SUMMARY
Intraflagellar transport (IFT) is the highly conserved process by which proteins are transported along ciliary microtubules by a train-like polymeric assembly of IFT-A and IFT-B complexes. IFT-A is sandwiched between IFT-B and the ciliary membrane, consistent with its putative role in transporting transmembrane and membrane-associated cargoes. Here, we have used single-particle analysis electron cryomicroscopy (cryo-EM) to determine structures of native IFT-A complexes. We show that subcomplex rearrangements enable IFT-A to polymerize laterally on anterograde IFT trains, revealing a cooperative assembly mechanism. Surprisingly, we discover that binding of IFT-A to IFT-B shields the preferred lipid-binding interface from the ciliary membrane but orients an interconnected network of β-propeller domains with the capacity to accommodate diverse cargoes towards the ciliary membrane. This work provides a mechanistic basis for understanding IFT-train assembly and cargo interactions.
Graphical Abstract
INTRODUCTION
Cilia, or flagella, are protrusions of eukaryotic cells with pleotropic functions that span from cellular motility to the sensation of physical, chemical and protein signals1. Ciliogenesis, cilia homeostasis, and the establishment of cilia signaling pathways rely on a dedicated and highly conserved trafficking mechanism called intraflagellar transport (IFT) which shuttles proteins in, out, and within cilia2. Consistent with the important role of IFT in most cilia functions, defective IFT is associated with a wide range of genetically heterogeneous human diseases including skeletal ciliopathies, polycystic kidney disease, retinal degeneration, and Bardet-Biedl syndrome (BBS)3.
Anterograde IFT toward the ciliary tip is powered by kinesin-24, whereas retrograde transport is driven by dynein-25. These two ATP-consuming motors travel processively along the outer microtubules of the axoneme at velocities between 2 to 4 μm/s4 while avoiding collisions6. The majority of IFT cargoes are coupled indirectly to the motors by IFT-A, IFT-B and the BBSome, which are multisubunit complexes of 6, 16, and 8 subunits, respectively. IFT-B is implicated in trafficking soluble proteins including tubulin7,8 and precursors of axonemal complexes9–11, whereas the BBSome and IFT-A are thought to transport transmembrane and membrane-associated proteins12–15. Cargoes are thought to be recognized by the IFT complexes by both direct interactions7,16 and through adaptor molecules10,13.
In the cilium and during assembly at the ciliary base, the IFT complexes assemble with their motors into polymeric assemblies known as “trains”4,17,18. Anterograde trains are compact structures approximately 200–500 nm in length with clearly defined periodicities, whereas retrograde trains have a less dense zig-zag pattern with a longer repeat length6. Subtomogram averaging has revealed repeating copies of IFT-B form the backbone of the anterograde train and make extensive contacts with dynein-2, which is carried as an inactive passenger19. IFT-A is situated between IFT-B and the ciliary membrane, where it presumably contacts its transmembrane cargoes. Although IFT-A is substoichiometric relative to IFT-B, it always occurs in linear arrays18,19.
Individual IFT proteins, including 4 IFT-A subunits, are rich in N-terminal β-propeller domains and C-terminal tetratricopeptide repeats (TPRs). Given that similar domain organizations occur in the subunits of vesicle coatomers (COPI, COPII and clathrin) and tethering complexes (CORVET, HOPs), it has been proposed that they share an evolutionary relationship20,21. Mounting evidence supports an extraciliary role for IFT proteins in vesicular trafficking, including localization of IFT46 to the surface of periciliary vesicles by in-situ immunogold labeling22 and the loss of densely coated periciliary vesicles in mouse cells deficient in the IFT-A subunit, IFT12123. How IFT-A binds vesicles, polymerizes into trains, and transitions between these states, are unanswered questions of fundamental importance toward a mechanistic understanding of IFT.
Unlike IFT-B7,24–30, dynein-231 and the BBSome32–34, structural data for the IFT-A complex is limited. Here, we have used single-particle cryo-EM to determine 3–4 Å resolution structures of native IFT-A complexes. In combination with spatial information from IFT trains in situ and by reconstituting lipid binding in vitro, we reveal the architecture of IFT-A, the mechanism by which IFT-A polymerizes into trains, and the surfaces capable of binding cargoes and membranes.
RESULTS
For a source of native IFT-A, we used Leishmania tarentolae, a single-celled uniflagellate of the Kinetoplastida order. Although L. tarentolae is not pathogenic to humans, it is closely related to organisms responsible for three major human diseases: leishmaniasis (various Leishmania species), African sleeping sickness (Trypanosoma brucei), and Chagas disease (Trypanosoma cruzi). Their flagella are crucial for their motility, pathogenicity, and viability35. For these reasons and their amenability to reverse genetics, the Kinetoplastida order has long been used for studies of flagella biology. Microscopy studies have demonstrated that the IFT machinery is conserved between Kinteoplastids and Chlamydomonas: both T. brucei and C. reinhardtii form anterograde trains of similar length (from ~200 nm to over 500 nm)18,36,37 and velocity (2–2.5 μm/s)36–39. Thin-section transmission electron microscopy revealed IFT trains in L. tarentolae flagella (Fig. S1A).
To purify the IFT-A complex from L. tarentolae, we created a strain expressing FLAG-tagged IFT43, the smallest of the IFT-A subunits. Anti-FLAG affinity purification recovered not only all six IFT-A subunits (IFT43, 121, 122, 139, 140 and 144), but most IFT-B subunits (Table S1A) indicating that IFT-A and -B coprecipitate. To simultaneously improve the purity and stability of the IFT-A complex, we centrifuged the sample through a sucrose gradient containing the chemical crosslinker, DTSSP (3,3’-Dithiobis (sulfosuccinimidylpropionate)). Fractions containing IFT-A were pooled from the gradient and purified further using ion-exchange chromatography (Fig. S1C). SDS-PAGE analysis (Fig. S1D) and mass spectrometry (Table S1B) revealed that the purified sample was almost exclusively IFT-A and that the interaction with IFT-B did not survive the purification process even in the presence of the crossslinker. Two-dimensional classification of particles from negative-stain electron microscopy revealed that the sample contained three major populations (Fig. 1A). One population was an elongated particle, approximately 300 Å in length and formed by two subcomplexes, one with a characteristic V shape. An almost identical class was obtained for IFT-A purified without DTSSP demonstrating that this state is not induced by the crosslinker (Fig. S2A). The other two populations were of these subcomplexes alone. Intriguingly, we did not observe IFT-A polymers in vitro, suggesting that polymerization of IFT-A is dependent on the prior oligomerization of IFT-B. This finding is consistent with the order of train assembly observed in vivo18.
