COPI coated vesicles mediate trafficking within the Golgi apparatus and between the Golgi and the endoplasmic reticulum. Assembly of a COPI coated vesicle is initiated by the small GTPase Arf1 that recruits the coatomer complex to the membrane, triggering polymerization and budding. The vesicle uncoats before fusion with a target membrane. Coat components are structurally conserved between COPI and clathrin/adaptor proteins. Using cryo-electron tomography and subtomogram averaging, we determined the structure of the COPI coat assembled on membranes in vitro at 9 Å resolution. We also obtained a 2.57 Å resolution crystal structure of βδ-COP. By combining these structures we built a molecular model of the coat. We additionally determined the coat structure in the presence of ArfGAP proteins that regulate coat dissociation. We found that Arf1 occupies contrasting molecular environments within the coat, leading us to hypothesize that some Arf1 molecules may regulate vesicle assembly while others regulate coat disassembly.https://doi.org/10.7554/eLife.26691.001
Coated vesicles mediate transport between intracellular organelles. Coat protein 1 (COPI) coated vesicles function in retrograde trafficking between the Golgi apparatus and the Endoplasmic Reticulum (ER), and between the Golgi cisternae in both retrograde and anterograde directions (Cosson and Letourneur, 1994; Letourneur et al., 1994; Orci et al., 1997). Assembly of a COPI coated vesicle is initiated by recruitment of the small GTPase Arf1 (ADP-ribosylation factor 1) to the membrane, where it is activated by a guanine exchange factor and transitions into a GTP-bound state in which its amphipathic N0 helix is inserted into the membrane. Arf1-GTP recruits the protein complex coatomer from the cytoplasm and anchors it at the membrane. Polymerization of coatomer at the membrane assembles the COPI coat, which recruits cargo molecules and membrane machinery to the assembly site (Harter and Wieland, 1998; Jackson et al., 2012). Growth of the coat, and the resulting membrane bud, increases until a vesicle is pinched off the donor membrane. GTP hydrolysis by Arf1, activated by ArfGAPs (Arf GTPase Activating Proteins), mediates uncoating of the vesicle which subsequently fuses with its target membrane.
Coatomer (the COPI complex) is a heteroheptameric protein complex, consisting of α, β, β’, γ, δ, ε and ζ subunits. Coatomer can be subdivided into an outer-coat and an adaptor subcomplex (Lowe and Kreis, 1995). The outer-coat subcomplex consists of α, β’ and ε subunits. α- and β’-COP each contain two WD40 β-propeller domains followed by an α-solenoid, a structural motif that is shared by other coat proteins such as clathrin and sec13/31 from COPII (Devos et al., 2004). The adaptor subcomplex consists of β, γ, δ and ζ subunits and is homologous to the clathrin adaptor AP complexes (Schledzewski et al., 1999). Within the adaptor subcomplex, βδ and γζ are structural homologues that are likely to have arisen by gene duplication. Each of the large adaptor subunits β- and γ-COP contains an α-solenoid ‘trunk’ domain, connected by an unstructured linker region to a small appendage domain. The trunk domains each interact with an Arf1 molecule to anchor the coat to the membrane. In contrast to clathrin and COPII, where outer-coat and adaptor are separate complexes, in COPI they form a single protein complex in the cytoplasm that is recruited to the membrane en bloc (Hara-Kuge et al., 1994).
Insights into the structure and function of coated vesicles have been obtained by crystallography and cryo-electron microscopy (cryoEM) (Faini et al., 2012; Fotin et al., 2004; Lee and Goldberg, 2010; Shen et al., 2015; Stagg et al., 2006). We recently determined the architecture of the COPI coat assembled on the vesicle membrane in vitro using cryo-electron tomography (cryoET) and subtomogram averaging (SA) (Dodonova et al., 2015). We were able to fit homology models and available crystal structures into our cryoET structure to build a molecular model of the assembled coat. Although the COPI adaptor and outer-coat subcomplexes share multiple structural and functional similarities with the clathrin coat, they appear to be organized very differently. The COPI adaptor and outer-coat subcomplexes both form arch-like arrangements on the membrane and together form a highly interconnected coat lattice: there is no clear division of function between outer-coat and adaptor subcomplexes (for consistency we will nevertheless use this nomenclature here).
Multiple subunits in the assembled COPI coat interact with cargo and membrane machinery. The outer-coat COPI subunits α and β’ interact with dilysine K(X)KXX cargo motifs (Jackson et al., 2012; Ma and Goldberg, 2013), while the adaptor subunit γ interacts with p23 transmembrane peptides (Cosson and Letourneur, 1994; Harter and Wieland, 1998). COPI coated vesicles are also involved in the transport of K/HDEL cargoes associated with specific receptors (Semenza et al., 1990) that interact with the COPI coat (Majoul et al., 2001). The transport of HDEL cargoes is dependent on the presence of an amphipathic helix in δ-COP (Arakel et al., 2016).
Disassembly of the coat is facilitated by ArfGAPs that induce GTP hydrolysis in Arf1 and coat disassembly using a highly conserved catalytic zinc-finger domain (Cukierman et al., 1995). Two classes, ArfGAP1 and ArfGAP2/3, have been implicated in COPI vesicular transport (Frigerio et al., 2007; Liu et al., 2005; Poon et al., 1999). ArfGAP1 contains membrane-binding ALPS (Amphipathic Lipid Packing Sensor) motifs that allow the protein to be recruited to membranes in the absence of coatomer (Bigay et al., 2005; Weimer et al., 2008). ArfGAP2 and 3 interact with COPI via their non-catalytic, C-terminal regions (Frigerio et al., 2007; Kliouchnikov et al., 2009; Yahara et al., 2006) and require coatomer for efficient membrane recruitment and activity (Szafer et al., 2001). ArfGAP2 but not ArfGAP1 was found to be associated with COPI coated vesicles budded from Golgi membranes in vitro (Frigerio et al., 2007). Although the functions of all three ArfGAP proteins overlap (Saitoh et al., 2009), ArfGAP2 deletion leads to a much stronger retrograde trafficking phenotype than ArfGAP1 deletion (Poon et al., 1999). Why COPI is regulated by multiple and different ArfGAP proteins remains unclear.
To understand the structure of the COPI coat, and how its assembly and disassembly are regulated, we wished to generate a complete structural model for the assembled coat at sufficient resolution to precisely position the secondary structure elements of the component proteins. Crystal structures were available for the majority of the components of the COPI coat, but not for the βδ-COP subcomplex. As a first step towards a complete structural model, we expressed and crystallized a complex containing β-COP19-391 and δ-COP1-175 from C. thermophilum (Ct), from which we obtained a molecular model of βδ-COP comprising residues 20–390 of β-COP (‘trunk region’) and residues 2–150 of δ-COP (Figure 1, Table 1). The overall architecture of βδ-COP is similar to that of the homologous γζ-COP (Yu et al., 2012). β-COP(20-390) is a curved HEAT-repeat type α-solenoid consisting of 20 consecutive α-helices. The right-handed super-helical arch of β-COP wraps around the δ-subunit. The structural model of δ-COP comprises a longin domain followed by a β-turn (117-122), and two helices a (127-135) and b (139-150), which are part of the linker region connecting the longin domain and the μ-Homology Domain (MHD). The stretch of residues between helix a and amphipathic helix b traverses the β-COP α-solenoid in the vicinity of helix 7. Helix b, including isoleucine residues 143, 146 and 147, projects away from the α-solenoid (Figure 1—figure supplement 1) where it would be accessible for binding partners. These isoleucine residues have been shown to be essential for correct trafficking of HDEL motif containing cargo by COPI (Arakel et al., 2016).
We prepared COPI coated vesicles in vitro as described previously (Dodonova et al., 2015) and vitrified them for cryoET. Tomographic data was collected making use of a direct electron detector and optimized data collection conditions (Hagen et al., 2017). Subtomogram averaging was performed essentially as described in the Materials and methods, and the structure was further refined by combining multiple locally-aligned structures. From 1733 vesicles and near-complete buds we obtained a structure of the leaf (representing one coatomer complex with two Arf1 molecules), the asymmetric unit of the COPI coat, at 9.2 Å resolution (Figure 2, Figure 2—figure supplement 1, Video 1). At this resolution, distinct α-helical densities and β-propeller blades are resolved (See also Figure 3A and B).
