Abstract All animal cells use the motor cytoplasmic dynein 1 (dynein) to transport diverse cargo toward microtubule minus ends and to organize and position microtubule arrays such as the mitotic spindle. Cargo-specific adaptors engage with dynein to recruit and activate the motor, but the molecular mechanisms remain incompletely understood. Here, we use structural and dynamic nuclear magnetic resonance (NMR) analysis to demonstrate that the C-terminal region of human dynein light intermediate chain 1 (LIC1) is intrinsically disordered and contains two short conserved segments with helical propensity. NMR titration experiments reveal that the first helical segment (helix 1) constitutes the main interaction site for the adaptors Spindly (SPDL1), bicaudal D homolog 2 (BICD2), and Hook homolog 3 (HOOK3). In vitro binding assays show that helix 1, but not helix 2, is essential in both LIC1 and LIC2 for binding to SPDL1, BICD2, HOOK3, RAB-interacting lysosomal protein (RILP), RAB11 family-interacting protein 3 (RAB11FIP3), ninein (NIN), and trafficking kinesin-binding protein 1 (TRAK1). Helix 1 is sufficient to bind RILP, whereas other adaptors require additional segments preceding helix 1 for efficient binding. Point mutations in the C-terminal helix 1 of Caenorhabditis elegans LIC, introduced by genome editing, severely affect development, locomotion, and life span of the animal and disrupt the distribution and transport kinetics of membrane cargo in axons of mechanosensory neurons, identical to what is observed when the entire LIC C-terminal region is deleted. Deletion of the C-terminal helix 2 delays dynein-dependent spindle positioning in the one-cell embryo but overall does not significantly perturb dynein function. We conclude that helix 1 in the intrinsically disordered region of LIC provides a conserved link between dynein and structurally diverse cargo adaptor families that is critical for dynein function in vivo.

Author summary The large size and complex organization of animal cells make the correct and efficient distribution of intracellular content a challenge. The solution is to use motor proteins, which harness energy from ATP hydrolysis to walk along actin filaments or microtubules, for directional transport of cargo. The multi-subunit motor cytoplasmic dynein 1 (dynein) is responsible for transport directed toward the minus ends of microtubules. An important question is how dynein is recruited to its diverse cargo, which includes organelles such as endosomes and mitochondria, proteins, and mRNA. In this study, we use nuclear magnetic resonance spectroscopy to show that the light intermediate chain (LIC) subunit of human dynein uses a short helix in its disordered C-terminal region to bind structurally distinct adaptor proteins that connect the motor to specific cargo. We then use genome editing in the animal model C. elegans to demonstrate the functional relevance of the C-terminal LIC helix for dynein-dependent cargo transport in neurons. Thus, dynein recruitment to cargo involves a highly conserved interaction between LIC and adaptor proteins.

Citation: Celestino R, Henen MA, Gama JB, Carvalho C, McCabe M, Barbosa DJ, et al. (2019) A transient helix in the disordered region of dynein light intermediate chain links the motor to structurally diverse adaptors for cargo transport. PLoS Biol 17(1): e3000100. https://doi.org/10.1371/journal.pbio.3000100 Academic Editor: Trina Schroer, Johns Hopkins University, UNITED STATES Received: September 7, 2018; Accepted: December 14, 2018; Published: January 7, 2019 Copyright: © 2019 Celestino et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Data Availability: The 1H, 13C, and 15N chemical shift assignments have been deposited in the BioMagResBank database (http://www.bmrb.wisc.edu) under the accession number 27401. Funding: This work was financed by the Fundo Europeu de Desenvolvimento Regional (FEDER) through the Norte Portugal Regional Operational Programme (NORTE 2020), Portugal 2020 (RG); by the Fundação para a Ciência e a Tecnologia (FCT)/Ministério da Ciência, Tecnologia e Ensino Superior in the framework of the project NORTE-01-0145-FEDER-030507 (RG); by FCT fellowships IF/01015/2013/CP1157/CT0006 (RG) and SFRH/BPD/101898/2014 (DJB); by the European Research Council under the European Union’s Seventh Framework Programme, ERC grant agreement no. ERC-2013-StG-338410-DYNEINOME (RG), and by a start-up package of the University of Colorado (BV). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Competing interests: The authors have declared that no competing interests exist. Abbreviations: ALM, anterior lateral mechanosensory; AVM, anterior ventral mechanosensory; ARP1, actin-related protein 1; BICD2, bicaudal D homolog 2; BICDR1, BICD-related protein 1; CC1, first coiled-coil segment; CD, circular dichroism; CRISPR/Cas9, clustered regularly interspaced short palindromic repeat/CRISPR-associated 9; DHC-1, dynein heavy chain 1; DLI-1, dynein light intermediate chain 1; dynein, cytoplasmic dynein 1; EM, electron microscopy; GCN4, general control nondepressible 4; GFP, green fluorescent protein; GST, glutathione S-transferase; HOOK3, Hook homolog 3; HSQC, heteronuclear single quantum coherence; HC, heavy chain; IC, intermediate chain; LC, light chain; LIC, light intermediate chain; LIC-C, C-terminal region of the LIC subunit; MBP, maltose-binding protein; MIRO, mitochondrial Rho; MST, microscale thermophoresis; NEBD, nuclear envelope breakdown; NIN, ninein; NMR, nuclear magnetic resonance; ns, not significant; ppm, parts per million; NTA, nitrilotriacetic acid; RAB11FIP3, RAB11 family-interacting protein 3; RILP, RAB-interacting lysosomal protein; RNAi, RNA interference; SNB-1, synaptobrevin 1; SPDL1, Spindly; SPR, surface plasmon resonance; SSP, secondary structure propensity; TOMM-20, translocase of outer mitochondrial membrane 20; TRAK1, trafficking kinesin-binding protein 1

