morphology and p125A expression. Data were analyzed using Windows Excel 2010 (Microsoft Corporation). Homoscedastic two-tailed Student's t-tests on the percentage of intact Golgi were performed using Microsoft Excel 2010.

Temperature-block analysis

HeLa cells expressing FP-proteins for 14–16 hours were incubated in media supplemented with 20 mM HEPES (pH 7.4) (Fisher Scientific) and the cells were incubated at 15°C or 10°C for 4 hours. Samples were fixed and processed for analysis.

Lipid blot-overlay

PIP Strips (Echelon) were blocked for 1 hour in TBS-Tween buffer [Tris-buffered saline (TBS, pH 8.0), 1% Tween-20) supplemented with 3% BSA (BSA, Fraction V, EMD) and incubated for 2 hours. in the same buffer supplemented with 1 µg/ml of GST or His6 tagged proteins. Strips were washed (6×5 minute incubations) in TBS-Tween and incubated for 1 hour in TBS-Tween supplemented with 3% BSA and HRP-conjugated murine anti His6 or GST antibodies. Subsequently, strips were washed (6×5 min.) in TBS-Tween and visualized using SuperSignal West Dura Extended Duration Substrate (Thermo Scientific) and HyBlot CL X-Ray film (Denville Scientific), according to manufacturers' protocols.

Immunofluorescence

Indirect immunofluorescence was carried out as described previously (Aridor et al., 1995). Images were acquired on an Olympus Fluoview 1000 confocal system using an inverted microscope (IX-81 Olympus) and 60×NA 1.42 PLAPON objective. Images were processed using FV10-ASW V. 02.00.03.10 (Olympus Corporation) and Adobe Photoshop CS3 (Adobe Photoshop Version: 10.0.1).

Zn2+ based polymerization assay

GST-SAM or GST-SAML690E were diluted to a final concentration of 10 µM in 50 mM Tris-HCl (pH = 7.5), 100 mM NaCl. An equal volume of 20 µM Zn(OAc)2 in the same buffer was added and polymerization was estimated by centrifugation at 18,000 g using a cooled microfuge (Sorvall). Supernatant and pellet fractions were visualized on SDS-PAGE gels using Coomassie Blue staining. For thrombin-cleaved proteins, SAM, SAML690E and Zn(OAc)2 (both at 0.455 mM) were incubated in buffer containing 50 mM Tris-HCl (pH 7.5) and 100 mM NaCl, and polymerization was determined as above. Proteins were visualized using SilverQuest Staining Kit (Invitrogen).

Coat recruitment to LUVs or ER microsomes

LUV-COPII recruitment assays using rat liver cytosol (RLC) and Sar1 proteins were performed as described previously (Bielli et al., 2005). p125A or SNX9 were depleted by immunoprecipitation from RLC. The resulting supernatants were adjusted for protein concentration, verified for similar Sec23 and HSP70 levels, and used to measure COPII recruitment to microsomes as described previously (Pathre et al., 2003).

Acknowledgments

We thank Katsuko Tani and Mitsuo Tagaya (Tokyo University, Japan), William E. Balch (TSRI, La Jolla, CA), Vivek Malhotra (GRC, Barcelona, Spain), Adam Linstedt (CMU, Pittsburgh, PA), and Jennifer Lippincott-Schwartz (NIH, Bethesda, MD) for valuable reagents.

Footnotes

  • Competing interests

    The authors declare no competing financial interests.

  • Author contributions

    D.K., K.R.L., K.S., S.C.W. and M.A. designed, performed and interpreted various experiments. M.A. conceived and directed the project and wrote the manuscript with input, editing and comments from all authors.

  • Funding

    The study was supported by the National Institutes of Health (NIH) [R01DK092807 to M.A.]. Deposited in PMC for release after 12 months.

  • Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.138784/-/DC1

  • Received July 19, 2013.
  • Accepted January 22, 2014.

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Research Article
A cascade of ER exit site assembly that is regulated by p125A and lipid signals
David Klinkenberg, Kimberly R. Long, Kuntala Shome, Simon C. Watkins, Meir Aridor
Journal of Cell Science 2014 127: 1765-1778; doi: 10.1242/jcs.138784
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