Structure of a monomeric IFT-A complex
To gain a better understanding of the architecture of IFT-A, we used cryo-EM single-particle analysis. Cryo-EM revealed two major populations of intact IFT-A that differed in the orientation of a large α-solenoid protein (IFT139) at one end of the complex (Fig. 1B). The structures of these populations were resolved separately to overall resolutions of 4.0 Å (class 1) and 3.6 Å (class 2). Because class 2 had more than twice the number of particles of class 1 and higher resolution, we focused our analysis on this conformational state. Even after separating the large-scale movement of IFT139, other parts of the complex showed a high degree of lability. By masking, and then combining, specific regions during processing we generated a composite map with improved local resolution (Figs. S1 and S3). The resolved sidechains of the map (Fig. S3C) allowed unambiguous modeling of all six subunits. High-resolution maps were necessary to confidently distinguish between the similar domain architectures of the IFT121/122/140/144 subunits (Fig. 1C). The final atomic model (Fig. 1D) is consistent with chemical crosslinking mass spectrometry (XL-MS) data obtained for Tetrahymena thermophila IFT-A40 (Fig. S4). The few outliers, with distances beyond the crosslinking capabilities of disuccinimidyl sulfoxide41, mostly involve residues within the TPR domains of IFT140 and IFT144, which, as we show later, are capable of large-scale movements37
Our structure revised a prior description of IFT-A as having core and peripheral subcomplexes13,42. We found that the IFT-A “core” (IFT122, 140 and 144) actually corresponded to the V-shaped subcomplex and that the “peripheral” subcomplex of IFT43, 121 and 139 also contained IFT122 as an integral component (Fig. 1D). Given that neither subcomplex is peripheral to the other, we redefine the core subcomplex as IFT-A1 and the peripheral subcomplex as IFT-A2, reflecting a similar nomenclature used for IFT-B subcomplexes28.
Both subcomplexes have distinctive architectures. The V-shape of the A1 subcomplex is generated by a heterodimer of IFT140/144 in which their tandem β-propeller domains are splayed ~50 Å apart. IFT121/122 form a similar heterodimer in the A2 subcomplex but with their β-propeller domains closed together into a tetrameric arrangement. The heterodimerization of IFT121/122 and IFT140/144 occurs through a common dimerization module that involves an antiparallel interaction between the first 5 helices of their respective TPR domains (Fig. 2A,B). This TPR–TPR interaction is different from how subunits with similar domain architectures heterodimerize in COPI coatamers43 and the HOPs complex44.
IFT139, at the bottom of the A2 complex, interacts with IFT122β-prop2 and the C-terminal TPRs of IFT121 (Fig. 2C). The interaction with IFT122β-prop2 involves the insertion of the positively charged sidechain of the penultimate residue of IFT139, R1642, into the center of the β-propeller (Fig. 2D). This interaction is likely highly conserved, as a basic residue occurs at or near the end of most IFT139 sequences (Fig. 2E). The interaction between IFT139 and IFT121TPR is stabilized by IFT43, which contributes short α-helices to the interface (Fig. 2C). The mediatory position of IFT43 explains why Chlamydomonas ift43 mutants have barely detectable levels of IFT13945. In humans, a mutation (W174R) within the IFT121–IFT43 interface (Fig. 2F) is associated with a skeletal ciliopathy46, demonstrating the importance of this interaction for proper IFT-A integrity and function. Notably, only the conserved region of IFT43 is resolved in our maps (Fig. 2G). The N-terminus, which is dispensable for ciliogenesis45, is predicted to be unstructured.
The subcomplexes are connected by IFT122TPR, which extends from the A2 subcomplex to engage with the TPR domains of IFT140 and IFT144 at the apex of the V-shaped A1 subcomplex (Fig. 2H). This interaction explains why IFT122 can form a stable complex with IFT140/144 in a Chlamydomonas mutant lacking IFT12142 and in human cell lines overexpressing fluorescently tagged IFT subunits47. To address whether IFT122 would segregate with either the A1 or the A2 subcomplex in their separated forms, we resolved a 3.8-Å resolution structure of the A2 subcomplex alone, which revealed IFT122 together with IFT43/121/139 (Fig. 1C and Fig. S1–2). In this subcomplex, IFT122TPR is not resolved, indicating that the connecting bridge is flexible in the absence of the A1 subcomplex. The isolated A1 subcomplex therefore likely consists of just IFT140/144, consistent with their ability to co-immunoprecipitate in Chlamydomonas ift122 mutants42.
In addition to IFT122TPR, the A1 and A2 subcomplexes are associated by an inter-propeller interaction between IFT140β-prop1 and a conserved surface of IFT121β-prop1 (Fig. 2I). Another, more labile connection (based on the relative weakness of the density) involves the C-terminus of IFT144, which extends from the A1 subcomplex to loosely contact IFT121TPR in the A2 subcomplex (Fig. 1D). As described later, both these interactions change when IFT-A polymerizes into anterograde trains. The C-terminus of IFT140TPR has no discernable density, indicative of extreme flexibility.
IFT121, IFT122 and IFT144 all terminate with two Cys4 zinc-finger (ZnF) domains, suggesting they share a common evolutionary origin (Fig. S5). These ZnF domains are highly conserved, although not all species have two ZnF domains per subunit; human IFT121, for example, lacks the first ZnF domain (Fig. S5A). In IFT122, the ZnF domains are separated by a small mixed α/β domain of unknown function (Fig. 2H and Fig. S5B). Loss of a Zn-coordinating cysteine (C1267Y) in human IFT144 is associated with short-rib thoracic dysplasia48, demonstrating that correct folding of the ZnF domain is required for proper IFT-A function. Proteins with similar WD40–TPR architectures were also found to terminate in ZnFs in the membrane-binding GATOR2 complex49 where they mediate heterodimerization. There is no evidence from our structure to support a similar dimerization function for the ZnF domains of IFT-A.
An atomic model of IFT-A polymerized on an anterograde IFT train
To determine how IFT-A polymerizes during IFT, we docked our atomic model into subtomogram averages of IFT-A from anterograde trains19,40, the best of which has a resolution of 23 Å40 (Fig. 3). The A2 subcomplex can be fitted unambiguously due to its distinctive arrangement of 4 β-propellers (two each from IFT121 and IFT122) and a large α-solenoid (IFT139) (Fig. 3B–C). Unexpectedly, the A1 subcomplex shows a substantial conformational rearrangement as a result of polymerization (Fig. 3D). It has rotated relative to A2, breaking the inter-propeller contacts between IFT121β-prop1 and IFT140β-prop1 (Video S1). The TPR domain of IFT122 straightens, and potentially rigidifies based on the strength of the density, in response to the rotation (Fig. 3D, inset). The liberated IFT140 rotates to contact IFT121β-prop2 of its distal neighbor. Despite this movement, the distance between the splayed β-propeller domains of IFT140/144 remains largely unchanged (~50 Å), consistent with a rigid-body motion. By generating lateral contacts between neighboring molecules, the swiveling of the A1 domain is a crucial step in IFT-A polymerization.