In order to generate a complete pseudo-atomic model of the COPI coat, we performed rigid body fitting of the available crystal structures of COPI components, including our new x-ray structure of βδ-COP, into the structure of the leaf. Next we fitted the homology models of domains for which high-resolution structures are not available (α-COP 327–813, β-COP 410–968, γ-COP 312–549), into the leaf EM map. It is important to note, that the COPI complex is highly conserved among eukaryotes. The sequence identity between the βδ-COP from Chaetomium thermophilum used for crystallography and Mus musculus used for subtomogram averaging is 53%. The excellent match between the fitted structures and the density further confirmed the structural conservation of the complex as well as the quality and validity of the map (see also Figure 1—figure supplement 1E). Flexible fitting (Trabuco et al., 2008) was performed to relax the structures and models into the map and generate an initial structural model for the assembled coat. While the core subunits are in positions very similar to those in our previous model (Dodonova et al., 2015), there are larger changes in the positions of subunits for which only homology models were available. The improved resolution allowed us to resolve ambiguities in the orientations of appendage domains (Figure 3). Most importantly, since individual secondary structure elements are resolved, we were able to identify and assign structural elements that function separately from the folded domains (see below). The higher resolution of the EM map when compared to our previously published study (Figure 2A–B, Figure 2—figure supplement 1C–E), and the availability of the βδ-COP structure, therefore allowed us to generate a more complete and significantly improved pseudo-atomic model of the coat (Figure 2C–D).
The γ- and β-COP appendage domains are homologous to the α and β2 ear/appendage domains of the clathrin adaptor AP-2. Both appendage domains are important for viability in yeast (DeRegis et al., 2008; Hoffman et al., 2003). In our previous model we found that the γ-COP appendage domain is bound to the second β-propeller of β’-COPI, while the β-COP appendage domain is tucked into the center of the leaf between α-COP and γ-COP. Our new structure shows that γ-COP interacts with β’-COP via a highly conserved interface in the base of the γ-COP β-sandwich subdomain (Figure 3A,B,C). The platform subdomain of the γ-COP appendage is exposed (Video 1) and easily accessible for other binding partners such as ArfGAP (Watson et al., 2004). The β2 and α2 appendage domains of AP2 are also thought to function as recruitment hubs for interaction partners, suggesting that these domains are also in an accessible location within the clathrin vesicle (Praefcke et al., 2004; Schmid et al., 2006). To date, there are no structures available showing the positions of the AP2 appendage domains relative to AP2 or the clathrin coat.
We were previously unable to determine the orientation of the β-COP appendage domain. In the new structure, the β-sandwich and the platform subdomains of the β-COP appendage are clearly visible in the EM map (Video 1). The platform interacts with the trunk domain of γ-COP, whereas the β-sandwich forms a conserved interface with α-COP including both α-COP β-propeller domains and part of the α-COP α-solenoid domain (Figure 3D). In the clathrin system, the β2-appendage domain of AP2 plays a conceptually similar role, interacting with clathrin and promoting cage formation (Owen et al., 2000). The β2-appendage is thought to be linked to the core of AP2 only by its flexible linker. In contrast, the β-COP appendage appears to form the main link between the adaptor and the outer-coat subcomplexes of the COPI coat. The other connection is a rather small interface between the β-COP trunk and β’-COP. We speculate that the position of the β-COP appendage domain would allow it to modulate the conformation of coatomer by fixing the angle between the two β-propeller domains of α-COP and by forming a rigid buttress between the adaptor subunit γ-COP and the outer-coat subunit α-COP (Figure 3D, Video 1), thereby determining their relative arrangements. Thus, the β-COP appendage appears to act as a keystone in the assembled COPI coat.
After fitting known structures and homology models to generate the initial structural model, we identified seven positions in the map where there were substantial regions of electron density not occupied by any of the fitted coatomer domains (see Materials and methods) (Figure 4A). We wished to identify the protein components that contribute these extra densities. We first performed secondary structure predictions for all COPI subunits using the Quick2D server (Jones, 1999; Ouali and King, 2000). We identified putative secondary structure elements in multiple COPI subunits that were outside the regions included in the crystal structures and homology models of the COPI domains. We compared the sequence positions of these elements with the locations of the unoccupied densities, as well as with available cross-linking mass-spectrometry (XL/MS) data, thereby assigning secondary structure elements to unoccupied densities.
The C-terminal region of β’-COP was predicted to contain two additional α-helices, consistent with the presence in the EM density of elongated, unoccupied densities adjacent to the C-terminus of β’-COP at the β’-α interface (Figure 4A,B ‘1’). We speculate that these helices may stabilize or regulate flexibility at the β’-α solenoid-solenoid interface.
We interpret the unoccupied density adjacent to the N-terminal β-propeller of α-COP, in the vicinity of the Arf1 bound to β-COP (βArf1), as being partly contributed by a flexible hydrophobic loop of the β-propeller itself that was absent from the crystal structure of the propeller domain (Figure 4C ‘2’) (PDB:4J87, [Ma and Goldberg, 2013]). Flexibility of the loop likely results in smearing of the density, and we cannot rule out that this density also contains other protein components (e.g. unstructured linker regions).
Five additional α-helices were predicted C-terminal to the β-trunk domain, consistent with the presence of several unoccupied densities near the C-terminal end of the fitted β-trunk model (Figure 4A,D ‘3’ and ‘4’). Cross-links between these extra helices and the ζ-longin domain support their location in that region (β617- ζ39, β618- ζ39) (Table 2). After these additional helices, the 103 amino acid unstructured β-COP linker extends in the sequence until the start of the β-appendage domain. The theoretical fully extended length of the linker is around 370 Å. Within the assembled coat it must span a distance of approximately 100 Å. Several cross-links between the linker, the β-COP appendage domain, β’-COP, and α-COP (Table 2, rows 5–15) are consistent with the expected route of the linker through the assembled coat, schematically shown in Figure 3F. The small density ‘6’ (Figure 4A), may represent a short helical stretch in the linker bound to the β-trunk. Density ‘5’ is discussed below.
Secondary structure predictions indicate the presence of three putative α-helices (referred to as ‘a, b and c’) in the linker-region between the δ-COP longin domain and MHD (Figure 5A).
Helix a and the first part of helix b are present in our x-ray structure (Figure 1). One short and one long extended rod-like density corresponding to these helices are also observed in equivalent positions immediately C-terminal to the fitted δ-COP longin domain in our COPI coat EM density (Figure 5B). While no electron density was visible for residues 151–175 of δ-COP in the x-ray structure, our cryoEM map of the COPI coat has clear density corresponding to a longer helix, and we modeled residues 139–165 of the predicted helix b into our density (Figure 5).
Helix a interacts with the δ-COP longin and the β-COP trunk domains. The N-terminal part of helix b interacts with the β-COP trunk domain, while its C-terminal part interacts with the peripheral Arf1 molecule (βArf1) in the triad (Figure 5B). The very C-terminal end of helix b also contacts the membrane surface (Figure 5C). An equivalent helix b is predicted in the sequence of the homologous ζ-COP (Alisaraie and Rouiller, 2012), but we saw no equivalent density near ζ-COP in our EM structure.
In an electron microscopy structure of a soluble trimeric assembly of AP-1, Nef, and Arf1 (Shen et al., 2015) an interaction was observed between the µ1 subunit of AP-1 (homologous to δ-COP), and Arf1. The interaction made by µ1 is very different to that made by δ-COP, since it involves the MHD of µ1 instead of the longin domain, and a different surface of Arf1.
Several cross-links (δ233-δ263, δ243-δ263, δ233-β’627 and δ241-β’627) (Table 2) suggest the approximate location of helix c (Figure 6—figure supplement 1), and based on these observations we speculate that density ‘5’ is occupied by this α-helix (Figure 4A,D ‘5’). In this position, helix c could help to coordinate the positioning of the C-terminal δ-COP MHD, which could otherwise be located a long distance from the vesicle: there are ~100 amino acids between helix b and the MHD.
