The spatiotemporal regulation of organelle positioning is critical for proper cellular function. The movement of organelles and other intracellular cargo is largely driven by the cytoskeletal motor proteins (Hirokawa et al., 2009; Reck-Peterson et al., 2018). Among them, conventional kinesin-1 is the major anterograde motor in cells that is essential for transporting mitochondria (Schwarz, 2013), ER, and Golgi-derived vesicles (Hirokawa et al., 2009) to microtubule plus ends, typically located at the cell periphery.

Kinesin-1 can exist in homodimeric and heterotetrameric forms. Kinesin-1 in the dimeric form is composed of two identical kinesin heavy chains (KHC) that can be divided into three parts: motor, stalk, and tail domains (Figure 1A; Hirokawa et al., 1989). The N-terminal motor domain functions as an engine that converts the chemical energy from ATP to the mechanical force to power kinesin walking along microtubules (Jon Kull et al., 1996). Downstream of the motor domain, a series of coiled-coils dimerize to form the stalk. The regulatory tail at the C-terminus is largely disordered (Seeger et al., 2012). In the heterotetramer form, two copies of the kinesin light chain (KLC) associate with the KHC stalk through its predicted N-terminal coiled-coil domain (Diefenbach et al., 1998; Verhey et al., 1998). The C-terminus of KLC consists of the TPR domain, which functions as a cargo binding platform to link kinesin-1 to various cellular cargo (Cockburn et al., 2018; Pernigo et al., 2013).

Figure 1 with 3 supplements see all

Download asset Open asset

Figure 1—source data 1

XL-MS data for KIF5B.

https://cdn.elifesciences.org/articles/86776/elife-86776-fig1-data1-v1.zip

Download elife-86776-fig1-data1-v1.zip

Figure 1—source data 2

XL-MS data for KIF5C.

https://cdn.elifesciences.org/articles/86776/elife-86776-fig1-data2-v1.zip

Download elife-86776-fig1-data2-v1.zip

Human kinesin-1 contains various isoforms, and dysfunction of them causes diseases. In humans, KHC is encoded by three genes: KIF5A, KIF5B, and KIF5C, which share a very similar motor domain and stalk with minor differences in the C-terminal tail, whereas KLC has four isoforms (KLC1–4). These isoforms showed distinct tissue expression patterns. KIF5B and KLC2 are ubiquitously expressed, whereas KIF5A, KIF5C, and KLC1 are neuron-enriched (Kanai et al., 2000; Rahman et al., 1998). Mutations in KIF5A and KIF5C result in defects in axonal transport and thus can cause neurodevelopmental and neurodegenerative pathologies (Crimella et al., 2012; Nicolas et al., 2018; Poirier et al., 2013; Willemsen et al., 2014).

When not bound to cargo, kinesin-1 exists in an autoinhibited state. Several early studies have proposed a model where the motility of KHC is regulated through the direct interaction between the motor and tail domains (Coy et al., 1999; Dietrich et al., 2008; Friedman and Vale, 1999; Stock et al., 1999). This interaction is facilitated by the folding of KHC at a break between CC2 and CC3, called ‘hinge,’ which positions an isoleucine–alanine–lysine (IAK) motif in the tail domain to bind in between the motor domains (Figure 1A; Kaan et al., 2011). The association between tail and motor domains restricts the relative movement of the motor domains, inhibiting KHC’s enzymatic and microtubule-binding activities (Hackney and Stock, 2000; Kaan et al., 2011).

Despite this prevailing model, recent data suggest that mutating the IAK motif or deleting the hinge does not increase the processivity of kinesin-1, making the current model controversial. Mutations in the IAK motif have been reported to disrupt kinesin-1 autoinhibition in cells, enzymatic assays, and single-molecule assays performed with cell lysates (Cai et al., 2007a; Kelliher et al., 2018; Seiler et al., 2000). However, recent in vitro single-molecule measurement using purified protein suggests that mutations in IAK did not result in a substantial increase in processive motility but increased the landing rate (Chiba et al., 2022). Similarly, deletion of the hinge was traditionally demonstrated to increase the ATPase activity and frequency of processive KHC dimer motility (Coy et al., 1999; Friedman and Vale, 1999; Kelliher et al., 2018), while recent data suggest that KHC without the hinge tends to oligomerize and shows little increase in processivity (Chiba et al., 2022). The contradictory data regarding the IAK motif and hinge indicate that kinesin-1 autoinhibition involves more than these two elements.