Introduction Microtubule-based cargo transport and force production are critical for a wide range of cellular and developmental processes. In animal cells, the 1.4-MDa complex cytoplasmic dynein 1 (dynein) is the major molecular motor with motility directed toward microtubule minus ends. Dynein-driven cargo transport is particularly crucial in highly polarized cells such as neurons. In axons, whose microtubule plus ends are uniformly oriented toward the axonal tip, dynein is responsible for the retrograde transport of diverse vesicle and organelle cargo toward the cell body. Mutations in dynein that alter axonal transport kinetics have been linked to a variety of nervous system disorders, including spinal muscular atrophy, motor neuron disease, Perry syndrome, and Charcot-Marie-Tooth 2 disease [1,2]. In addition to transporting cargo, dynein can exert pulling forces on microtubules when the motor is stably anchored at subcellular sites such as the cell cortex. A striking example of dynein-dependent force production occurs during mitosis, when cortically localized dynein pulls on astral microtubules to position and orient the bipolar spindle, which in turn defines the axis along which the cell will divide. Dynein's functional versatility implies tight regulation of localization and motor activity, the molecular basis of which has only recently begun to be understood. Dynein is a 12-subunit complex consisting of a dimerized heavy chain (HC) with a C-terminal motor domain and two copies each of five accessory chains that bind along the HC N-terminal tail: dynein intermediate chain (IC), light intermediate chain (LIC), and three types of light chain (LC). In vivo, dynein function requires the cofactor dynactin, which is itself a 1.1-MDa complex built around a short filament of actin-related protein 1 (ARP1) [3]. In recent years, a number of coiled-coil proteins, referred to as activating adaptors [4], have been shown to recruit dynein to cargo while simultaneously promoting the association of dynein with dynactin [5,6]. Dynein, dynactin, and the N-terminal coiled-coil region of activating adaptors form a stable three-way assembly capable of highly processive motility in vitro [7,8], whereas the C-terminal region of adaptors links to cargo [4]. Cryo–electron microscopy (EM) studies with the adaptors bicaudal D homolog 2 (BICD2), BICD-related protein 1 (BICDR1), and Hook homolog 3 (HOOK3) have revealed the molecular arrangement within the dynein-dynactin-adaptor assembly [3,9–11]: the adaptor coiled-coil region binds along the length of the dynactin ARP1 filament with the adaptor N terminus located at the filament's barbed end, and the N-terminal tail of dynein HC makes contact with both the ARP1 filament and the coiled-coil region of the adaptor. An additional contact between dynein and adaptors involves the C-terminal region of the LIC subunit (LIC-C) [12,13]. Whereas the highly conserved N-terminal GTPase-like domain of LIC binds tightly to the HC [12,14], the LIC-C sequence is more divergent and predicted to be disordered. Vertebrates possess two genes for LIC (LIC1 and LIC2), which may specify distinct dynein populations. Adaptors known to bind LIC1-C include BICD2 and Spindly (SPDL1), which are likely related [15,16], as well as the structurally distinct adaptors HOOK3, RAB11 family-interacting protein 3 (RAB11FIP3; hereafter referred to as FIP3), and RAB-interacting lysosomal protein (RILP) [12,17]. Whether LIC2-C also interacts with these adaptors has not been examined. SPDL1/BICD2 and HOOK3 bind to LIC1-C through a motif in their first coiled-coil segment (the CC1 box) and the N-terminal Hook domain, respectively, and point mutations in these adaptors that abrogate binding to LIC1-C compromise the formation of the dynein-dynactin-adaptor assembly [16–18]. Consequently, HOOK3 mutants that fail to bind LIC1-C do not support processive dynein runs in vitro [17]. A mutation in the CC1 box of Drosophila melanogaster BicD causes a hypomorphic loss-of-function phenotype [19], indicating that the BicD-LIC interaction is functionally relevant in vivo. Numerous other loss-of-function studies have implicated LIC in many dynein-dependent processes, including mitosis and retrograde cargo transport in axons. However, the extent to which LIC loss-of-function phenotypes reflect an important role for LIC-C is less clear, as the dynein complex becomes destabilized when LIC is absent in D. melanogaster and Aspergillus nidulans [20,21], as well as in LIC1-deficient mice [22]. Indeed, biochemical analysis indicates that the N-terminal LIC domain plays an important structural role within the dynein complex [14,23]. Thus, although LIC is clearly essential for dynein function in vivo, the specific contributions of LIC-C remain to be determined. Here, we dissect the interaction between LIC-C and dynein adaptors in vitro and in the animal model C. elegans. Nuclear magnetic resonance (NMR) analysis shows that human LIC1-C is intrinsically disordered and possesses two short segments with helical propensity. In agreement with a recent report [13], we show that helix 1 of LIC1-C is essential for binding to BICD2, SPDL1, and HOOK3, and we extend this finding to the adaptors FIP3, RILP, ninein (NIN), and trafficking kinesin-binding protein 1 (TRAK1), as well as to LIC2-C. Finally, we show that LIC-C mutants generated by genome editing in C. elegans have major defects in postembryonic cell division and retrograde axonal cargo transport, demonstrating the crucial importance of LIC-C helix 1 for dynein function in vivo.