Further validation of the interaction between δ-COP helix b and Arf1 is provided by published photo-cross-linking data (Sun et al., 2007), which showed a cross-link between Arf1 residue 167 and δ-COP (Figure 6C, Arf167 is marked), as well as by mass-spectrometry cross-linking data that showed cross-links between the second longer helix b and Arf1 (δ142-Arf36, δ164-Arf36) (Table 2, Figure 6C). We prepared δ-COP variants containing photolabile amino acids in position 156 or 159. These variants were expressed as subcomplexes with β-COP(19-391) and recruited to liposomes in an Arf- and GTP-depended manner. Subsequent photo-cross-linking resulted in an 80 kDa product that was confirmed by western blotting to consist of Arf1 and δ-COP (Figure 6A). We also prepared an Arf1 variant with two photolabile amino acid derivatives in positions 46 (known to cross-link to β-COP [Sun et al., 2007]) and 167. This Arf1 could be photo-cross-linked within vesicles to give a 180 kD product, corresponding to β-COP, δ-COP and Arf1 (Figure 6B, and Figure 6—figure supplement 2), confirming that one Arf1 molecule interacts with both β- and δ-COP within the same coatomer complex.
To dissect the role of the δ-COP linker-region (δCOP 117–271) in binding to Arf1, we made four δ-COP constructs incorporating different parts of this linker region. These terminate after helix a (δ1–137), helix b (δ1–175) or helix c (δ1–243) or code for full-length δ-COP. These proteins were expressed as subcomplexes with β-COP(19-391) and tested for binding to Arf1 in its GMPPNP-loaded state using pulldown assays (Figure 6D). The complex containing δ1–137 showed very little GTP dependent Arf1 binding, δ1–175 showed an intermediate level, while δ1–243 bound Arf1 in a GTP dependent manner at the same level as the complex containing full length δ-COP (Figure 6E). These results indicate that the interaction between βδ-COP and Arf1-GTP is stabilized by δ-COP helix b and is further stabilized by downstream regions of δ-COP including helix c. These observations are consistent with our structural model for the assembled coat in which helix b interacts directly with Arf1, and helix c is bound to β-COP to stabilize the complex.
δ-COP helix a is analogous to the α5 helix in the AP2 μ2-subunit linker, and they are positioned similarly near the β-COP/β2-AP trunk. Helix b is a conserved, COPI-specific feature, at a sequence distance similar to that of the ‘bind-back’ helix in AP2 μ2 (Arakel et al., 2016; Jackson et al., 2010). Arakel and colleagues proposed two models to explain a requirement of helix b for retrieval of HDEL/KDEL proteins (Arakel et al., 2016). In a first model, this helix was proposed to bind back into a furrow in β-COP, thereby preventing destabilization of the β-COP α-solenoid. In a second model, helix b, which is amphipathic in nature, binds to the membrane, bringing coatomer into close proximity where it can more easily interact with retrieval signals. Our EM structure confirms that helix b is located proximally to the membrane (Figure 5C), however the contact with the membrane is small and involves only the C-terminal end of the helix. The distance between the membrane and the nearest conserved hydrophobic residue is too large for a direct contact. This argues against it being a conventional membrane-inserting amphipathic helix (see Figure 5C). Our structure suggests that the main interaction partner of helix b is Arf1.
δ-COP helix b interacts directly with the Arf1 Switch I region, Interswitch and C-terminal helix (Figure 5B) (in contrast to the interactions observed between AP1 µ1 subunit and Arf1, which do not involve the switch regions of Arf1 [Shen et al., 2015]). δ-COP helix b binds the surface of Arf1 in a region that is accessible when Arf1 is in the GTP state and the amphipathic N0 helix is inserted into the membrane, but is occupied by the N0 helix when Arf1 is in the GDP state (PDB: 1MR3, [Amor et al., 1994]) (Figure 5D). The exposure of this region in Arf1-GTP when N0 moves to insert into the membrane may contribute to the nucleotide-dependent recruitment of coatomer. These interactions also reconcile the apparent nucleotide independence of the interaction between isolated δ-COP and NΔ17Arf1 in solution (Sun et al., 2007) with the nucleotide dependence of the interaction between δ-COP and Arf1 during coatomer recruitment to membranes (Figure 6B): the truncation of the N0 helix in NΔ17Arf1 exposes the δ-COP binding site regardless of Arf’s nucleotide state.
These observations suggest a mechanistic model for the role of δ-COP helix b in COPI function. Upon binding of the Arf1 N0 amphipathic helix to the membrane, the binding site for δ-COP is exposed, and the resulting interaction contributes directly to Arf1-dependent coatomer recruitment to the membrane. The interaction of δ-COP may additionally stabilize Arf1 in its active GTP-bound state by binding its Switch and Interswitch regions. The interaction of δ-COP helix b with the Arf1 GTPase, with the membrane, and possibly with cargo confirms and explains its critical role in regulating COPI function.
We note a further implication of this model: GTP-hydrolysis by Arf1 can directly modulate the conformation of δ-COP. Such modulation may be transmitted to other binding partners such as the KDEL cargo receptor, providing a possible route to link coat hydrolysis to cargo binding/release.
We also determined the structures of the linkages between triads as previously described (Dodonova et al., 2015) (Figure 2—figure supplement 2). The linkage structures are at lower resolution than that of the leaf, so we fitted our leaf structure as a rigid body (see Materials and methods and Figure 2—figure supplement 2) to assess which subunits are close to one another in the linkages. Consistent with our previously published structures (Dodonova et al., 2015) the central contacts in linkages I and IV are made by the αε-COP subunits, and those in linkage II are made by the δ-COP MHD. Interestingly neither ε-COP nor the δ-COP MHD are essential for COPI function and the combined deletion of both is not lethal for yeast (Arakel et al., 2016; Kimata et al., 2000). While the contacts formed by these proteins at the linkages may play a regulatory role, they are therefore not essential for COPI function.
The slightly improved resolution of the linkage structures obtained here compared to our previously published maps showed that the density that we interpret as being contributed by a flexible hydrophobic loop of the N-terminal β-propeller of α-COP, is in the vicinity of γ-COP in a neighbouring triad. We found that βArf1 is close to the N-terminal β-propeller of β’-COP in a neighbouring triad, and approaches the trunk domain of β-COP (Figure 2—figure supplement 2C). None of these contacts are similar to the previously described crystal contact between the α1 subunit of AP1 and the back side of βArf1 (Ren et al., 2013) suggesting that an equivalent interaction is not relevant within the COPI coat. While the low-resolution of the linkage structures precludes more detailed interpretation, the proximity of the peripheral βArf1 molecule to neighbouring triads suggests that βArf1 could modulate inter-triad interactions.
The γArf1 and βArf1 molecules have different interaction partners within the coat (γ-COP, and δ-COP and β-COP, respectively) and are in very different molecular environments (Figure 2D). These observations suggest that the two different Arf1 molecules in the coat are differentially regulated. To further investigate the regulation of Arf1 in the coat, we incubated COPI coated vesicles with ArfGAP1 or ArfGAP2 in the presence of a non-hydrolysable GTP analogue and determined their structures. Previously published biochemical data indicates that almost no ArfGAP1 binds to COPI vesicles generated in vitro in the presence of poorly hydrolysable GTP analogs, while ArfGAP2 is abundant in such vesicle fractions (Frigerio et al., 2007). Note, that in the presence of hydrolysable nucleotide GTP, ArfGAP1, 2 and 3 facilitate efficient vesicle uncoating (Weimer et al., 2008). We were unable to identify any ArfGAP1 bound to vesicles produced in the presence of GTPγS, while in vesicles incubated with ArfGAP2 we identified an additional density near the central γArf1 molecules in a subset of COPI leaves (Figure 7A,B and Figure 7—figure supplement 1). No significant additional density was observed near the peripheral βArf1 molecules or at any other position in the structure. The size and shape of the additional mass corresponded very well to the catalytic domain of the ArfGAP2 protein, which we fitted into the density (Figure 7C). To further validate this fit, we superimposed Arf6 from a structure in which it had been co-crystallized with the catalytic domain of the ArfGAP homologue ASAP3 (PDB:3LVQ, [Ismail et al., 2010]), with Arf1 in our coat structure. The resulting position of the catalytic domain of ASAP3 coincided with the additional density that we observe (compare Figure 7 and Figure 7—figure supplement 1C). In this structure, the catalytic domain of ArfGAP2 is positioned directly near the Arf1 nucleotide-site, where it can provide the Arginine ‘finger’ essential for stimulation of GTP-hydrolysis. The observed position of the ArfGAP catalytic domain differs from a previous ArfGAP1-Arf1 structure (Goldberg, 1999) in which the ArfGAP1 catalytic domain is distant from the nucleotide-site of Arf1 (Figure 7—figure supplement 1D).
We were not able to resolve the C-terminal, non-catalytic part of ArfGAP2, known to be involved in coatomer binding via interactions with the γ-COP appendage domain (Kliouchnikov et al., 2009; Watson et al., 2004), most likely because the non-catalytic part of the ArfGAP2 protein is largely disordered and may not form a globular domain that can be positioned at 12 Å resolution (Pietrosemoli et al., 2013).