Kinesin-1 motility is further regulated by KLCs, where the incorporation of KLCs into heterotetrameric kinesin-1 motors inhibits microtubule binding in vitro and in vivo (Verhey et al., 1998). For instance, cellular FRET experiments implied that KLC keeps kinesin-1 in the folded state but separates the motor domain further apart (Cai et al., 2007a). Recently, in vitro single-molecule assays further demonstrated that the presence of KLCs suppresses the kinesin-1 motility (Chiba et al., 2022). However, it remains unclear whether KLCs stabilize the inhibited state of the heavy chain or promote the formation of a new structural state. Furthermore, how cargo binding to KLCs activates kinesin-1 is still an open question.

Microtubule-associated proteins function as an additional layer to regulate motor protein activity. For instance, MAP7 increases the landing rate and processivity of kinesin-1 by binding to coiled-coil 2 and recruiting kinesin-1 onto microtubules (Ferro et al., 2022; Hooikaas et al., 2019; Monroy et al., 2018). MAP7 contains an N-terminal microtubule-binding domain and a C-terminal kinesin-binding domain connected by an unstructured linker (Hooikaas et al., 2019). Interestingly, the MAP7 kinesin binding domain alone is sufficient to promote the landing rate and processivity of kinesin-1 (Hooikaas et al., 2019), suggesting that binding MAP7 to CC2 disrupts the autoinhibited state. Furthermore, truncated kinesin-1 homodimer KIF5B(1–560) that lacks the hinge and downstream elements can be further activated by MAP7 kinesin binding domain (Hooikaas et al., 2019), suggesting that autoinhibition is still present in KIF5B(1–560), which the current model can not explain.

Recently, a study combining AlphaFold structure prediction and negative staining EM proposed a new hinge position to allow the stalk to fold on itself (Weijman et al., 2022). While this study is an important prediction of KHC bends, there is minimal data to provide an overall description of how kinesin-1 adopts a folded, inhibited state. Moreover, this study did not address how the folded conformation relates to motor domain inhibition.

Here, we combined crosslinking mass spectrometry (XL-MS), negative stain electron microscopy (EM), and AlphaFold structure prediction to probe the conformation of autoinhibited full-length kinesin-1 homodimer and kinesin-1 heterotetramer. First, we performed structural studies with negative stain EM to characterize the folded conformation of kinesin-1 homodimers and heterotetramers. Both biochemical and structural analysis showed that kinesin-1 exists in folded and extended conformations. To determine the folding pattern of kinesin-1, we combined protein structure prediction using AlphaFold and XL-MS to derive an integrative molecular model of the autoinhibited full-length kinesin-1. For the kinesin-1 homodimer, we found that it does not simply fold in half via the originally proposed hinge but rather adopts a hierarchical folding pattern. A newly identified hinge in coiled-coil 3 allows the stalk to fold back on itself. The motor domain also scaffolds back to dock onto coiled-coil 2, positioning it in a head-to-tail conformation. XL-MS analysis of the heterotetramer revealed that the KLC does not change the folding pattern of the heavy chain, indicating that the light chain does not promote a new structural state but stabilizes the inhibited state. We validated our proposed autoinhibition model via mutagenesis studies to show that disruption of multiple regions is required to fully activate the full-length kinesin-1. We demonstrate that autoinhibition requires more interactions than the inhibitory tail peptide, providing the first comprehensive explanation of autoinhibition. Our model offers a conceptual framework for understanding how activation factors such as MAP7 and cargo adaptors relieve autoinhibition to promote full activation.

Our integrative modeling approach provides the first comprehensive description of the kinesin-1 autoinhibition mechanism (Figure 7). Overall, our results point to hierarchical intramolecular interactions that generate a folded autoinhibited state of kinesin-1, and several important findings can be highlighted.

Figure 7

Download asset Open asset

First, kinesin-1 does not simply fold in half via the originally proposed hinge between the CC2 and CC3a (Coy et al., 1999; Friedman and Vale, 1999). Our results demonstrate that this previously proposed hinge is instead a highly structured region that adopts a short antiparallel coiled-coil in both the dimeric (KIF5B and KIF5C) and tetrameric (KIF5B/C+KLC1) states (Figure 2—figure supplement 1 and Figure 4—figure supplement 1). This finding explains why deletion of the hinge did not result in an extended conformation for Drosophila KHC (Coy et al., 1999) and why removal of the hinge resulted in an only a partial increase in kinesin-1 activity (Chiba et al., 2022; Friedman and Vale, 1999; Kelliher et al., 2018). Our results also demonstrate that the folding of kinesin-1 depends on a short (~4 aa) break in CC3, which serves as a hinge that allows CC3b and CC4 to interact with CC2 and CC3a. Our results are consistent with and extend the work of Weijman et al., 2022. Surprisingly, mutation of this hinge (called the elbow by Weijman et al., 2022) to generate an extended coiled-coil across CC3 did not fully relieve autoinhibition in single-molecule assays. We interpret this result to mean that other regions of KIF5B can generate breaks in the coiled-coil stalk region that enable intramolecular interactions between the motor domain, stalk, and tail.