We also observed large changes in the positions of the TPR domains of IFT140 and IFT144 on the underside of the IFT-A complex. The C-terminus of IFT144TPR is released from its loosely bound position on IFT121TPR to engage IFT121β-prop2 of its distal neighbor (Fig. 3D–E). This appears to induce ordering of the C-terminal region of IFT140TPR, which curls to interact with IFT144TPR in its new position (Fig. 3E). It should be noted that these positions are tentative as accurate modeling of the TPRs was challenging due to the 23 Å resolution of the subtomogram average. The position of IFT144 in the train may explain some of the outlier crosslinks that were incompatible with the model of monomeric IFT-A (Fig. S4). TPR–TPR interactions also occur between IFT139 and IFT121 from neighboring complexes. In order for these interactions to occur, IFT139 has rotated further from its position in either state 1 or state 2. The need for IFT139 to be dynamic may explain why we observed multiple states. In conclusion, collective cooperativity provided by multiple lateral interactions explain why IFT-A polymerizes in linear arrays on IFT-B rather than binds sporadically18.
To visualize how these IFT-A polymers are oriented on IFT trains, we next docked our atomic model into a composite map of an IFT train assembled at the ciliary base prior to anterograde transport (EMD-15261)18 (Fig. 3F). This revealed that IFT-A adopts a slanted configuration on the train relative to IFT-B with a footprint covering approximately 3 IFT-B complexes. IFT-A is tethered to IFT-B through unassigned densities either side of the IFT139 subunit (Fig. 3G). Candidate IFT-B subunits that might mediate these contacts include IFT70, IFT88, and IFT172 which crosslink with IFT-A40, and IFT74/81, which integrative modeling with AlphaFold2 predictions have proposed to be positioned near IFT-A29. Interestingly, these five proteins are the top five IFT-B subunits that copurify with L. tarentolae IFT-A (Table S1A).
The largest unassigned density is between the C-terminus of IFT144TPR and IFT139 (Fig. 3G). A crucial role for the C-terminus of IFT144 in binding IFT-B would agree with recent biochemical evidence from IFT144 truncations50,51. The C-terminus of IFT172 has been proposed to bridge IFT144 and IFT139 on anterograde trains29, potentially explaining this unassigned density. The link with IFT172 could explain why mutations in IFT17252,53 phenocopy those caused by IFT-A mutants (Table S3).
A consequence of the orientation of IFT-A on the IFT-B polymer is that all six β-propeller domains are on the upper face, closest to the ciliary membrane and all TPR domains face the IFT-B complex (Fig. 3F). The BBSome is also thought to position multiple β-propeller domains toward the ciliary membrane32,33. In the subtomogram average of anterograde IFT-A40, we observed additional density above the interface between IFT140β-prop1 and IFT140β-prop2 that cannot be explained by our atomic model (Fig. 3A). This density may correspond to a cargo or cargo adaptor that is present in vivo but absent from the purified IFT-A used for cryo-EM analysis. A role for the IFT140 β-propeller domains in cargo binding would be consistent with loss of ciliary localization of GTPases, lipid-anchored proteins, and cell signaling proteins in Chlamydomonas mutant strains lacking the IFT140 β-propeller domains54.
Orientation of IFT-A on membranes
The proposed evolutionary relationship between IFT-A and vesicle-binding coatomers and tethers20,21 and its membrane-proximal location on IFT trains19 support a direct interaction between IFT-A and membranes. Previous work with a recombinant Chlamydomonas IFT-A subcomplex (IFT43/121/139) had identified preferential binding to anionic phosphoglycerates23. To test if native L. tarentolae IFT-A has similar lipid specificity, we used a lipid-protein overlay assay (Fig. 4A). Of the 15 lipids tested, only phosphatidic acid (PA) and 3-sulfogalactosylceramide showed binding to IFT-A. Ceramide binding is consistent with the ability of ceramide to pull-down IFT complexes from Chlamydomonas flagella55. Because PA was also the dominant interaction detected for Chlamydomonas IFT43/121/139 subcomplexes23, we next tested whether IFT-A could bind PA-containing liposomes. Negative-stain electron microscopy showed partial decoration of the liposome surface but not the formation of cage-like coats (Fig. 4B). IFT-A did not decorate liposomes formed without PA (Fig. S6). Furthermore, two-dimensional class averaging revealed a consistent mode of attachment of IFT-A to the liposome surface, in which the curvature of the IFT-A complex complements the convex curvature of the liposome (Fig. 4C). By overlaying our atomic model onto these class averages (Fig. 4D), we could determine the approximate surfaces that face, and potentially interact, with the membrane. The closest contact is with the apex of the A1 module, which consists of the entwined TPR domains of IFT122, IFT140 and IFT144 (Fig. 2C). At the other end of the complex, IFT139 approaches the membrane in some but not all cases (Fig. 4B). A role for IFT139 in membrane binding would be consistent with the ability of IFT43/121/139 but not IFT43/121 to bind PA-containing liposomes and PA in lipid overlap assays23. Thus, the preferred membrane binding interface of the IFT-A complex involves its TPR domains. The ability of TPR domains to bind lipids is supported by a recent crystal structure of the TPR domain of a mitochondrial protein, PTPIP51, bound to PA56. Surprisingly, the same TPR surfaces face, and even engage with, IFT-B in anterograde trains rather than the ciliary membrane (Fig. 3F), rendering these two binding modes incompatible.
Ciliopathy-associated IFT-A variants reveal potential cargo-binding sites
A structure of IFT-A allowed us to analyze mutations that cause human ciliopathies. First, we exploited the structural conservation of the IFT-A complex and the predictive power of AlphaFold2 to build an atomic model of human IFT-A. Onto this model, we mapped all missense variants in IFT-A that are annotated as either pathogenic or likely pathogenic in the ClinVar database (Fig. 5A and Table S3). The mutations are dispersed throughout the structure with all six subunits harboring variants. Most mutations occur within the hydrophobic cores of individual subunits, where they may disrupt complex formation by instigating defective folding pathways. A smaller subset, including IFT43 W174R (Fig. 2F), map to intersubunit interfaces, where they could destabilize complex formation. We also observed 18 mutations occurring at surface-exposed sites (Fig. 5B). We speculate that mutation of residues that face the ciliary membrane and occur within conserved regions, such as IFT144 D159 (Fig. 5C), may elicit their deleterious effects by disrupting cargo binding rather than IFT-A complex formation.