The ArfGAP2 catalytic domain is bound into a niche in the assembled coat formed by Arf1, the γ- and β-COP adaptor subunits, and the β’-COP outer-coat subunit (Figure 7C and D). ArfGAP2 activity is increased by 100-fold in the presence of coatomer (Luo et al., 2009; Szafer et al., 2001). This stimulation requires the presence of both the adaptor subcomplex and outer-coat subcomplex (Pevzner et al., 2012), indicating that multiple ArfGAP-coatomer interactions are functionally important. Within a triad, an ArfGAP2-binding niche is formed by β’-COP and β-COP from one leaf and by γ-COP from a neighboring leaf (Figure 7D), with the central γArf1 molecules at the bottom. Thus, the complete binding site for the ArfGAP2 catalytic domain is formed when the coat is assembled. This suggests a possible proofreading mechanism: ArfGAP2 recruitment, and the resulting GTP-hydrolysis and coat dissociation can only occur once the coat is assembled, minimizing premature dissociation of coatomer from the membrane.
βArf1 molecules in the triad do not provide an equivalent binding niche for the catalytic domain. We did not observe ArfGAP2 bound near the βArf1 molecules. This is consistent with yeast-two hybrid experiments which showed an interaction between the ArfGAP2 Glo3 and β’-COP and γ-COP but not with other coatomer subunits (Eugster et al., 2000). The membrane surface is more exposed near the βArf1 molecules; we suggest that this may facilitate interaction with ArfGAP1 recruited directly to the membrane via its ALPS domains.
By combining the βδ-COP crystal structure and the in vitro EM structure of the COPI coat on the vesicle membrane, we have generated a model that reveals molecular details of the coat at the level of protein secondary structure, allowing precise positioning of protein domains and the interpretation of isolated secondary structure elements. The resulting structural model has provided novel functional insight and offers a basis for future concept-driven investigation of molecular mechanisms that underlie vesicular transport.
The two halves of the COPI adaptor subcomplex, γζ-COP and βδ-COP, are thought to have evolved by gene duplication. The structure reveals that they have functionally diverged in two important ways. Firstly, the appendage domains have divergent functions - the γ-appendage sits at the outside of the coat where it provides a binding site for regulatory factors. The β-COP appendage domain links the adaptor and the outer-coat COPI subunits within one coatomer molecule. This functional divergence seems to be mirrored in AP-1 and AP-2, although the details of the interactions are different. Secondly, the two halves of the adaptor subcomplex recruit and position Arf1 in two different regulatory environments – at the center of the triad, bound to γ-COP (γArf1), and at the periphery of the triad bound to β-COP and δ-COP (βArf1). Our structure shows that the two Arf1 molecules can be differentially regulated both during coatomer recruitment and coat disassembly - βArf1 by interactions of its Switch 1 and N0 helix regions with the essential helix b downstream of the longin domain in δ-COP, and γArf1 by recruitment of ArfGAP2. Differential regulation would allow fine-tuning of coat assembly and cargo recruitment. We speculate that βArf1 molecules may be primarily regulators of coat assembly – modulating coatomer recruitment, dissociation, and the interactions between triads that may be influenced by the presence of cargo, and in which the δ-COP helix b plays a key role. ArfGAP1, bound to the membrane, may function at this stage. We speculate that the γArf1 molecules, via their interaction with ArfGAP2, are the primary regulators of coat disassembly, unlocking the triad and triggering coat collapse upon hydrolysis.
Recombinant M. musculus coatomer was expressed and purified from SF9 insect cells (Invitrogen, Karlsruhe). Original SF9 cells were cloned from the parental IPLBSF-21 (Sf-21) cell line that was derived from the pupal ovarian tissue of the fall army worm, Spodoptera frugiperda. Invitrogen Sf9 cells were tested by the manufacturer for contamination of bacteria, yeast, mycoplasma and virus and were characterized by isozyme and karyotype analysis. We used a baculoviral expression system essentially as described previously (Sahlmüller et al., 2011) with a ‘One-Strep-Tag’ at the C-terminus of α-COP. Recombinant S. cerevisiae myristoylated Arf1, and human nucleotide exchange factor ARNO, were purified from E. coli as described previously (Chardin et al., 1996; Randazzo et al., 1992). Full-length N-terminally His-tagged R. norvegicus ArfGAP2 and ArfGAP1 proteins were expressed in insect cells and purified through Ni-NTA chromatography (Weimer et al., 2008), and gel filtration.
To determine the most stable domains of coatomer suitable for crystallization, limited proteolysis with subtilisin (Sigma Aldrich, St. Louis Missouri USA) was performed with C. thermophilum (Ct) coatomer and subcomplexes. 45–250 μg of the respective complex was treated at various molar ratios. The reaction was incubated for 15 min on ice and then stopped by addition of PMSF to a final concentration of 1 mM. The samples were separated by SDS-PAGE and the resulting fragments were analyzed by MS.
Expression plasmids for truncated forms of Ctβδ-COP subcomplexes were constructed using the pFBDM vector with a One-Strep-Tag fused the N-terminus of β-COP (Berger et al., 2004; Fitzgerald et al., 2006). Baculoviruses were generated for the subcomplexes Ctβ19-391δ, Ctβ19-391δ1–243, Ctβ19-391δ1–175 and Ctβ19-391δ1–137 by infecting SF9 cells (Invitrogen, Karlsruhe) with recombinant Bacmids prepared using E.coli Dh10b MultiBac cells. The dimeric coatomer subcomplexes were produced by co-expression of both subunits in Sf9 insect cells infected with the corresponding baculovirus. Insect cells were harvested 72 hr post infection. Cells were lysed in buffer (25 mM Tris, 300 mM NaCl, 1 mM DTT pH 8.0) using a high pressure Microfluidizer (Microfluidics, Newton USA) and cell debris was pelleted by centrifugation at 100 000 x g for 1 hr. The protein complex was purified using strep-tactin affinity chromatography according to the supplier´s instructions, followed by size exclusion chromatography (Superdex 75 column, GE Healthcare). Purified protein was concentrated using Amicon spin concentrators (Merck Millipore, Darmstadt) and stored at −80°C.
In order to obtain phases by anomalous x-ray diffraction Ctβ19-391δ1–175 subcomplex incorporating selenomethionine (MSE) was produced in Sf9 insect cells. Sf9 cells were grown in Δ921 Series methionine deficient medium (Expression Systems LLC, USA) supplemented with 150 mg/ml MSE.
Crystallization trials were performed with all construct variants described above and diffracting crystals could be obtained with polyethylene glycols as precipitant with a size range from PEG 3350 to PEG 8000. Best diffracting crystals were obtained with construct Ctβ19-391δ1–175 using sitting drop vapor diffusion in 96-well MRC UVP plates at 18°C with a precipitant and reservoir composition of 0.2 M magnesium formate, 0.1 M Tris pH 7.0, and 24% PEG3350. Crystallization drops consisted of 400 nl protein solution of Ctβ19-391δ1–175 at a concentration of 13 mg/ml and 400 nl of precipitant. The reservoir volume was 95 μl. Crystals were visible after 10 days. All measured crystals had the orthorhombic space group C2221 with typical cell dimensions of a = 137.59, b = 177.47, c = 62.72, α=β=γ=90°.
All X-ray diffraction data were collected at ESRF beamlines ID-29 (native) and ID-21 (MSE). Anomalous diffraction data was collected from crystals containing MSE labelled Ctβ19-391δ1–175. All data were integrated using XDS (Kabsch, 2010) and scaled with AIMLESS (Evans, 2006). Experimental phases were obtained with Phenix. autosol (Terwilliger et al., 2009) from anomalous diffraction data and an initial model was obtained using Phenix. autobuild (Terwilliger et al., 2008). The final model was built with Coot (Emsley et al., 2010) and refined with Phenix.refine (Afonine et al., 2012) using the native dataset (see Table 1). The asymmetric unit contains one molecule of Ctβ20-390δ1–150. Structure quality was evaluated using MolProbity (Chen et al., 2010). Interfaces and crystal contacts were analyzed using PISA (Krissinel and Henrick, 2007). Structure figures were created with Chimera (Pettersen et al., 2004).