Discussion How dynein achieves specificity for cargo is a key question underlying the functional versatility of the motor. Studies over the past few years have revealed that the N-terminal region of cargo adaptor proteins helps bring together the motor with its essential cofactor dynactin and that multiple distinct protein–protein interactions participate in the assembly of a processive dynein-dynactin-adaptor transport machine. Here, we dissected the structural determinants and the functional relevance of the interaction between the C-terminal region of the dynein LIC subunit and cargo adaptors. We present high-resolution molecular evidence that LIC-C is disordered and that a conserved segment with helical propensity, helix 1, is the main binding site for seven adaptors that differ in structure and cargo specificity. Our results agree with recent work that identified LIC1-C helix 1 as the critical structural element for the interaction with HOOK1/3, SPDL1, and BICD2 [13], and we extend these findings to RILP, NIN, FIP3, and TRAK1. The highly conserved phenylalanines and leucines of the amphipathic helix 1 are essential for the interaction, which occurs with single-digit micromolar affinity for SPDL1, BICD2, HOOK3, and RILP, suggesting a similar binding mechanism. NMR spectroscopy analysis shows that helix 1 is transient. The presence of transiently sampled secondary elements in intrinsically disordered proteins is common and may facilitate the transition to more rigid states upon binding [39–41]. The observed signal quenching upon binding to adaptors is unlikely to be caused entirely by an increased local tumbling time. Instead, micro-millisecond conformational exchange between various states of the helical elements or binding kinetics contribute significantly to the quenching. Both mechanisms suggest residual structural disorder. Indeed, various examples of disordered proteins that maintain partial disorder during interaction with other proteins have recently been reported [40,42–44]. A prior study used X-ray crystallography to show that LIC1-C helix 1 inserts into a hydrophobic cleft on HOOK3(1–160) [13]. An analogous hydrophobic cleft is likely formed by the CC1 box, the LIC1-C binding region in SPDL1 and BICD2 [13,16]. CC1 boxes are also present in other adaptors whose N-terminal regions are structurally similar to SPDL1 and BICD2, such as HAP1 and TRAK [45]. Consistent with this, we show that the first coiled-coil segment of TRAK1 interacts with LIC1-C in a helix 1-dependent manner. RILP, NIN, and FIP3 contain neither a Hook domain nor a CC1 box. The structural element that accommodates the LIC-C helix 1 in these adaptors remains to be identified. LIC1-C helix 1 is essential for adaptor binding, but we find that helix 1 on its own—i.e., LIC1(440–455)—is not sufficient for efficient binding to 5 out of 6 adaptors tested, RILP being the exception. Pull-down assays with LIC1-C truncations show that segments N-terminal to helix 1 contribute to adaptor binding. LIC1-C binding to SPDL1, FIP3, and NIN appears to be particularly sensitive to N-terminal LIC1-C truncations, as we find that LIC1(424–471) has markedly reduced affinity relative to LIC1(388–471). This is in agreement with NMR analysis, which indicates that residues 418–421 participate in the interaction with SPDL1. NMR analysis also indicates that LIC1-C helix 2 participates in the interaction with adaptors, but in vitro binding experiments show that the contribution of helix 2 to the overall binding affinity is relatively minor. Vertebrates possess two LIC paralogs that differ significantly in their C-terminal region. Prior studies in cultured cells suggest that there is functional redundancy among LIC1 and LIC2 but also hint at functional specialization in dynein-dependent cell division processes and membrane trafficking [46–53]. One attractive possibility is that the C-terminal regions of LIC1 and LIC2 discriminate between adaptors. Our results show that LIC1 and LIC2 use the same helix 1–based mechanism for adaptor binding, and we did not detect any striking differences in the ability of the two C-terminal regions to interact with our set of six adaptors. Nevertheless, it is plausible that LIC1 and LIC2 exhibit subtle preferences for specific adaptors. For example, we find that NIN binds slightly better to LIC2 than LIC1. Posttranslational modifications could possibly further modulate the affinity for adaptors in an isoform-specific manner. Previous work in vitro showed that the dynein-dynactin-adaptor assembly fails to form in HOOK3 and SPDL1 mutants that cannot bind to LIC1 [16,17] and that addition of excess LIC1-C helix 1 inhibits processive movement mediated by HOOK3 or BICD2 fragments in motility assays [13]. Here, we present direct evidence that LIC-C helix 1 is important for dynein function in vivo. Like other invertebrates, the nematode C. elegans expresses only one LIC isoform, which facilitates the identification of functionally critical regions. In all assays, the outcome of mutating the two conserved phenylalanines or leucines in the C-terminal helix 1 of DLI-1 is identical to that of deleting the entire C-terminal region. Unlike DLI-1 depletions, the phenotype of the C-terminal DLI-1 mutations cannot be attributed to destabilization of DHC-1 and therefore likely reflects defective adaptor binding. On the animal level, sterility and other developmental defects suggest failure of postembryonic cell division, similar to what was described for dli-1 null mutants [30,31]. We show on the cellular level that the C-terminal DLI-1 mutations disrupt the distribution of early endosomes and presynaptic vesicles in axons of touch receptor neurons. Since we find that axonal length is normal in day 1 adults of dli-1(L396A/L397A) and dli-1(Δ369–443) mutants, this is unlikely to be an indirect consequence of a developmental defect. Instead, the misaccumulation at the axonal tip is suggestive of impaired retrograde transport by dynein, as previously described for other dynein mutants, including the dli-1 null mutant js351 [31]. Consistent with this, we show that the frequency, velocity, and run length of early endosome movement is significantly decreased in our dli-1 mutants, with a predominant effect on retrograde movement. Neurodegeneration (i.e., axon beading) starts to become evident in dli-1 mutant day 1 adults and likely contributes to the animals' severe locomotory deficit and shortened life span. The residual retrograde movement of early endosomes in axons of dli-1 mutants could indicate that dynein retains a limited ability to form processive dynein-dynactin-adaptor complexes without the interaction between DLI-1 and adaptors. This may also explain the modest effect of dli-1 mutants on mitochondrial transport. However, it is difficult to rule out that a small fraction of wild-type maternal DLI-1, passed on from the heterozygous mother to homozygous mutant progeny, could persist in touch receptor neurons at the L4 stage and promote residual dynein activity. Nevertheless, the observation that dli-1 mutants have a more pronounced effect on early endosomes and synaptic vesicles than on mitochondria indicates that DLI-1 binding to adaptors may not be equally important for all cargo transport. In contrast to the point mutants in the DLI-1 C-terminal helix 1, there are no obvious developmental defects in animals expressing the DLI-1(Δ414–443) mutant that lacks the C-terminal helix 2, and mKate2::RAB-5 is not misaccumulated at the axonal tip of touch receptor neurons in this mutant. We have not examined neuronal cargo transport kinetics, but given that dli-1(Δ414–443) animals are healthy and propagate normally, any defects are unlikely to be substantial. We do, however, observe a delay in mitotic spindle positioning in one-cell dli-1(ΔΔ414–443) embryos. The mild phenotype contrasts with the failure of spindle assembly observed in one-cell embryos after DLI-1 depletion [30]. Furthermore, the mitotic defects of dli-1(Δ414–443) embryos are not enhanced by a null allele of the dynein cofactor nud-2, which partially compromises dynein [36,38]. Thus, analysis in vivo and in vitro indicates that LIC-C helix 2, despite its high sequence conservation, makes a relatively modest contribution to dynein function compared to helix 1. Together with the work of Lee and colleagues [13], our study establishes the molecular mechanism used by LIC to interact with structurally diverse cargo adaptors. An interesting open question is whether LIC-C could have additional binding partners besides adaptors. Two recent cryo-EM studies revealed that two dyneins can be recruited by a single dynactin-adaptor complex [10,11]. In one of the structures, an extra density, most likely corresponding to LIC-C of the first dynein, packs against the N-terminal coiled-coil of BICDR1 while simultaneously contacting one of the HCs of the second dynein [10]. It is tempting to speculate that this interaction between LIC-C and HC facilitates the incorporation of a second dynein into dynein-dynactin-adaptor assemblies.