DISCUSSION
Intraflagellar transport, essential for ciliogenesis and the signaling capabilities of cilia, requires the formation of megadalton polymeric assemblies that transport cargoes to and from the cilium. Here we have used cryo-EM to determine structures of native L. tarentolae IFT-A complexes. The structures revealed that IFT-A has a bilobal architecture of two subcomplexes, A1 and A2. Although the IFT-A assembly pathway remains to be elucidated, the structure supports a model where IFT121/122 and IFT140/144 independently heterodimerize through antiparallel TPR interactions, before recruiting each other and IFT43/139. Contemporary work reporting the cryo-EM structure of a human IFT-A complex, reconstituted from recombinant proteins, also shows a bilobal architecture57. However, the A1 and A2 subcomplexes are highly flexible relative to one another and are only connected through the IFT122TPR domain. No interaction between IFT140β-prop1 and IFT121β-prop1 was observed. This striking disparity in architecture and dynamics may reflect species-specific differences or a consequence of sample preparation methods (native versus recombinant). In negative-stain electron microscopy of L. tarentolae IFT-A purified without a crosslinker we observed the “closed” state and also less populated “open” states (Fig. S2A,B). A partially open state, in which the IFT140β-prop1 and IFT121β-prop1 are no longer in contact, was also observed as a minor class in our cryo-EM dataset of the crosslinked IFT-A sample (Fig. S2C). From these data, we conclude that native IFT-A exists as an equilibrium between closed the open states. The ability of the IFT140β-prop1 and IFT121β-prop1 interface to open is consistent with our mechanism for oligomerization (Fig. 3), in which this interface has to break to allow the IFT-A1 domain to swivel to contact its neighboring complex.
We show, using in-vitro reconstitution with liposomes, that the fully assembled complex can bind membranes through it TPR domains with specificity for phosphatidic acid and ceramides. Ceramides were shown recently to promote ciliogenesis, regulate retrograde IFT, and pull-down Chlamydomonas IFT subunits55. The ability of IFT-A to bind membranes provides further evidence for a shared evolutionary relationship with coatomers. Although we saw no evidence for IFT-A forming cage-like structures on liposomes, further work will be needed to discover if IFT-A can form ordered vesicle coats in cells.
Surprisingly, we discovered that IFT-A uses a common interface to engage both liposomes and IFT-B, with the N-terminal TPRs of IFT139 integral to both interactions. This contradicted our expectation that IFT-A would engage both membranes and IFT-B through opposing interfaces given that it is sandwiched between the two in anterograde trains. We hypothesize that our membrane-associated form, with lipid contacts mediated by TPR domains, corresponds to IFT-A on preciliary vesicles and/or retrograde trains (Fig. 4E). If our preciliary vesicle hypothesis is correct, competition between binding membranes or IFT-B at the ciliary base might explain how IFT-A transitions from a vesicle- or membrane-bound state to one that engages IFT-B on assembling trains in the cytosol. This is not to say that IFT-A in anterograde trains cannot bind membranes. Membrane binding could still be achieved through a lower affinity interface or through adaptors such as TULP3, which associate with IFT-A during anterograde transport13,57,58 and have affinity for phosphoinositides13.
If our retrograde train hypothesis is correct, it would require IFT-A to invert during the conversion from anterograde to retrograde trains at the ciliary tip (Fig. 4E). Rotation on a hinge provided by the flexible IFT139 subunit could allow inversion to occur without breaking contact with IFT-B, which is thought to be preserved during the transition59. A consequence of the inversion model is that it would physically break interactions with anterograde cargoes, releasing them from futile recycling back to the cytosol. By exposing a different interface to the ciliary membrane, the inverted IFT-A could recognize a different set of cargoes from those delivered to the cilium. A role for IFT-A in binding lipids specifically in retrograde trains may explain why ceramide biosynthesis inhibitors slow retrograde but not anterograde trains in Chlamydomonas flagella55. In-situ structures of IFT-A-coated periciliary vesicles and retrograde IFT trains will be needed to test these hypotheses. How and when IFT-A converts between monomeric, polymeric, and membrane-bound forms has fundamental implications for the assembly and conversion of IFT trains, and the pickup and release of ciliary cargoes.
Implications for cargo binding
Our structures and structural analysis suggest that both the outer edges and upper surface of the β-propeller domains may be involved in cargo recognition, and that the β-propeller domains of IFT140 and IFT144 in the IFT-A1 subcomplex may be particularly important. First, surface-exposed residues that are mutated in ciliopathies, and which may correspond to cargo binding sites, map to both the edges and upper surfaces of the β-propeller domains (Fig. 5). Second, density in the subtomogram average of an anterograde IFT-A train that cannot be explained by our model (Fig. 3A) is associated with the edges of the IFT140 β-propeller domains. This region includes G296, one of residues mutated in short-rib thoracic dysplasia (Table S3). Although we cannot identity the origin of this additional density, it indicates that factors may be able to bind this interface. Finally, a role for IFT140 in cargo binding is consistent with biochemical evidence showing a direct association between IFT140 and the ciliary targeting sequence of the ciliary G-protein coupled receptor, somatostatin receptor 3 (SSTR3)16.
A recent study has mapped the TULP3 binding site on human IFT-A to an acidic patch on the outside of IFT122 ZnF domain57. Using AlphaFold2 multimer, this region was predicted to bind the N-terminal helix of TULP3, which had previously been shown to be required for IFT-A binding13. Mutation of five conserved IFT122 residues within this interface, including charge reversal of three acidic residues, abolished TULP3 binding without disrupting the incorporation of IFT122 into IFT-A57. Binding to the IFT122 ZnF domain would place the phosphoinositide-binding TUBBY domain of TULP313 in the vicinity of the A1 subcomplex β-propeller domains on the end of a flexible tether. Our structure of the L. tarentolae IFT-A demonstrates that the putative binding site for the TULP3 N-terminal helix is conserved, including the acidic residues that are proposed to interact electrostatically with the basic residues of the helix. However, the L. tarentolae genome does not encode an obvious homolog of TULP3, and no TUBBY domain-containing proteins copurified with IFT-A. Whether L. tarentolae has an adaptor equivalent of TULP3 awaits investigation.