For pulldown assays Arf1 (NΔ20CtArf) was cloned in the petM11 vector, adding an N-terminal HIS6-TAG, and expressed in E. coli BL21 (DE3) cells (Novagen, Merck, Darmstadt). NΔ20CtArf was purified by Ni-NTA affinity chromatography followed by gelfiltration on a Superdex 75 column (GE Healthcare, Buckinghamshire UK) in PD (Pulldown) buffer (25 mM Tris, 150 mM NaCl, 5 mM MgCl2, 1 mM DTT, pH 7.5).
To test coatomer βδ subcomplexes for binding of Arf1, 200 to 400 μg of the respective subcomplex was immobilized on 50 μl strep-tactin sepharose beads by incubation for one hour at 4°C on a rotary wheel. After immobilization of the subcomplex the beads were washed with PD buffer to remove unbound protein. The soluble form of Arf1-GTP was added to the immobilized subcomplexes and incubated at 4°C for one hour on a rotary wheel. After the incubation the beads were washed extensively with PD buffer. Next the beads were pelleted by centrifugation, resuspended in 50 μl buffer and samples were taken for each subcomplex and mixed with SDS sample buffer. After centrifugation, the proteins in the supernatants were analyzed by SDS-PAGE and western blot.
For photo-cross-linking experiments human Arf1 and yeast N-myristoyl-transferase were cloned in pETDuet vector. Amber stop codons were introduced in Arf1 at positions 46 and 167 by point mutation. Methionine aminopeptidase for improvement of myristoylation efficiency was cloned into pRSF-Duet1 vector. Both plasmids were co-transformed in E. coli Bl21 (DE3) cells harbouring the pEVOL plasmid coding for orthogal tRNA recognizing the Amber stop codon and the tRNA synthetase (Chin et al., 2002). Cells were grown to OD600 of 0.6 at 37°C. After addition of 65 μM of sodium myristate and 1 mM of p-benzoyl-I-phenylalanine, cells were shifted to 27°C. Expression was induced 1 hr after the temperature shift by addition of IPTG (0.5 mM) and arabinose (0.5%) and was continued for 22 hr at 27°C. Photolabile Arf-I46Bp-Y167Bp was purified from the cleared lysate in the presence of 2 mM GDP by size exclusion chromatography on Superdex 200 (GE Healthcare, Buckinghamshire UK) in 25 mM Tris, 150 mM KCl, pH 7.0.
This bivalently photolabile Arf1 was used in a reconstitution assay with liposomes, GTP and complete COPI. For the reaction 5 μg Arf-I46Bp-Y167Bp was mixed with 0.4 μg ARNO, 100 μM GTPγS and 500 μM Golgi-like liposomes in a final volume of 100 μl in 20 mM MOPS pH 7.2, 150 mM KOAc 2 mM Mg(OAc)2, and incubated at 37°C for 10 min. Alternatively 50 μg of Golgi membranes in the presence of 200 mM Sucrose were incubated with 5 μg of the respective photolabile Arf1 with or without 100 μM GTPγS in a final volume of 100 μl buffer at 37°C for 10 min. In a second step, coatomer was added and incubation was resumed at 37°C for 10 min. Membranes were pelleted by centrifugation at 16,000 x g for 30 min at 4°C. Golgi membranes from the liver of male 200 g Wistar rats (Rattus norvegicus) were pelleted through 330 μl of 15% (v/v) Sucrose. The pellet was re-suspended in 10 μl buffer and irradiated on ice with 15 × 1 s UV366 pulses with 1 s pauses. After irradiation the sample was analysed by MS, SDS-PAGE, and western blot using antibodies directed against β-COP, δ-COP and Arf1. Additionally photolabile amino acids were introduced in position 156 or 159 of δ-COP (M. musculus) using the same approach. These δ-COP variants were introduced in βδ-COP subcomplexes together with Ctβ19–391 producing a chimeric βδ subcomplex. This was done as no antibodies were available against Ctδ-COP. The chimeric subcomplexes were tested in the reconstitution assay with liposomes, GTP and Arf1 as described above.
Giant Unilamellar Vesicles (GUVs) were prepared by electroformation (Angelova et al., 1992) from the Golgi-like lipid mix (Bigay and Antonny, 2005). COPI-coated vesicles were produced in vitro by incubating coatomer (840 nM), Arf1 (2 μM), GTPγS (1 mM), ARNO (1.5 μM) and 2 μl GUVs in a total volume of 40 μl for 30 min at 37°C. The budding reaction buffer contained 50 mM HEPES pH 7.4, 50 mM KOAc, 1 mM MgCl2. Protein-A conjugated 10 nm gold was added to the reaction mix in 1:6 vol ratio and the sample was applied onto glow-discharged (30 s, 20 mA) C-flat (Protochips Inc.) multihole grids. The grids were blotted from the back side for 11 s at room temperature in a chamber at 85% humidity and plunge-frozen into liquid ethane using a manual plunger.
In order to test activity of the ArfGAP1 and ArfGAP2 proteins, the COPI budding reaction was performed in the presence of GTP, and the reaction mix was incubated for 30 min at 37°C. ArfGAP1, ArfGAP2 or buffer (as a control) were added to the mix in 10 molar excess to coatomer and after 15 min incubation the reaction was plunge-frozen. All samples were imaged in an electron microscope. The control samples contained coated vesicles and buds, whereas the samples incubated with ArfGAP1 or ArfGAP2 contained only naked liposomes. The functionality of ArfGAP proteins in vitro was also shown previously in budding assays containing rat liver Golgi membranes (Weimer et al., 2008).
To explore the structure of the coat in the presence of ArfGAP1 or ArfGAP2, the budding reaction mix was incubated for 30 min prior to addition of a 10 molar excess of ArfGAP1 or ArfGAP2 in the presence of GTPγS nucleotide. The mix was incubated for a further 15–20 min, protein-A conjugated 10 nm gold was added in a 1:6 vol ratio, and the sample was plunge-frozen.
The plunge-frozen COPI-coated vesicles were imaged in a FEI Titan Krios electron microscope operated at 300 kV and equipped with a Gatan Quantum 967 LS energy filter with a 20 eV energy slit and Gatan K2xp direct electron detector (Gatan Inc.). Tomographic tilt series were acquired with the dose-symmetric tilt-scheme (Hagen et al., 2017) over an angular range of ±60° with a 3° increment and a total electron dose of approximately 85 e/Å2. The defocus values ranged from −2.0 to −5.0 um. Data acquisition was controlled using the SerialEM software package (Mastronarde, 2005). Five frames were collected in super-resolution and electron-counting mode at each tilt. The super-resolution pixel size at the specimen level was 0.89 Å. The frames were Fourier-cropped, motion-corrected with the K2Align package based on the MotionCorr algorithms (Li et al., 2013) and integrated together. Each of the images in the tilt series was low-pass filtered according to the electron-dose acquired by the sample (Grant and Grigorieff, 2015). Motion-corrected and dose-filtered tomograms were reconstructed in Imod (Kremer et al., 1996).
1733 vesicles and near-complete buds were picked from 61 tomograms. CTF-determination for each individual tilt image was performed using CTFFIND4 (Rohou and Grigorieff, 2015). Strip-based CTF-correction and tomogram reconstruction was performed in Imod. Subtomograms were extracted from the surface of the vesicles. Subtomogram averaging was performed using scripts derived from the TOM and Av3 software packages (Förster et al., 2005; Nickell et al., 2005) and Dynamo (Castaño-Díez et al., 2012). Each dataset was split into two halves, each half including odd- and even-numbered vesicles. The COPI triad structure (EMDB-2985, [Dodonova et al., 2015]) was low pass filtered to 55 Å and used for the initial alignment step. All further processing steps were performed completely independently on the two half datasets. Preliminary processing was performed using 4x binned data without CTF correction (pixel size 7.12 Å). C3 symmetry was applied. The references were low pass filtered to 35 Å at each alignment step. Iterative rotational and translational alignments were performed until convergence. Next the dataset was cleaned to remove misaligned subtomograms based on cross-correlation coefficient threshold, and duplicates were removed based on the mutual distance between neighboring subtomograms. Final subtomogram alignments were performed on the unbinned (pixel size 1.78 Å) and CTF-corrected data with the low pass filter set to 16 Å. The half datasets contained 19,343 and 19,155 asymmetric units. At the final alignment steps each asymmetric unit of the triad, ‘the leaf’, representing a COPI molecule and two Arf1 molecules, was processed separately in order to compensate for coat movements and deviation from C3 symmetry. The FSC was calculated for the two final references masked with a soft cylindrical mask and the measured resolution at FSC = 0.143 was 9.4 Å (Figure 2—figure supplement 1A).