Materials and methods DNA constructs for protein expression The cDNAs for expression of human DYNC1LI1 (UniProt ID: Q9Y6G9; residues 388–523, 388–471, 402–471, 414–471, 424–471, 440–471, 440–455, 440–523, and 472–523) and human DYNC1LI2 (UniProt ID: O43237; residues 375–492, 375–450, and 451–492) were cloned into vector pGEX-6P-1 with a single N-terminal tryptophan and a C-terminal linker (GSGSG) followed by 6xHis. The cDNAs for human BICD2 (UniProt ID: Q8TD16; residues 2–422), RAB11FIP3 (UniProt ID: O75154; residues 2–756), HOOK3 (UniProt ID: Q86VS8; residues 2–239 and 2–552), NIN (UniProt ID: Q8N4C6; residues 1–693), SPDL1 (UniProt ID: Q96EA4; residues 2–359), and TRAK1 (UniProt ID: Q9UPV9; residues 103–167 and 103–187, with and without a C-terminal fusion to the GCN4 dimeric coiled-coil sequence VKQLEDKVEELLSKNAHLENEVARLKKLV [GCN4CC]) were cloned into a 2CT-derived vector containing an N-terminal 6xHis::MBP fusion followed by a linker with a Tobacco Etch Virus nuclear-inclusion-a endopeptidase (TEV protease) cleavage site and containing a C-terminal linker (GSGSGR) followed by the Strep-tag II. The cDNA of RILP (UniProt ID: Q96NA2; residues 1–401) was cloned into the pACEBac1 vector with a C-terminal linker (GSGSGR) followed by the Strep-tag II. Protein expression and purification from bacteria All bacterial expression vectors were transformed into the Escherichia coli strain BL21, except for the NIN and HOOK3 constructs, which were transformed into the E. coli strain Rosetta. Expression was induced with 0.1 mM IPTG at 18°C overnight at an OD 600 of 0.9, and cells were harvested by centrifugation for 20 min at 4,000g. For GST::LIC::6xHis constructs used in pull-down experiments, bacterial pellets were resuspended in lysis buffer A (50 mM HEPES, 250 mM NaCl, 0.1% [v/v] Tween 20, 10 mM EDTA, 10 mM EGTA, 5 mM DTT, 1 mM phenylmethanesulfonyl fluoride [PMSF], 2 mM benzamidine-HCl, 1 mg/mL lysozyme [pH 8.0]), disrupted by sonication, and cleared by centrifugation at 34,000g for 45 min. GST::LIC::6xHis was purified by tandem affinity chromatography using glutathione agarose resin (Thermo Fisher Scientific) followed by HisPur Ni-NTA resin (Thermo Fisher Scientific). Glutathione agarose resin was incubated in batch with the cleared lysate and then washed with wash buffer A (25 mM HEPES, 250 mM NaCl, 0.1% Tween 20, 1 mM DTT, 2 mM benzamidine-HCl [pH 8.0]), and proteins were eluted on a gravity column with elution buffer A (50 mM HEPES, 150 mM NaCl, 10 mM reduced L-glutathione, 1 mM DTT, 2 mM benzamidine-HCl [pH 8.0]). Fractions containing the recombinant proteins were pooled, incubated in batch with Ni-NTA resin, and washed with wash buffer B (25 mM HEPES, 250 mM NaCl, 25 mM imidazole, 0.1% Tween 20, 1 mM DTT, 2 mM benzamidine-HCl [pH 8.0]). Proteins were eluted on a gravity column with elution buffer B (50 mM HEPES, 150 mM NaCl, 250 mM imidazole, 1 mM DTT, 2 mM benzamidine-HCl [pH 8.0]). Fractions containing the proteins were pooled and dialyzed against storage buffer (25 mM HEPES, 150 mM NaCl [pH 7.5]) or further purified by size-exclusion chromatography using a Superose 6 10/300 column (GE Healthcare) equilibrated with storage buffer. Glycerol and DTT were added to final concentrations of 10% (v/v) and 1 mM, respectively, and aliquots were flash-frozen in liquid nitrogen and stored at −80°C. Purification of LIC1::6xHis (residues 388–523, 388–471, and 472–523) for NMR spectroscopy, SPR, and MST experiments was carried out as described above with the following modifications: GST::LIC1::6xHis was captured using a GSTrap FF column (GE Healthcare) and eluted with elution buffer A. The GST moiety was cleaved off in solution with PreScission Protease, glutathione was removed by dialysis (50 mM HEPES, 150 mM NaCl [pH 8.0]), and the sample was applied again to a GSTrap FF column to remove GST and GST-tagged Prescission Protease. The flow-through containing LIC1::6His was subjected to size-exclusion chromatography using a Superdex 75 increase 10/300 GL column (GE Healthcare) in NMR buffer (50 mM sodium phosphate, 150 mM NaCl [pH 6.5]). For purification of cargo adaptors, bacterial pellets were resuspended in lysis buffer B (50 mM HEPES, 250 mM NaCl, 10 mM imidazole, 0.1% Tween 20, 1 mM DTT, 1 mM PMSF, 2 mM benzamidine-HCl, 1 mg/mL lysozyme [pH 8.0]), disrupted by sonication, and cleared by centrifugation at 34,000g for 45 min. The 6xHis::MBP::adaptor::Strep-tag II proteins were purified by tandem affinity chromatography using HisPur Ni-NTA resin followed by Strep-Tactin Sepharose resin (IBA). HisPur Ni-NTA resin was incubated in batch with the cleared lysate and then washed with wash buffer B, and proteins were eluted on a gravity column with elution buffer B. Fractions containing the recombinant proteins were pooled, incubated overnight with TEV protease to cleave off the 6xHis::MBP moiety (except for TRAK1 fragments, which were not cleaved), incubated in batch with Strep-Tactin Sepharose resin, and washed with wash buffer A. Proteins were eluted on a gravity column with elution buffer E (100 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 2.5 mM desthiobiotin [IBA] [pH 8.0]). Fractions containing the proteins were pooled and dialyzed against storage buffer or further