Limitations of the study
Here, we have shown that IFT-A displays lipid specificity and can partially coat liposomes in vitro, consistent with the in-vivo visualization of IFT121-dependent densely coated periciliary vesicles in mammalian cells23. However, further work will be needed to demonstrate exactly how IFT-A binds native vesicles, and whether it co-migrates with IFT-B during vesicular trafficking. Are they delivered to the ciliary base separately or as a preformed complex? How IFT-A transitions from its vesicle-bound form to the IFT train also needs further clarification. Does uncoating require membrane fusion, or is uncoating a prerequisite for membrane fusion? To aid these studies, our high-resolution structures will provide templates to identify IFT-A at different stages of trafficking in electron micrographs and tomograms60. While we show that lateral polymerization of IFT-A into IFT trains requires large subcomplex rearrangements, whether IFT-B polymerization follows similar principles will require structures of IFT-B before train formation. The ability to determine structures of native IFT complexes provides a viable avenue to achieve this aim. Similarly, although our structures reveal potential cargo-binding sites, the molecular basis for substrate recognition will require structures of co-complexes with membrane-protein cargoes and their adaptors.
STAR★Methods
RESOURCE AVAILABILITY
Lead contact
Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Alan Brown (alan_brown@hms.harvard.edu).
Materials availability
A pLEXSY-hyg2.1 plasmid expressing Flag-tagged IFT43 has been deposited with Addgene (https://www.addgene.org/) with ID #194433.
Data and Code Availability
Composite cryo-EM maps of L. tarentolae IFT-A state 1 and 2 have deposited in the Electron Microscopy Data Bank (EMDB) with accession numbers EMD-28866 and EMD-28867. Half maps and masks are deposited as additional maps associated with these entries. Atomic models of IFT-A state 1 and state 2 have been deposited in the Protein Data Bank (PDB) with accession numbers 8F5O and 8F5P.
This paper does not report original code.
Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.
EXPERIMENTAL MODEL AND SUBJECT DETAILS
Leishmania tarentolae
Leishmania tarentolae strain P10 (Jena Bioscience, #LT-101) cells were grown in BHI medium (HIMEDIA, #N210) containing 5 μg/mL hemin chloride (Sigma, #3741) and 10 U/mL Penicillin-Streptomycin (Gibco, #15070063) at 26°C in the dark. Cells were maintained as static suspension cultures as described in the LEXSY expression kit manual (Jena Bioscience).
METHOD DETAILS
Sample preparation for thin-section transmission electron microscopy
Cells in late logarithmic phase were diluted 1:10 and left overnight. The next day, the cells were fixed by a 5-min incubation with microscopy-grade glutaraldehyde at a final concentration of 2.5%. The cells were collected by centrifugation at 800 × g for 5 min. The pellet was resuspended in 1.5 mL fresh BHI medium without hemin and antibiotics. To this, an equal volume of fixation reagent containing 2% formaldehyde, 2.5% glutaraldehyde and 0.3% picric acid in 0.1 M sodium cacodylate buffer (pH 7.4) was added and left for 1 hr on a rocking platform. 200 μL of cell suspension (containing ~20–50 million cells) was then loaded onto 12.7 mm Aclar coverslips (Electron Microscopy Sciences, #50425–25) precoated with poly-L-lysine (Sigma-Aldrich, #P8920) and allowed to settle for 30 min. Excess liquid was aspirated, and the coverslip was spun at 4680 × g on a plate spinner for 3 min. The coverslip was then washed three times with 0.1 M sodium cacodylate buffer followed by fixation with 1% osmium tetroxide. After rinsing with 50 mM maleate buffer (pH 5.2), the sample was further incubated with 1% aqueous uranyl acetate for 1 hr at room temperature. The fixed sample was dehydrated by washing with escalating ethanol concentrations (70, 90 and 100%). Following dehydration, the sample was treated with propyleneoxide for 1 hr, infiltrated with epoxy resin and finally embedded in freshly mixed Epon. 50 nm sections were collected from the Epon blocks and mounted on the specimen grid. For contrast, the grid was stained with lead citrate and uranyl acetate. The images were collected on a 120 kV Tecnai T12 microscope (Thermo Fisher Scientific).
Construct design
To constitutively express IFT43-FLAG in L. tarentolae from a genome-integrated position, we modified expression vector pLEXSY-hyg2.1 (Jena Bioscience, #EGE-1310hyg) to insert a linker, 3X FLAG peptide sequence, and a XbaI restriction site into the KpnI site of the vector. The L. tarentolae gene encoding IFT43 was amplified from genomic DNA using primer oligonucleotides (Key Resources Table) and Phusion High-Fidelity DNA polymerase (Thermo Fisher Scientific, #F-530S). The amplified gene was inserted into the modified pLEXSY-hyg2.1 vector between the 5´ BglII or NcoI and 3´ XbaI sites. The ligated vector was transformed into E. coli DH5α cells and plated onto agar plates containing ampicillin. Bacterial colonies were screened by colony PCR using the gene-specific primers (Key Resources Table). The plasmids were purified and sequence verified before transfection.