The local resolution of the EM map was estimated within a small floating window moving within the whole map. The local resolution was highly variable and ranged from 8.7 to 12.7 Å within the protein density (Figure 2—figure supplement 1B). We therefore performed multiple local alignments using soft cylindrical masks focused on different parts of the structure. Finally, all the local averages were masked with the corresponding alignment masks, added together, and the map was normalized within the volume enclosed by all local masks. The local resolution ranged from 8.1 to 11.5 Å. The global resolution was 9.2 Å at FSC 0.143. The final structure (Figure 2B) was B-factor sharpened (B-factor=−1400) (Rosenthal and Henderson, 2003).
The COPI triads can be arranged into four distinct types of linkages within the coat (Faini et al., 2012). We used the approach described in detail in (Dodonova et al., 2015; Faini et al., 2012) to determine their positions and orientations. Next we performed subtomogram averaging on each of the linkage datasets to determine their structures (Figure 2—figure supplement 2). Each set was processed in two independent halves, which were split based on the odd or even sequential number of the vesicle. C2 symmetry was applied to the linkage III and IV sets. The starting references were obtained by averaging the subvolumes at their initial positions. The alignments were performed iteratively starting with a full search for the in-plane Euler angle and continued with a progressively decreasing angular search step. Duplicate and misaligned subtomograms were removed. Next the subvolumes were extracted at the positions defined during previous iterations from the CTF-corrected unbinned tomograms, and final alignments were performed. The linkage datasets contained 2547, 3312, 140, 1640 asymmetric units from both half sets. The final resolutions of linkages I, II, and IV at FSC 0.143 were respectively 17, 15, 17.3 Å. Linkage III had low abundance in the dataset and the resulting resolution was 31 Å.
In order to generate pseudo-atomic models of the linkages, first, the model of the COPI leaf generated by homology modeling and flexible fitting into the 9.2 Å leaf EM map (see below) was fitted as a rigid body into each of the linkage EM maps. Next, homology models of linkage-specific subunits (α-COP C-terminal domain together with ε-COP and the MHD of δ-COP) were generated in Modeller and fitted as rigid bodies into the central densities in the linkage EM maps.
First the available crystal structures of COPI components were fitted as rigid bodies into the structure of the leaf: Arf1-γζ-COP (PDB:3TJZ [Yu et al., 2012]), αβ’-COP (PDB:3MKQ [Lee and Goldberg, 2010]), the N-terminal β-propeller of α-COP (PDB:4J87 [Ma and Goldberg, 2013]), and the γ-COP appendage (PDB:1PZD [Hoffman et al., 2003]). Moreover, several crystal structures corresponding to the densities in the inter-triad linkages were also available: the αε-COP structure (PDB:3MKR and 3MV2 [Hsia and Hoelz, 2010; Lee and Goldberg, 2010]), the δ-COP MHD (PDB:4O8Q [Lahav et al., 2015]).
Homology models of COPI subunits were generated automatically using the HHpred and Modeller servers (Söding et al., 2005; Webb and Sali, 2014), as described previously (Dodonova et al., 2015). The model for β-COP (residues 410–968) was generated based on the structures of the homologous AP2 β2 subunit (PDB:2XA7 and PDB:2G30) (Edeling et al., 2006; Jackson et al., 2010); the model for γ-COP (residues 312–549) was generated based on the structure of the AP2 α2 (PDB:2XA7) (Jackson et al., 2010). α-COP residues 327–813 were modeled based on the β’-COP structure (PDB:3MKQ) (Lee and Goldberg, 2010). We performed ‘structure-guided’ modeling for the C-terminal parts of the β-COP and γ-COP trunk domains, which were absent from the starting x-ray structural models. The γ-COP and β-COP trunk C-terminal parts were modeled as separate batches of 2–3 helices, which were sequentially fitted into the distinct α-helical EM densities. Loops connecting the batches of helices were added in Modeller.
The structures of all remaining COPI subunits or parts of subunits were available from the PDB and were modeled to represent the M. musculus sequence.
After flexible fitting of all COPI subunits into the map, several small regions of the map remained unoccupied. The unoccupied ‘extra’ densities were identified by subtracting the density occupied by the fitted model from the EM map. The density occupied by the fitted model was generated by the chimera molmap command at 9 Å resolution. The resulting difference map was segmented and the volumes of all separate extra densities were plotted (Figure 4—figure supplement 1). The seven large extra densities were numbered according to their sizes (Figure 4—figure supplement 1, numbered): ‘1’ near the β’-α-COP interface; ‘2’ near the N-terminal β-propeller domain of α-COP; ‘3’ and ‘4’ near the β-γ-COP trunks interface; ‘5’ near a β-β’-COP contact site; ‘6’ near the β-δ interface; ‘7’ near the Arf1-β-δ interface;
The COPI-ArfGAP2 dataset consisted of 27 tomograms containing 690 vesicles and near-complete buds. Subtomogram averaging and alignments were performed for two independent half datasets, which contained either odd- or even-numbered vesicles and consisted of 11,331 and 11,004 asymmetric units respectively. The subtomogram averaging processing pipeline was exactly the same as for the COPI dataset. The final measured resolution of the leaf at the FSC = 0.143 was 9.8 Å. We calculated the difference map between the final COPI-ArfGAP2 EM map and the control COPI EM map. An additional density was visible near the Arf1 molecules located at the center of the triad. Multireference alignment and classification on the complete leaf structure was performed in order to identify the subpopulation of leaves with bound ArfGAP2. To do this, the subtomograms were aligned against the reference from the COPI-ArfGAP2-dataset and from the COPI-dataset and divided into two classes based on cross-correlation. The average structures were generated for both classes. This process was iterated a total of three times. The final COPI-ArfGAP2 class contained 6280 and 6092 asymmetric units in two independent half datasets, comprising in total approximately 65% of the initial dataset. The resolution was 10.1 Å at FSC = 0.143. The second class, which did not contain additional ArfGAP density, consisted of 3497 and 3481 asymmetric units and the resolution at the 0.143 FSC was measured to be 11.7 Å. The final difference map was calculated between the structures produced from the two classes (Figure 7—figure supplement 1).
The linkage maps from the COPI-ArfGAP2 were calculated and upon comparison with the control maps did not show any additional densities except that described above.
The COPI-ArfGAP1 dataset comprised 24 tomograms containing 833 vesicles and near-complete buds. Subtomogram averaging and alignments were performed for two independent halves of the dataset. The two half datasets contained 14,613 and 14,379 asymmetric units. The subtomogram averaging processing pipeline was exactly the same as for the COPI and COPI-ArfGAP2 datasets. The final measured resolution of the COPI leaf structure at the FSC = 0.143 was 9.6 Å. We calculated the difference map between the final COPI-ArfGAP1 leaf EM map and the control COPI EM map. No large additional densities were observed. The structures of the linkages were calculated and upon comparison with the control maps did not show any additional densities.
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Margaret S RobinsonReviewing Editor; University of Cambridge, United Kingdom
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
Thank you for submitting your article "9Å structure of the COPI coat reveals diverged functions of γζ-COP-Arf1 and βδ- COP-Arf1" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Randy Schekman as the Senior Editor. The following individual involved in review of your submission has agreed to reveal his identity: Benjamin D Engel (Reviewer #3).
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
The authors have used state-of-the-art cryo-EM tomography, combined with X-ray crystallography and photo-crosslinking, to determine the structure of the COPI coat at 9Å resolution. The structure provides many mechanistic insights, not only into the workings of the COPI coat, but also into how the related AP complex-containing coats might function, and into how the whole family may have evolved. It will be of interest to both structural and cell biologists – including evolutionary cell biologists.
All three reviewers acknowledge the excellence and importance of the work. Although a few additional experiments are suggested, in every case the authors could address the reviewers' comments by making modifications to the text (e.g. by softening interpretations), unless they have the data already in hand. Many of the reviewers' suggestions are about ways to make an inherently complicated structure as clear as possible to the reader. These are constructive comments that the authors should take on board, so that the paper can be understood by as many readers as possible. Other comments include concerns about a few discrepancies, and about the speculative nature of their model, which the authors should easily be able to address in the text. The reviewers have done such a thorough job that instead of trying to paraphrase their comments, I am appending their complete reviews below.