purified by size-exclusion chromatography using a Superose 6 10/300 column equilibrated with storage buffer. Glycerol and DTT were added to final concentrations of 10% and 1 mM, respectively, and aliquots were flash-frozen in liquid nitrogen and stored at −80°C. Protein expression and purification from insect cells Bacmid recombination and virus production were performed as described previously [54]. A 500-mL culture (SFM4 medium; Hyclone) of Sf21 cells (0.8 × 106 cells/mL) was infected with RILP::Strep-tag II-encoding virus. Cells were harvested by centrifugation at 800g for 5 min. Pellets were resuspended in lysis buffer C (50 mM HEPES, 250 mM NaCl, 1 mM DTT [pH 8.0]) supplemented with EDTA-free cOmplete Protease Inhibitor Cocktail (Roche), sonicated, and cleared by centrifugation at 34,000g for 45 min. RILP::Strep-tag II was purified by batch affinity chromatography using Strep-Tactin Sepharose. The resin was washed with wash buffer C (25 mM HEPES, 250 mM NaCl, 0.1% [v/v] Tween 20, 1 mM DTT [pH 8.0]), and the protein was eluted on a gravity column with elution buffer E. Fractions containing RILP::Strep-tag II were pooled and dialyzed against storage buffer. Glycerol and DTT were added to final concentrations of 10% and 1 mM, respectively, and aliquots were flash-frozen in liquid nitrogen and stored at −80°C. GST pull-down assays Purified GST::LIC::6xHis constructs (50 pmol) were incubated with 250 pmol SPDL1(2–359)::Strep-tag II, 50 pmol BICD2(2–422)::Strep-tag II, 250 pmol NIN(1–693)::Strep-tag II, 50 pmol HOOK3(2–552)::Strep-tag II, 50 pmol RAB11FIP3(2–756)::Strep-tag II, 50 pmol RILP(1–401)::Strep-tag II, 250 pmol 6xHis::MBP::TRAK1(103–187)::Strep-tag II, 250 pmol 6xHis::MBP::TRAK1(103–167)::GCN4CC::Strep-tag II, or 250 pmol 6xHis::MBP::TRAK1(103–187)::GCN4CC::Strep-tag II for 1 h at 4°C in 150 μL pull-down buffer (50 mM HEPES, 100 mM NaCl, 5 mM DTT [pH 7.5]) containing 0.1% Tween 20 and supplemented with 15 μL of glutathione agarose resin. After washing the resin with 3 × 500 μL of the same buffer, proteins were eluted with pull-down buffer containing 15 mM reduced L-glutathione. Immunoblotting For immunoblots of purified proteins, samples were resolved by 10% SDS-PAGE and transferred to 0.2-μm nitrocellulose membranes (GE Healthcare). Membranes were blocked in PBS (4 mM KH 2 PO 4 , 16 mM Na 2 HPO 4 , 115 mM NaCl [pH 7.4]) containing 3% (w/v) BSA and 0.5% (v/v) Tween 20 and probed at 4°C overnight with mouse StrepMAB-Classic antibody (IBA) at 1 μg/mL in PBS containing 0.2% BSA and 0.1% Tween 20. Membranes were washed three times with PBS/0.1% Tween 20 (PBST), incubated with goat anti-mouse antibody coupled to HRP (Jackson ImmunoResearch, 1:10,000) for 1 h at room temperature, and washed again three times with PBST. Proteins were visualized by chemiluminescence using Pierce ECL Western Blotting Substrate (Thermo Fisher Scientific) and X-ray film (Amersham, GE Healthcare). For immunoblots of C. elegans, 100 adult hermaphrodites were collected into M9 buffer and processed for immunoblotting as described [37]. Samples were resolved on a gradient gel (4%–20%) and transferred to 0.2-μm nitrocellulose membranes. Membranes were blocked with 5% (w/v) nonfat dry milk in TBST (20 mM Tris, 140 mM NaCl, 0.1% Tween [pH 7.6]) and probed at 4°C overnight with rabbit anti-DHC-1 antibody GC4 (1:1,400, made in-house), mouse anti-FLAG M2 antibody (1:1,000, Sigma-Aldrich), or mouse anti-α-tubulin B512 antibody (1:5,000, Sigma-Aldrich). Membranes were sequentially rinsed 3× with TBST, 1× with 5% nonfat dry milk in TBST, and 3× with TBST. Membranes were incubated with goat secondary antibody coupled to HRP (Jackson ImmunoResearch, 1:10,000) for 1 h at room temperature and washed again 3× with TBST, 1× with 5% nonfat dry milk in TBST, and 3× with TBST. Proteins were visualized as described above. NMR spectroscopy For backbone resonance assignment of LIC1(388–523)::6xHis, 15N-1H HSQC, HNCACB, CBCA(co)NH, HNCO, HN(ca)CO, and HNN spectra [55] were recorded on a triple-resonance Varian 900 NMR cryoprobe spectrometer at 25°C using a 13C/15N-labeled sample in 50 mM sodium phosphate and 150 mM NaCl at pH 6.5 and 375 μM protein concentration in a standard 5-mm Shigemi tube. The 3D spectra were acquired with a nonuniform sampling (NUS) scheme generated by the NUS@HMS scheme generator software [56] with 1,024 complex data points in the direct dimension and 30% sampling of the original 96 and 80 points in the indirect 13C and 15N dimension, respectively. The spectral widths were 14,045 Hz (1H), 3,200 Hz (15N), 3,770 Hz (C = O), and 15,835 Hz (Cα/Cβ); the interscan delay was 1.7 s; and the number of scans was 16 for all experiments. The NUS-acquired data were reconstructed using the hmsIST software [56]. Zero-filling was achieved by addition of 256 points in both indirect dimensions. A solvent subtraction function was applied in the direct dimension. Further data processing and visualization were performed using NMRPipe/NMRDraw [57] and NMRFAM Sparky [58]. Resonance assignment was performed using CCPNmr Analysis software [59]. Because of high sequence redundancy and extensive peak overlap, we used a “divide-and-conquer” approach for chemical shift assignment. We measured and overlaid 15N-1H HSQC