REAGENT or RESOURCE | SOURCE | IDENTIFIER |
---|---|---|
Antibodies | ||
Rat anti-FLAG antibody | BioLegend | Cat #637301; RRID:AB_1134266 |
Horseradish peroxidase-conjugated anti-rat secondary antibody | Cell Signaling Technology | Cat #7077S |
Biological Samples | ||
Chemicals, Peptides, and Recombinant Proteins | ||
1,2-dipalmitoyl-sn-glycero-3-phosphate (PA) | Avanti Polar Lipids | Cat #830855P |
1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE) | Avanti Polar Lipids | Cat #850757 |
1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1’-rac-glycerol) (POPG( | Avanti Polar Lipids | Cat #840457 |
3,3’-dithiobis(sulfosuccinimidyl propionate) (DTSSP) | Thermo Fisher Scientific | Cat #21578 |
3X FLAG peptide | Pepmic | Custom synthesis, no identifier |
Anti-FLAG M2 resin | Sigma-Aldrich | Cat #A2220 |
BHI medium | HIMEDIA | Cat #N210 |
Hemin chloride | Sigma | Cat #3741 |
Penicillin-Streptomycin | Gibco | Cat #15070063 |
Poly-L-lysine | Sigma-Aldrich | Cat #P8920 |
Phusion High-Fidelity DNA polymerase | Thermo Fisher Scientific | Cat #F-530S |
Polycarbonate membrane | Avanti Polar Lipids | Cat #610005 |
Trypsin | Promega | Cat #90057 |
Critical Commercial Assays | ||
Novex ECL chemiluminescence kit | Thermo Fisher Scientific | Cat #WP20005 |
Membrane lipid strips | Echelon Bioscience | Cat #P-6002 |
Deposited Data | ||
Subtomogram average of C. reinhardtii IFT-A obtained from anterograde trains | McCafferty et al., 202240 | EMDB: EMD-26791 |
Subtomogram average of C. reinhardtii IFT-A obtained from anterograde trains | Jordan et al., 201819 | EMDB: EMD-4304 |
Composite subtomogram average of an assembled C. reinhardtii IFT train at the ciliary transition zone | van den Hoek et al., 202218 | EMDB: EMD-15261 |
Composite cryo-EM map of Leishmania tarentolae IFT-A state 1 | This paper | EMDB: EMD-28866 |
Composite cryo-EM map of Leishmania tarentolae IFT-A state 2 | This paper | EMDB: EMD-28867 |
Atomic model of Leishmania tarentolae IFT-A state 1 | This paper | PDB: PDB 8F5O |
Atomic model of Leishmania tarentolae IFT-A state 2 | This paper | PDB: PDB 8F5P |
Chemical crosslinking mass spectrometry data obtained for Tetrahymena thermophila IFT-A | McCafferty et al., 202240 | MassIVE/ProteomeXchange: PXD032818 |
Experimental Models: Cell Lines | ||
Experimental Models: Organisms/Strains | ||
Leishmania tarentolae strain P10 | Jena Bioscience | Cat #LT-101 |
Bacteria: Escherichia coli DH5α | New England Biolabs | Cat #C2987H |
Oligonucleotides | ||
LtIFT43_Fw1 CCTTGCCACCAGATCTGCCATGTCCACGACTTCTATGACTGCC |
Sigma | N/A |
LtIFT43_Rv1 CCGGAGCCTCTAGACTTCGAAACCTGCTTGGATG |
Sigma | N/A |
Recombinant DNA | ||
Plasmid: pLEXSY-hyg2.1 | Jena Bioscience | Cat #EGE-1310hyg |
Plasmid: pLEXSY-hyg2.1_LtIFT43_Flag | This paper | Addgene: Cat #194433 |
Software and Algorithms | ||
AlphaFold2 | Jumper et al., 202170 | https://www.deepmind.com/ |
ChimeraX v1.4 | Pettersen et al., 202169 | https://www.rbvi.ucsf.edu/chimerax/ |
Clustal Omega v1.2.2 | Sievers et al., 201174 | http://www.clustal.org/omega/ |
Coot v.0.9.8.3 | Brown et al., 201571 | https://www2.mrc-lmb.cam.ac.uk/personal/pemsley/coot/ |
cryoSPARC v3.3.2 | Punjani et al., 201766 | https://cryosparc.com/ |
DeepEMhancer | Sanchez-Garcia et al., 202162 | https://github.com/rsanchezgarc/deepEMhancer |
Prism v9 | Graphpad Software, USA | https://www.graphpad.com/ |
phenix.combine_focused_maps v1.18.2–3874 | Liebschner et al, 201968 | https://www.phenix-online.org/ |
phenix.molprobity v1.18.2–3874 | Chen et al., 201073 | https://www.phenix-online.org/ |
phenix.real_space_refine v1.18.2–3874 | Afonine et al., 201872 | https://www.phenix-online.org/ |
RELION 4.0 | Zivanov et al., 201867 | https://www3.mrc-lmb.cam.ac.uk/relion/ |
SBGrid | Morin et al., 201365 | https://sbgrid.org/ |
SerialEM v.3.8.5 | Schorb et al., 201964 | https://bio3d.colorado.edu/SerialEM/ |
SEQUEST | Thermo Fisher Scientific | https://www.thermofisher.com |
Other | ||
Aclar coverslips | Electron Microscopy Sciences | Cat #50425–25 |
Carbon-coated copper grids | Electron Microscopy Sciences | Cat #CF200-Cu |
R2/1, 200 mesh gold, QUANTIFOIL grids | Electron Microscopy Sciences | Cat #220210 |
Transfection
For transfection, L. tarentolae cell density was adjusted to 108 cells/mL, followed by incubation on ice for 10 min. 380 μL of this culture was mixed with 2 μg of linearized (SwaI digested and gel purified) plasmid. This mixture was immediately transferred to a pre-cooled electroporation cuvette (d=2 mm) and electroporated using a Gene Pulser Xcell Eukaryotic System (Bio-Rad) using time constant protocol at 450 V for 3.5 ms. The cuvette was then put back on ice for 10 min. The cells were allowed to recover overnight in fresh BHI-hemin medium as a static suspension culture. The cells were harvested by centrifugation at 2,000 × g for 3 min, resuspended in 200 μL and plated onto BHI agar containing 100 μg/mL hygromycin. The plate was incubated for 4–5 days until colonies were visible. Individual colonies were transferred to 150 μL BHI-hemin medium in a 96-well plate. For protein production, the cells were grown at scale as agitated suspension cultures.
IFT-A purification (with crosslinker)
L. tarentolae cells from 32 L culture volumes were harvested by centrifugation at 5,422 × g for 10 min. The cell pellets were resuspended in ~450 mL ice-cold lysis buffer (35 mM HEPES pH 7.4, 100 mM KCl, 5 mM MgSO4, 1 mM CaCl2, 1 mM DTT, Protease inhibitor cocktail, 20 mM NaF, 50 mM β-glycerophosphate, 1% Triton X100, and DNAse) and incubated at 4°C for 15 min with continuous stirring. The lysate was centrifuged at 42,510 × g for 30 min. The clarified lysate was applied to 4 mL anti-FLAG M2 resin (Sigma-Aldrich, #A2220) preequilibrated with wash buffer (35 mM HEPES pH 7.4, 100 mM KCl, 5 mM MgSO4, 1 mM CaCl2, 1 mM DTT). The column was washed with 200 mL wash buffer and eluted with wash buffer containing 100 μg/mL 3X FLAG peptide (Pepmic). The eluate was concentrated using a 15 mL concentrator with a 100 kDa cutoff Amicon Ultra filter (Millipore) to less than 1 mL and loaded onto a 13 mL 10–40% continuous sucrose gradient prepared in wash buffer with a gradient maker (Gradient Master, BioComp). Crosslinking reagent 3,3’-dithiobis(sulfosuccinimidyl propionate) (DTSSP; Thermo Fisher Scientific, #21578) was added to the 40% sucrose solution to a final concentration of 1 mM prior to making the gradient. The sample was centrifuged at 200,000 × g for 16 hrs using a SW40Ti rotor in an Optima LE-80K ultracentrifuge (Beckman Coutler) at 4°C. The gradient was fractionated into 600 μL aliquots with the crosslinking reaction quenched by adding 40 mM Tris pH 7.4. The fractions containing the complex were pooled and concentrated with a 4 mL Amicon Ultra concentrator with a 100 kDa cutoff (Millipore). The sample was loaded onto a MonoQ5/50 GL column (Cytiva, #17516601) preequilibrated with buffer A (20 mM HEPES pH 7.4, 5 mM MgSO4, 1 mM CaCl2, 50 mM KCl, and 1 mM DTT) and eluted using a gradient up to buffer B (20 mM HEPES pH 7.4, 5 mM MgSO4, 1 mM CaCl2, 1 M KCl, and 1 mM DTT). Peak fractions were analyzed by SDS-PAGE, mass spectrometry, and negative-stain electron microscopy.