Vesicle coat complexes enable vesicular traffic between different compartments of the endomembrane system. They are highly dynamic macromolecular complexes that reversibly associate with membranes, select protein and lipid cargo, orchestrate vesicle budding and scission from the donor compartment, and – via the interaction with tethering complexes – contribute to the selection of the acceptor compartment. All these different steps must be properly coordinated and proofread to ensure smooth operation of vesicular traffic, which is a fundamental cellular process. Structural insight into some vesicle coat complexes such as clathrin or COPII is very far advanced whereas the structural analysis of COPI and its conformational dynamics has lagged behind for many years.
The manuscript by Dodonova et al., is an impressive illustration of how the combination of cutting edge cryo-electron tomography methods with X-ray crystallography and elegant functional experiments can bring about a major leap in understanding. This work intrigues and fascinates by providing many deep insights into the mechanics of the COPI coat.
Specifically, the following major advances are presented:
The manuscript finally closes a gap by providing a higher-resolution structure of β-δ-COP. As evident from the previous ET paper by Dodonova et al., such crystal structures are important to validate ET structures and to harness their full potential. Homology models of COPI subunits based on clathrin adaptor structures were of limited use in this context and the data presented in this paper elevates the resulting structural model of the entire COPI complex on the membrane to a stunning level of detail and quality.
Next, the manuscript makes substantial progress in understanding the structural integration of the appendage domains of β- and γ-COP respectively. The insights provided into their different accessibility and packing with other subunits in the case of the β-COP appendage are truly striking and will be key to a full functional understanding of the COPI vesicle formation.
Finally, the manuscript is of fundamental importance because it provides detailed insights into COPI's functional interplay with the small GTPase Arf1. This interaction is crucial for COPI recruitment to the membrane, coat formation, and uncoating. The data reveal two structurally and functionally distinct modes by which the small GTPase contacts COPI. This functional angle of the manuscript is developed a lot further by presenting yet another structure including the catalytic domain ArfGAP2.
Given the array of novel and exciting findings one aspect is treated with surprising lack of depth and rigorousness: The issue of the linkages in the COPI coat. The authors start the short description of the pertinent data and their interpretation by saying "We determined the structure of the linkages between triads as previously described (Dodonova, 2015)", suggesting that the following sentences logically build on the previous paper. But they don't. In the current model other subunits are delineated as being important for the linkages between triads. Also, work demonstrating that yeast remains viable without epsilon-COP and the mu-homology domain (Arakel et al., 2016) is not discussed in this context. It is clearly communicated that the resolution of the structure is lower in the relevant regions but a comparison between the previous paper by Dodonova et al., and their current data is warranted. A more conceptual discussion of why the COPI coat may be so different compared to COPII and clathrin in terms of the regularity of its structure would be very useful to the field.
A control experiment for Panel 6A would be important. We agree with the assignment of the specific cross-linked products in the anti-δ-COP IB. However, in the anti-Arf1 IB on the left, the difference between lanes 4 and 8 is very subtle, indeed. It would be an important corroboration of this experiment if a mutant form of δ-COP where the hydrophobic side chains of the helix are mutated were tested as a negative control.
This manuscript by Dodonova et al., presents the highest resolution structure to date of the COPI coat by fitting a new 2.6 Å crystal structure of β-δ-COP, existing atomic structures of other COP proteins, and homology models of the remaining components into a new 9 Å subtomogram average of the membrane-bound COPI coat reconstituted on membranes in vitro. The authors then build upon this structure with a subtomogram average of COPI bound to ArfGAP2, which binds only one of the two Arf1 sites, providing insights into the mechanisms of COPI coat assembly and disassembly.
While these findings are somewhat incremental, they are important and greatly increase our structural understanding of COPI function. Moreover, the work is technically excellent; a sub-nanometer subtomogram average of a relatively flexible and modular complex is certainly at the leading edge of what is possible with current technology. The jump in resolution from 13 Å (Dodonova et al., Science 2015) to 9 Å (this study) is biologically significant because it allows the secondary structure of the assembled COPI triad to be resolved and fairly accurately modeled for the first time.
I have no critical concerns that would preclude the publication of this study. The structures presented here are of high quality and will be very useful to the scientific community. However, the issues below must be addressed:
1) I do not believe the data in this paper supports the model proposed by the authors for the recruitment of coatomer by Arf1: "These observations suggest a mechanistic model for the role of δ-COP helix b in COPI function. Upon binding of the Arf1 N0 amphipathic helix to the membrane, the binding site for δ-COP is exposed, and the resulting interaction contributes directly to Arf1-dependent coatomer recruitment to the membrane."
The authors state in the introduction that Arf1-GDP is recruited to the membrane, where it is activated into Arf1-GTP by a GEF protein. Then Arf1-GTP recruits the coatomer via an interaction with β-δ-COP. The low binding affinity of Arf1-GDP for β-δ-COP is confirmed by the pulldown assays in Figure 6D to E, conflicting with the author's model that Arf1-GDP recruits the coatomer. I agree that the amphiphatic NO helix of Arf1-GDP (which is absent from Arf1-GTP) likely helps recruit Arf1-GDP to the membrane. However, the less conflicting model to propose would be that a GEF protein then converts it into Arf1-GTP, which removes the NO helix, enabling the "b" helix of δ-COP to bind in the site that was previously occupied by the NO helix. Interestingly, while losing the amphiphatic N0 helix could weaken the association of Arf1-GTP with the membrane, the "b" helix of δ-COP is also amphiphatic and so may be able to compensate and stabilize the membrane interaction (although the authors state that the nearest hydrophobic residues of the "b" helix are too far from the membrane, the "b" helix does occupy the same location as the NO helix).
If the authors want to propose a model where membrane binding by Arf1-GDP increases the affinity of Arf1-GDP for δ-COP, thus recruiting coatomer to the membrane prior to GEF-mediated conversion to Arf1-GTP, then they must do this experiment or provide strong supporting references.
2a) The mouse sequences for β-COP and δ-COP (mouse proteins were used for the EM structure) are 953 and 511 amino acids long, respectively, according to NCBI and Figure 5A. However, the structure presented in Figure 1 as the "crystal structure of the β-δ-COP subcomplex" only contains β-COP 19-391 and δ-COP 1-175 from C. thermophilium (the COP proteins are highly conserved, so mixing organisms in the model should be fine). This is only 39% of β-COP and 34% of δ-COP. While this is not a problem in itself, the title of the Results section and the title of the figure legend should state that this is a "partial structure of β-δ-COP" or a "structure of the interaction between β-COP and δ-COP". In order to clearly convey which regions of the proteins are included in the structure, an inset should be added to Figure 1 that diagrams the domains of β-COP and δ-COP (similar to Figure 5A), illustrating which parts are in the crystal structure and which parts are not.
2b) Figure 5 returns to the structure of β-δ-COP, this time looking at the subtomogram average to show the interaction with Arf1. Compared to the crystal structure in Figure 1, more (all?) of β-COP is displayed. However, the C-terminal half of δ-COP, corresponding to the MHD domain, is still absent. Later in the manuscript, it is briefly mentioned that the δ-COP MHD domain is part of the linkage between COPI triads (along with epsilon-COP and the C-terminus of α-COP; Figure 6—figure supplement 2, and crucially, only in the figure legend of Figure 2—figure supplement 2 is it mentioned that "δ-COP MHD is located in the linkage, and not in the COPI leaf, so it is not enclosed by the EM density". Finally, I understood why the MHD domain is not pictured in Figure 5, but this took far too much detective work. Thus: 1) the legend for Figure 2C should state which COP domains are not resolved within the 9A leaf structure due to their location in the linkage region outside of the triad, and 2) the legend of Figure 5 should reiterate that the MHC domain is not shown because it is outside the leaf structure.