spectra for two smaller LIC1 constructs, 388–471 and 472–523, with 388–523 to facilitate the verification of assignment. The 15N-1H HSQC spectra of the small constructs were measured with 128 scans and 128 complex points in the indirect dimension. We assessed residual secondary structure using the SSP score program developed by Forman-Kay and colleagues with the re-referencing algorithm for 13CA and 13CB shifts [24]. The method combines different chemical shifts into a single residue–specific SSP score. Our input shifts were those of 1HN, 15N, 13CO, 13CA, and 13CB. The 15N R 1 and R 1ρ relaxation rate constants and 15N-1H heteronuclear NOEs of LIC1(388–523)::6xHis were measured on a Bruker 700 MHz spectrometer equipped with a triple-resonance cryoprobe at 25°C using a 15N-labeled sample at 400 μM protein concentration in 50 mM sodium phosphate buffer, 50 mM NaCl, 0.05% (w/v) NaN 3 , and 5 mM DTT at pH 6.5. For the R 1 and R 1ρ experiments, the sampling time points were 40, 88, 136, 192, 288, 392, 592, 688, 792, and 992 ms and 30, 60, 120, 150, 180, and 210 ms, respectively. During the R 1ρ relaxation time, a 15N spin-lock field of 1,433-Hz strength was applied. R 2 was calculated from R 1 and R 1ρ using the following equation: R 2 = R 1ρ + (R 1ρ –R 1 )*tg2(θ), where θ = tan-1(2πΔν/γ N B 1 ), Δν is the resonance offset, |γ N B 1 |/2π is the strength of the spin-lock field B 1 , and γ N the gyromagnetic ratio of the 15N spin. The 15N-1H heteronuclear NOEs were determined from two spectra recorded in presence and in the absence of 1H saturation in an interleaved manner. For NMR titration analysis, 15N-1H HSQC spectra with 128 complex points in the indirect dimension and 128 scans were recorded on a triple-resonance Varian 900 NMR cold-probe spectrometer at 25°C of 40 μM samples of 15N-labeled LIC1(388–523)::6xHis alone and in the presence of 0.5 and 1 equivalent of unlabeled SPDL1(2–359)::Strep-tag II, BICD2(2–422)::Strep-tag II, or HOOK3(2–239)::Strep-tag II in 50 mM sodium phosphate and 150 mM NaCl at pH 6.5. We used freshly prepared samples for each titration step to limit confusion with potential degradation peaks. Data processing and visualization were performed using NMRPipe/NMRDraw [57] and NMRFAM Sparky [58]. The 1H, 13C, and 15N chemical shift assignments have been deposited in the BioMagResBank database (http://www.bmrb.wisc.edu) under the accession number 27401. SPR SPR analysis with SPDL1, BICD2, and HOOK3 fragments was conducted with a Biacore 3000 system. His-tagged LIC1(388–523) and LIC1(388–471) constructs (ligand) were immobilized on three different flow cells (FC1–3) on an NTA sensor chip at densities of 200–300 RU, whereas the fourth flow cell (FC4) was spared for blank sensogram measurement. Concentrated stocks of Strep-tagged SPDL1(2–359), BICD2(2–422), and HOOK3(2–239) (analyte) were dialyzed exhaustively against HBS-P flow buffer (10 mM HEPES, 150 mM NaCl, 0.005% [v/v] surfactant P 20 [pH 7.4]). The background-blank sensogram was subtracted from sensograms measured with immobilized ligands. Injections for each analyte concentration were performed in triplicate. Data processing was done on Biacore evaluation software. SPR analysis with RILP::Strep-tag II was performed using a Biacore X100 system equipped with a CM5 sensor chip (GE Healthcare). Anti-GST antibody was immobilized using amine-coupling chemistry using the Amine Coupling Kit (GE Healthcare) and the GST Capture Kit (GE Healthcare) according to manufacturer's instructions. The surfaces of flow cells 1 and 2 were activated for 7 min with a 1:1 mixture of 0.1 M N-hydroxysuccinimide and 0.4 M 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide at a flow rate of 5 μL/min. Anti-GST antibody at a concentration of 30 μg/mL in 10 mM sodium acetate (pH 5.0) was immobilized at a density of 7,500 RU on flow cells 1 and 2. Surfaces were blocked with a 7-min injection of 1 M ethanolamine (pH 8.0). Anti-GST antibody high-affinity sites were blocked with 3 cycles of a 3-min injection of recombinant GST at 5 μg/mL in running buffer (10 mM HEPES, 150 mM NaCl, 3 mM EDTA, 0.05% surfactant P 20 [pH 7.4]) followed by a 2-min injection of regeneration solution (10 mM glycine-HCl [pH 2.1]). To collect kinetic binding data, at the beginning of each cycle, GST::LIC1(388–523)::6xHis or GST::LIC1(440–455)::6His in running buffer (ligand) was injected over flow cell 2 at a density of 720–530 and 550–460 RU, respectively. Flow cell 1 was injected with GST::6xHis at a density of 575–430 RU to serve as a reference surface. RILP::Strep-tag II (analyte) was injected in running buffer over the two flow cells at concentrations of 50, 11, 4.5, 1.8, and 0.3 μM at a flow rate of 30 μL/min and a temperature of 25°C. The complex was allowed to associate and dissociate for 120 and 600 s, respectively. Surfaces were regenerated with two 2-min injections of regeneration solution. Three independent runs were performed for each condition. The data were fit to a 1:1 interaction model using the evaluation module of the Biacore X100 software, version 2.0.1 (GE Healthcare). MST Measurements were carried out on a NanoTemper Monolith NT.115 pico instrument (NanoTemper Technologies) at 25°C using medium power and 20% excitation power (auto-detect-pico-red). LIC1(388–523)::6xHis was fluorescently labeled by interacting 100 μL protein solution (200 nM concentration) with 100 μL Red-Tris NTA dye (100 nM concentration). The reaction mixture was incubated at room temperature for 30 min followed by centrifugation for 10 min at 4°C with 15,000g. Then, 10 nM of labeled LIC1(388–523)::6xHis and 16 two-fold dilution series of the adaptor were loaded into 16 standard capillaries (NanoTemper Technologies) (SPDL1[2–359]::Strep-tag II highest concentration 42.5 μM; BICD2[2–422]::Strep-tag II highest concentration 36.5 μM; HOOK3[2–239]::Strep-tag II highest concentration 70 μM). We observed sigmoidal behavior of the fluorescence level over time, which allowed characterization of the interactions. Raw data were analyzed using the NanoTemper software (MO affinity analysis v 2.2.7). The signal-to-noise ratios for SPDL1(2–359)::Strep-tag II, BICD2(2–422)::Strep-tag II, and HOOK3(2–239)::Strep-tag II were 18.8, 22.1, and 7.1, respectively. CD CD spectra of LIC1(388–523)::6xHis were collected on a J-815 CD spectrometer (Jasco) with a wavelength range from 190 to 250 nm, data pitch 1 nm, standard sensitivity, 1 nm bandwidth and 20 nm/min scanning speed, at temperatures of 25, 37, 50, 60, and 70°C. The baseline of the spectrum was obtained from measurement of the buffer (50 mM sodium phosphate, 150 mM NaCl [pH 6.5]) and subtracted from the spectra of the samples to remove artificial CD signals that might originate from the optical system. Data measurement and analysis were performed using Spectra Manager version 2 (Jasco). Molar ellipticity was calculated as m°M/(10LC), where m° is the degree value (mdeg), M is the molecular weight in g/mole, L is the path length in cm, and C is the concentration in g/L. C. elegans strains Worm strains (S1 Table) were maintained at 20°C on standard nematode growth media (NGM) plates seeded with OP50 bacteria. A Mos1 transposon-based strategy (MosSCI) was used to generate strains stably expressing mKate2::RAB-5, SNB-1::mKate2, and TOMM-20(1–54)::mKate2 in touch receptor neurons [33]. Transgenes were cloned into pCFJ151 for insertion on Chromosome 2 (ttTi5605 locus), and transgene integration was confirmed by PCR. C-terminal mutants of dli-1 (Δ369–443, F392A/F393A, L396A/L397A, and Δ414–443) and 3xflag::dli-1 were generated by CRISPR/Cas9-mediated genome editing, as described previously [60,61]. Genomic sequences targeted by sgRNAs are listed in S2 Table. Modifications in genomic DNA sequence were confirmed by sequencing, and strains were outcrossed 6 times against the wild-type N2 strain to remove potential background mutations. Other fluorescent markers were subsequently introduced by mating. The dli-1 mutants Δ369–443, F392A/F393A, and L396A/L397A were maintained as heterozygotes using the GFP-marked genetic balancer nT1 [qIs51]. Homozygous F1 progeny from balanced heterozygous mothers were identified by the absence of GFP fluorescence. Life span assay Animals were collected at the L4 stage (day 0) and transferred every 2 d to a new NGM plate with bacteria. Animals were scored as alive or dead every 1–3 d. Animals were considered dead if they did not respond when touched with a platinum wire and if there was no evidence of pharyngeal pumping. Animals that were found dead on the edge of the plate, escaped, or died because of internal hatching of progeny were excluded from the assay. Imaging Body bending assay. L4 hermaphrodites were transferred to a new NGM plate with bacteria 16 h before performing the assay. For imaging, animals were transferred to a slide containing a 2-μL drop of M9. Movements were tracked at 20°C for 1 min at 40 frames per second using an SMZ 745T stereoscope (Nikon) mounted with a QIClic CCD camera (QImaging) and controlled by Micro-Manager software (Open Imaging). The wrMTrck plugin for Image J (http://www.phage.dk/plugins/wrmtrck.html) was used for automated counting of body-bends. Differential interference contrast imaging of adults animals. L4 hermaphrodites were transferred to a new NGM plate with bacteria for 16 h, paralyzed with 50 mM of sodium azide for 5 min, mounted on a freshly prepared 2% (w/v) agarose pad, and covered with an 18 mm × 18 mm coverslip (No. 1.5H, Marienfeld). Imaging was performed on an Axio Observer microscope (Zeiss) equipped with an Orca Flash 4.0 camera (Hamamatsu) controlled by ZEN 2.3 software (Zeiss). Multiple images covering the entire animal were recorded at 1 × 1 binning with a 40x NA 1.3 Plan-Neofluar objective and assembled into one image using the tiles mode in ZEN. Touch receptor neurons. To assess the axonal distribution of mKate2::RAB-5, SNB-1::mKate2, and TOMM-20(1–54)::mKate2, hermaphrodites at the day 1 adult stage were paralyzed with 50 mM of sodium azide for 5 min, transferred to a 2% agarose pad, and imaged on the Axio Observer microscope described above using an HXP 200C Illuminator (Zeiss). For mKate2::RAB-5 and SNB-1::mKate2, z-stacks (1-μm z steps) of the axonal tip and the nerve ring in ALM and AVM neurons (marked by soluble GFP) were acquired at 1 × 1 binning with a 63x NA 1.4 Plan-Apochromat objective. For TOMM-20(1–54)::mKate2, z-stacks capturing the entire ALM neuron were recorded at 1 × 1 binning with a 40x