IFT-A purification (without detergent or crosslinker)
Cell pellets were resuspended in ~300 mL ice-cold lysis buffer without detergent (35 mM HEPES pH 7.4, 100 mM KCl, 5 mM MgSO4, 1 mM CaCl2, 1 mM DTT, Protease inhibitor cocktail, 20 mM NaF, 50 mM β-glycerophosphate, 8% sucrose, and DNAse). The cells were lysed with a high-pressure homogenizer-EmulsiFlex-C3 (Avestin) with an applied pressure of 50–60 psi. The lysate was centrifuged at 42,510 × g for 30 min. The clarified lysate was applied to, and eluted from, a 4 mL anti-FLAG M2 resin as described above. The eluate was concentrated to ~0.5 mL and loaded onto Superpose 6 Increase 10/300 GL column (Cytiva, #29091596) preequilibrated in 20 mM HEPES pH 7.4, 100 mM KCl, 5 mM MgSO4, 1 mM CaCl2, and 1 mM DTT. Peak fractions were analyzed immediately by negative-stain electron microscopy.
Mass spectrometry
Mass spectrometry was performed at the Taplin Mass Spectrometry Facility at Harvard Medical School. In brief, the IFT-A samples were provided to the facility as dehydrated SDS-PAGE gel pieces. The gel pieces were rehydrated with 50 mM ammonium bicarbonate solution containing 12.5 ng/mL trypsin (Promega, Cat. #90057). After 45 min at 4°C, the trypsin solution was replaced with 50 mM ammonium bicarbonate solution and left at 37°C overnight. Peptides were extracted by removing the ammonium bicarbonate solution, followed by a wash with a solution containing 50% acetonitrile and 1% formic acid. The extracts were then dried using a vacuum concentrator for one hour and stored at 4°C. For mass spectrometry, the samples were reconstituted in 5–10 mL of solvent A (2.5% acetonitrile, 0.1% formic acid) and loaded onto a pre-equilibrated reverse-phase capillary column (100 mm inner diameter, and ~30 cm length) containing 2.6 mm C18 spherical silica beads using a Famos auto sampler (LC Packings). A gradient was formed, and peptides were eluted with increasing concentrations of solvent B (97.5% acetonitrile, 0.1% formic acid). As peptides eluted, they were subjected to electrospray ionization and entered into an LTQ Orbitrap Velos Pro ion-trap mass spectrometer (Thermo Fisher Scientific). Peptides were detected, isolated, and fragmented to produce a tandem mass spectrum of fragment ions for each peptide. Protein identity was determined using Sequest (Thermo Fisher Scientific). The data were filtered to between a one and two percent peptide false discovery rate.
Lipid-binding assays
Membrane lipid strips (Echelon Bioscience, #P-6002) were incubated with 5 mL Tris-buffered saline (TBS) containing 3% bovine serum albumin (BSA) overnight on an orbital shaker at 4°C. The next day, the buffer was renewed, and the strip incubated for another hour. The strip was then incubated with a 5 mL solution of ~0.05 mg/mL IFT-A in buffer containing 20 mM HEPES pH 7.4, 5 mM MgSO4, 1 mM CaCl2, 50 mM KCl, and 1 mM DTT at room temperature with constant shaking. After 2 hrs, the protein solution was discarded, and the strip washed twice (5 min per wash) with a total of 10 mL blocking buffer (TBS with 0.001% tween-20 (TBST) and 5% milk). The strip was then incubated with 5 mL blocking buffer containing a 1:500 dilution of anti-FLAG antibody (BioLegend, #637301) overnight with constant shaking at 4°C. Next, the strip was washed three times with 10 mL TBST for a total of 30 min before incubating with a horseradish peroxidase-conjugated anti-rat secondary antibody (Cell Signaling Technology, #7077S) for 1 hr at room temperature. Finally, the strip was washed three times with TBST and once with TBS (10 mL per wash, each for 10 min). Signal was detected using the Novex ECL chemiluminescence kit (Thermo Fisher Scientific, #WP20005).
Liposome-binding assay
Liposomes were generated using a lipid composition (11.4% PA, 63.2% POPE, 25.4% POPG) that replicates the C. reinhardtii ciliary membrane23,63. 1,2-dipalmitoyl-sn-glycero-3-phosphate (16:0 PA, #830855P), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (16:0–18:1 POPE, #850757) and 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1’-rac-glycerol) (16:1–18:1 POPG, #840457) were purchased from Avanti Polar Lipids. Liposomes were prepared using thin-film hydration followed by extrusion. Briefly, lipid stock solutions in chloroform were mixed to get 100 nM total lipid in the desired composition in a glass tube. The mixture was dried using N2 gas to form a thin layer followed by drying overnight in a vacuum desiccator. The dried film was hydrated with 1 mL buffer containing 30 mM HEPES pH 7.4 and 100 mM KCl. The buffer was first warmed to the phase transition temperature of PA (65°C) to help liposome formation. The solution was sonicated in a water bath sonicator for 5 min. The liposome solution was then extruded 50–100 times through a 100 nM polycarbonate membrane (Avanti Polar Lipids, #610005). Liposomes containing POPE (63.2%) and POPG (36.8%) were prepared using same method. Purified IFT-A was mixed with liposomes in molar ratios of 10:1, 20:1, 40:1 and 60:1 and incubated 30 on ice and analyzed by negative stain electron microscopy.
Negative-stain electron microscopy
Carbon-coated copper grids (Electron Microscopy Sciences, #CF200-Cu) were glow discharged at 30 mA for 30 s. A 4 μl aliquot of IFT-A sample (0.02 mg/mL) was applied to the grid. After incubating for 1 min, the grid was washed twice with 10 mM HEPES pH 7.4 and twice with 1.5% Uranyl formate followed by staining with 1.5% Uranyl formate for 2 min. The grids were imaged using either a 100 kV CM10 (Philips) or a 120 kV Tecnai T12 (Thermo Fisher Scientific) microscope. Images were recorded using an Ultrascan 895 CCD camera (Gatan).