3) Related to the above issue, additional labels should be added to many of the figures in order to make it easier to understand which components are being displayed in the images without having to continually return to the Figure 2 legend for the color code. If only specific domains are being shown instead of full proteins, this should also be labeled. This will make the results much easier to follow for readers who are not intimately familiar with the COPI structure. Specifically:
a) Color-coded labels on figure panels adjacent to the corresponding subunits:
Figure 1: dark green "β-COP" (in addition to existing labels); Figure 3A: red "appendage sandwich subdomain", purple: "appendage platform subdomain"; Figure 3B and C: black "γ-COP appendage", light blue "β'-COP"; Figures 3D and E: black "β-COP appendage", dark blue "α-COP"; Figure 4B: light blue "β'-COP", dark blue "α-COP"; Figure 4C: dark blue "α-COP N-terminus"; Figure 4D: dark green "β-COP", orange "δ-COP", light green "γ-COP", yellow "zeta-COP", pink "Arf1"; Figure 5B: dark green "β-COP", black "Longin domain" (in addition to "a", "b", "c"); Figure 7C: red "ArfGAP2", pink "Arf1".
b) The degree of rotation (e.g. 90°) should be added next to the rotation symbols in each of the Figure 2A,B, 3C,E, 4A,B, 5B, Figure 2—figure supplement 2 and 2D Figure 7—figure supplement 1. Additional rotation symbols and degrees should be added to Figures 4C and D.
c) In Figure 5D, it would be very helpful to illustrate the position of the δ-COP "b" helix from the cryo-EM structure relative to Arf1-GDP in order to show how the N0 amphiphatic helix of Arf1-GDP blocks the binding site for δ-COP on Arf1. Since there is space, this could either be incorporated into the current image or be shown in a second image.
d) Figure 6C would be much more clear if the "a" and "b" helices of δ-COP were colored red and yellow, respectively, to be consistent with their coloration in Figures 1 and 5, and Figure 1—figure supplement 1 Upon doing so, the yellow spheres representing crosslinked residues should be recolored to match the red "b" helix (perhaps a dark orange would look good).
4) No PDB structures are listed for the atomic models that were fit into the EM density (α-COP, Arf1-GTP, etc.). While these structures may have been named in the previous Science paper, at least in the methods of the current manuscript, these structures should be listed and the appropriate papers should be cited.
5) As in situ structures within cells are now possible, it should be made more explicitly clear in the abstract, introduction, and summary that the structure being presented is from an in vitro reconstituted system. Specifically:
Abstract: change "we determined the structure of the COPI coat assembled on membranes" to something like "we determined the structure of the COPI coat assembled on membranes in vitro".
Introduction: change "we recently determined the architecture of the COPI coat assembled on the vesicle membrane" to something like "we recently determined the architecture of the in vitro reconstituted COPI coat assembled on vesicle membranes".
Summary: change "by combining the β-δ-COP crystal structure and the EM structure of the COPI coat on the vesicle membrane" to something like "by combining the β-δ-COP crystal structure and the in vitro EM structure of the COPI coat on the vesicle membrane".
Dodonova and colleagues have determined a new crystal structure of part of the β/δ-COP subcomplex, which revealed several interesting new features in the 'linker region' of δ-COP (between the longin domain and μ-homology domains). They use this crystal structure, together with published X-ray structures, homology models, and their cryo-ET data, to generate an improved model of the mammalian COPI coat at ~9Å resolution using vesicles formed in vitro. The improved resolution allows them to locate and place key COPI subunits within coats assembled on membranes, including the β and γ appendage domains, and important helices in δ- COP. Photo cross-linking data support their structural model that places Arf1 in close proximity to β/δ-COP and specifically near to the δ-COP helix b. The authors also present structural evidence that Arf1 molecules in the coat are not equivalent and that only specific Arfs (bound to γ-COP) would be accessible for ArfGAP2 binding.
Overall, the paper and associated movie presents beautiful new crystallographic and EM data that provide insight into the overall structure of the COPI coat. But the paper currently has significant weaknesses. It is not written especially well: it would 'read' much better if the authors clearly separated results from discussion and speculation, and several important discussion aspects seem to be missing (e.g. a more complete comparison of COPI versus AP2 appendage domains) or glossed over. The figures could be significantly improved in terms of color selection, consistency, and labelling. If you're not already very familiar with these structures, you will likely struggle to understand this story. There is a great deal of speculation here.
There are two weaknesses in this story. Many questions remain around the biochemical data, which is currently insufficient to demonstrate how δ-COP helices interact with Arf1. This seems to be a key interaction in the coat that unites membrane recruitment, cargo recognition, and coat assembly. The pulldown data also do not seem to support placement of δ-COP helixC in the electron density.
Secondly, they emphasize the possibility and importance of non-equivalent Arfs in the coat but provide no biochemical or in vitro data to support this. The structure may imply two different functions, but they have certainly not shown it. One would prefer to see specific structure-based point mutants in the β/δ and/or γ/ζ sub-complexes (or perhaps just in the δ-COP linker?) used in liposome budding assays. One might predict mutations in the δ-COP linker alone or in the β/δ-COP interface would result in inefficient coat recruitment or budding, while mutations in γ/ζ subcomplexes would not. Finally, coat disassembly is only one model for ArfGAP function; yeast data suggest a role in coat assembly too. The authors should comment on based on their new structure.
1. δ-COP helices section. Does it make sense for helixC to be assigned to this position in the density? It seems located far away: what's the distance the δ-COP unstructured linker must span? And where does that imply the MHD sits? Why is helix c needed to help coordinate the MHD? This is very speculative, and the biochemical data don't seem to support the placement of helixC (below).
2) The authors need to describe more clearly experimental details for pulldowns between β/δ-COP and Arf1 (GMPPNP) in the main text and figure labels (ie. strep-tagged sub-complexes on sepharose). Importantly, why don't we see two Coomassie bands in all lanes of Figure 6D, corresponding to β- and δ-COP? And why haven't the authors shown pulldowns with both GDP- and GTP-locked forms of Arf1 if they say the interaction is nucleotide 'agnostic'?
Although the pulldowns demonstrate a biochemical interaction between purified β-/δ-COP and Arf1, such an interaction has been shown previously by Jonathan Goldberg. What's new here seems to be the importance of the linker region for the interaction. The authors state the main interaction partner of δ-COP helix b is Arf1, but their pulldown data indicate helix C is important (Figure 6D and E). But then they place helixC very far away from Arf1 in the density (Figure 6B). Why not do pulldowns with δ-COP linkers (of different lengths) and Arf1 in both nucleotide states to nail down this interaction? Is the affinity too low, even by Western blotting? Either way, the biochemistry suggests helixC is located much closer to Arf1. This story is interesting but is 'left hanging'.
3) The authors claim δ-COP helix b makes interactions with specific Arf1 regions (Switch I/Interswitch). I'm not convinced the resolution is sufficient to make these claims. And they have not shown us biochemical data to support the idea that δ-COP is agnostic to the nucleotide state of Arf1.
4) The authors propose that only the Arfs near γ-COP would be hydrolyzed by ArfGAP2? They suggest only these γArfs would be accessible for ArfGAP2 binding while βArfs would facilitate interaction with ArfGAP1. Is there evidence somewhere that supports the idea that different ArfGAPS bind different coatomer subunits (yeast two hybrid etc)? And is there any relation at all between one of these Arfs and the so-called 'backside' Arf1 seen in Hurley's open AP1 structure?
5) The idea of non-equivalent Arfs could be tested biochemically by combining structure-based mutagenesis and their liposome budding assay to count budding vesicles per area. This would be an important confirmation of their structure-based hypothesis.https://doi.org/10.7554/eLife.26691.068
- Felix Wieland
- John AG Briggs
- Irmgard Sinning
- Felix Wieland
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
EM maps and corresponding structural models are deposited in the Electron Microscopy Data Bank (accession codes EMD-3720, EMD-3721, EMD-3722, EMD-3723, EMD-3724) and the Protein Data Bank (accession codes PDB: 5NZR, 5NZS, 5NZT, 5NZU, 5NZV). Coordinates and structure factors for the X-ray structure of βδ-COP are deposited in PDB under accession code 5MU7. We thank P Schultz for the vectors used for site directed photolabeling, K Bacia for providing S. cerevisiae Arf1 protein; A. von Appen and J Kosinski for help with the MS data; F Schur, W Wan, O Avinoam, S Mattei, Y Bykov for discussions. We are grateful to J Goldberg for providing the coordinates of the Arf1-ArfGAP1 model. We thank C Siegmann from the BZH/Cluster of Excellence:CellNetworks crystallization platform for support in protein crystallization. We thank the staff of ESRF and of EMBL-Grenoble for assistance and support in using beamlines ID-21 and ID-29. Our work was technically supported by EMBL IT services, and was funded by the Deutsche Forschungsgemeinschaft within SFB638 (A16) to JAGB and FW, and SFB638 (Z4) to IS; and WI 654/12–1 to FW. FW and IS are investigators of the Cluster of Excellence:CellNetworks.
- Margaret S Robinson, Reviewing Editor, University of Cambridge, United Kingdom
© 2017, Dodonova et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.