NA 1.3 Plan-Neofluar objective. For live imaging of mKate2::RAB-5 and TOMM-20(1–54)::mKate2, hermaphrodites at the L4 stage were paralyzed with 5 mM levamisole in M9 buffer for 10 min and mounted onto freshly prepared 2% or 5% agarose pads, respectively. ALM neurons were identified using the GFP signal, and only neurons with morphologically healthy axons were imaged. Time-lapse sequences of an axonal region approximately 50 μm away from the cell soma were recorded in a temperature-controlled room at 20°C, using a Nikon Eclipse Ti microscope coupled to an Andor Revolution XD spinning disk confocal system, composed of an iXon Ultra 897 CCD camera (Andor Technology), a solid-state laser combiner (ALC-UVP 350i, Andor Technology), and a CSU-X1 confocal scanner (Yokogawa Electric Corporation) controlled by Andor iQ3 software (Andor Technology). For mKate2::RAB-5, a single image was acquired every 200 ms for a total of 30 s at 1 × 1 binning using a 100x NA 1.45 Plan-Apochromat objective. For TOMM-20(1–54)::mKate2, a 5 × 0.5 μm z-stack was acquired every 5 s for a total of 5 min at 1 × 1 binning using a 60x NA 1.4 Plan-Apochromat objective. One-cell embryos. Adult gravid hermaphrodites were dissected in a watch glass filled with a 0.7× dilution of Egg Salts medium (1× medium is 118 mM NaCl, 40 mM KCl, 3.4 mM MgCl 2 , 3.4 mM CaCl 2 , 5 mM HEPES [pH 7.4]), and embryos were mounted on a 2% agarose pad. Imaging was performed using the spinning disk confocal system described above at 1 × 1 binning using an 60x NA 1.4 Plan-Apochromat objective. A 12 × 1 μm z-stack of mCherry::HIS-11 (histone H2B) and GFP::TBB-2 (β-tubulin) was acquired every 10 s from the start of pronuclear migration until the onset of cytokinesis. Image analysis Image analysis was performed using Fiji software (Image J version 1.52d). mKate2::RAB-5 and SNB-1::mKate2 levels. For axonal tip measurements, the GFP signal was used as a reference to draw a region around the last 20 μm of the ALM or AVM axon, and the integrated intensity of mKate2 fluorescence within that region was measured. The region was then expanded by a few pixels along the length of the axonal segment. The difference in integrated fluorescence intensity between the outer and inner region was used to define the integrated background intensity after normalization to the area of the inner region. The final integrated intensity of mKate2 signal at the axonal tip was then calculated by subtracting the integrated background intensity from the integrated intensity of the inner region. The same approach was used to determine integrated mKate2 intensity in the nerve ring. TOMM-20(1–54)::mKate2 distribution. To determine the number and position of mitochondria along the ALM axon (marked by GFP), line segments connecting adjacent TOMM-20(1–54)::mKate2 particles were drawn with the first segment starting at the cell body and the last segment terminating at the axonal tip. The sum of the lengths of individual line segments was equal to the total length of the axon. The distance of each TOMM-20(1–54)::mKate2 particle from the cell body was then normalized to axon length (i.e., 0% corresponds to the beginning of the axon at the cell body and 100% denotes the axonal tip). Axonal transport parameters. Only time-lapse sequences during which there was no discernable movement of the animal and that had uniformly bright mKate2 signal along the axonal segment were considered. Kymograph generation and image analysis were performed with KymoAnalyzer, a semiautomated open-source ImageJ package of macros designed to quantify transport parameters of fluorescently labeled stationary or moving particles [62]. A track is defined as a single particle trajectory, and a segment corresponds to a portion within a track of an individual moving particle framed by a pause or a reversal. Values for run length and velocity reported in this study are for segments. To determine the fraction of particles moving in the anterograde or retrograde direction, the position of the start and end point of the track was considered (i.e., a net anterograde moving particle may also have one or more segments of retrograde movement along its track and vice-versa; a net stationary particle may have one or more segments of anterograde or retrograde movement along its track). Centrosome–centrosome axis tilt in the one-cell embryo. After maximum intensity projection of z-stacks, the angle between the centrosome–centrosome axis and anterior–posterior axis at NEBD was calculated from the coordinates of the two centrosomes and the two outermost points along the embryo long axis, using the cytoplasmic GFP signal to visualize the embryo outline. Statistical analysis Statistical analysis was performed with GraphPad Prism 7.0 software. Values in figures and text are reported as mean ± SEM. The type of statistical analysis performed is described in each figure legend. Differences were considered significant at P < 0.05.

Acknowledgments The authors thank Tiago Dantas (IBMC/i3S) for critical reading of the manuscript and Joana Leite (IBMC/i3S) for help with rose diagrams. The authors also thank David Jones and Shaun Bevers (University of Colorado, Denver) for help and support with NMR, SPR, and MST measurements.