Cryo-grid preparation
Cryo-EM grids of IFT-A were prepared using a Vitrobot Mark IV (Thermo Fisher Scientific). 3 μL aliquots of purified complex at concentrations of 1.1 mg/mL were applied onto glow-discharged 2-nm thick carbon coated QUANTIFOIL grids (R2/1, 200 mesh gold, Electron Microscopy Sciences, #220210). The grids were blotted for 5.5 s with a blot force of 12 and 100% humidity before being plunged into liquid ethane cooled by liquid nitrogen.
Cryo-EM data collection
Images were acquired on a Titan Krios microscope equipped with a BioQuantum K3 Imaging Filter (slit width 20 eV) and a K3 direct electron detector (Gatan) and operating at an acceleration voltage of 300 kV. Images were recorded at a defocus range of −1.5 μm to −2.5 μm with a nominal magnification of 105,000 ×, resulting in a pixel size of 0.83 Å. Each image was dose-fractionated into 50 movie frames with a total exposure time of 2.963 s, resulting in a total dose of ~57.8 electrons per Å2. SerialEM v.3.8.5 was used for data collection64.
Cryo-EM data processing
Software used for cryo-EM data processing and model building were installed and managed by SBGrid65. Data were processed using cryoSPARC v3.3.266 and RELION 4.067. Individual movies were motion corrected in RELION before being exported to cryoSPARC for Patch-based CTF estimation. Curate Exposures was used to select 25,415 good micrographs. Next, we used a template-free blob picker to pick particles on a subset of 5,492 micrographs from which we generated 14 distinct two-dimensional classes with well-resolved features. Using these 2D classes as templates for picking, we obtained 1,203,589 “large” and 696,079 “small” particles from the full dataset. Large particles corresponded to the intact IFT-A complex, whereas the small particles corresponded to individual IFT-A2 subcomplexes. After alignment in cryoSPARC, the large and small particles were transferred to RELION 4.0 for further processing. The large particles were classified with or without alignment into 4 to 6 classes. The classes were then combined into two states that differ in the position of IFT139. State 1 particles (239,280 in total) were reextracted without binning, polished and refined to a final resolution of 4.0 Å. Next, we performed signal subtraction for the A1 and A2 subcomplexes followed by 3D classification. Mask-focused refinement was performed on five local regions (shown in Fig. S1E) to obtain higher-resolution maps. Unbinned state 2 particles (563,466 in total) were polished and refined to a final resolution of 3.6 Å. To improve local resolution, mask-focused refinement was performed on the A2 subcomplex. In addition, mask-focused classification and refinement was performed on the A1 subcomplex and IFT139 to improve the resolution of these dynamic regions.
Mask-focused refinement maps were sharpened by RELION postprocessing and merged into a composite map using PHENIX combine_focused_maps68 with default settings. These composite maps were used for model building, refinement, and analysis. Maps sharpened using DeepEMhancer62 were used to make figures in ChimeraX v1.469.
Model building and refinement
Atomic models of the L. tarentolae IFT-A subunits were predicted using AlphaFold270 and docked into the cryo-EM maps using Coot v.0.9.8.371. The models were then adjusted in Coot to better fit the maps using real-space refinement. Zinc ions were added to the zinc-finger domains. The final model was refined in real space using Phenix72 with Ramachandran and secondary structure restraints applied. Model statistics were calculated with MolProbity73 implemented within Phenix. In general, only protein regions with resolved sidechains or secondary structure are included in the final, deposited models. Flexible loops and domains have been removed including the C-terminal zinc finger domain of IFT144 and the N-terminal TPRs of IFT139. More of the IFT139 N-terminus could be modeled in state 1 than state 2. An exception was made for IFT144β-prop1, which has relatively weak density due to flexibility but was left in the model.
Atomic models of IFT-A from Tetrahymena thermophila (used for validation against crosslinking data; Fig. S4), Chlamydomonas reinhardtii (used for modeling IFT-A in anterograde trains; Fig. 3) and Homo sapiens (used for interpreting human mutation data; Fig. 2F and Fig. 5) were built by superposing sections of AlphaFold2 models onto the atomic model of L. tarentolae IFT-A. Sequence alignments were performed with Clustal Omega v1.2.274.
The atomic model of IFT-A polymerized on IFT trains was generated by docking the homology model of the C. reinhardtii IFT-A into the subtomogram average of IFT-A obtained from anterograde trains (EMD-26791)40. Docking revealed that the segmented density did not correspond with the monomeric unit of IFT-A. We therefore used a lower resolution subtomogram average that contains approximately 3 copies of IFT-A (EMD-4304)19 to proliferate EMD-26791 into a linear array. Docking was guided by the A2 subcomplex, where the shape and curvature allowed unambiguous positioning. The A1 domain was then rotated into position as a rigid body. The IFT140TPR and IFT144TPR domains were then manually adjusted in Coot to fit the density. To resolve the position of IFT-A relative to IFT-B, the atomic model was docked into the composite subtomogram average of an assembled C. reinhardtii IFT train at the ciliary transition zone (EMD-15261)18.
QUANTIFICATION AND STATISTICAL ANALYSIS
Estimations of the resolution of the cryo-EM density maps (reported in Figures S1 and S3) are based on the 0.143 FSC criterion75. All statistical validation performed on the deposited models (PDB: 8F5O and PDB: 8F5P) was done using the PHENIX package73 (Table S2).
Supplementary Material
Highlights.
Cryo-EM structures of native Leishmania tarentolae IFT-A complexes
Subcomplex rearrangements enable IFT-A to polymerize into lateral IFT trains
IFT-A has lipid specificity and can coat liposomes
Anterograde IFT-A positions cargo-binding beta-propellers towards the ciliary membrane
ACKNOWLEDGMENTS
We thank S. Sterling, R. Walsh, and M. Mayer for assistance with electron microscopy at the Harvard Cryo-EM Center for Structural Biology. We thank R. Tomaino for mass spectrometry analysis, P. Aschauer for advice on preparing liposomes, J.R. Anderson for testing postprocessing strategies, and SBGrid for computing support. M.G. was supported by a Charles A. King Trust Postdoctoral Research Fellowship. A.B. is supported by NIGMS (GM141109 and GM143183), the Smith Family Foundation, and the Pew Charitable Trusts.
Footnotes
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DECLARATION OF INTERESTS
The authors declare no competing interests.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
Data Availability Statement
Composite cryo-EM maps of L. tarentolae IFT-A state 1 and 2 have deposited in the Electron Microscopy Data Bank (EMDB) with accession numbers EMD-28866 and EMD-28867. Half maps and masks are deposited as additional maps associated with these entries. Atomic models of IFT-A state 1 and state 2 have been deposited in the Protein Data Bank (PDB) with accession numbers 8F5O and 8F5P.
This paper does not report original code.
Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.