Second, we identified a novel role for CC2 in the autoinhibition of the two motor domains. In the folded state of the full-length KIF5B model, the two motor domains fold back, and one of them docks on top of CC2 in a manner that blocks the microtubule-binding interface of the motor domain. Positioning of one motor domain along CC2 also enables the C-terminus of CC4 and the tail to be positioned in between the two motor heads and contribute to autoinhibition. In the context of the truncated (1–565) protein, the motor-CC2 interaction results in a protein that, while active in single-molecule motility assays, is still inhibited when compared to the minimally dimeric construct (1–420).

Third, we confirm and extend the role of the tail domain, particularly the IAK motif, in kinesin-1 autoinhibition. Early studies suggested that the IAK motif plays an important role in autoinhibition based on truncations of the KHC from the C-terminus and a tail-bound motor domain structure (Hackney and Stock, 2000; Kaan et al., 2011; Stock et al., 1999). More recent work, however, showed that mutating IAK to AAA did not cause a substantial increase in processive motility but instead increased the landing rate (Chiba et al., 2022). In our hands, mutation of the IAK region resulted in only a partial increase of kinesin-1 activity for Drosophila KHC (Kelliher et al., 2018) and human KIF5B in cell lysate (Figure 5). Furthermore, KIF5B with IAK changed to AAA still adopts a folded conformation under EM and XL-MS (Figure 5—figure supplement 2), suggesting that the removal of IAK alone is insufficient to open up kinesin-1. Since our purified IAK mutant has mScarlet protein at the C-terminus, we observed an additional globular density in between the motor domains (Figure 5—figure supplement 2B), indicating that the regulatory tail is still in proximity to the motor domain in the absence of IAK motif.

Interestingly, we observed that deleting the whole tail (constructs 1–890 and 1–909) can achieve a higher landing rate than simply mutating the IAK residues, but the tail deletions showed lower dwell times (Figure 5B and C, Figure 5—figure supplement 1A and B ) and increased velocities (Figure 5—figure supplement 1C). We believe the tail functions as a whole element to regulate kinesin-1 activity. It may stabilize the autoinhibited state of kinesin-1 and reduce the off rate when kinesin-1 walks along microtubules. In our model, the tail domain should lie very close to the motor domain as its upstream CC4 is crosslinked to the motor domains (Figure 2C). However, since the tail region of kinesin-1 lacks enough lysine residues and is disordered, we could not observe it via EM and XL-MS, and thus cannot comment on the conformation of the tail and location of IAK motif in the autoinhibited state.

Fourth, the addition of KLC did not change the folding pattern of the heavy chain. From this, we propose that the role of KLCs is to stabilize the folded state of the heavy chain, consistent with previous work (Chiba et al., 2022; Verhey et al., 1998). Compared to the kinesin-1 homodimer, the addition of the light chain pushes CC4 closer to the motor domain (Figure 4B), which may facilitate its insertion in between the motor domain. This finding is consistent with our previous work showing that the addition of KLC results in a lower FRET efficiency (larger distance) between the two motor domains (Cai et al., 2007a) and that the N-terminal coiled-coil region of KLC, not the TPR domain, is required for driving KHC further into the autoinhibited state (Chiba et al., 2022; Verhey et al., 1998).

Our model of autoinhibition provides a framework to propose the mechanism of kinesin-1 activation by cargoes and MAPs (Figure 7). Kinesin-1 binding partners such as JIP3 (Sun et al., 2011), TRAK1/2 (Canty et al., 2021; Fenton et al., 2021; Henrichs et al., 2020), JIP1 (Fu and Holzbaur, 2013), and HAP1 (Twelvetrees et al., 2019) increase kinesin-1 dimer motility in vitro. Here, we selected TRAK1 as our target and further map the TRAK1 binding site on kinesin-1 to be in between the CC3b and CC4 (Figure 6E), which overlaps with the KLC binding site, suggesting the incompatibility for simultaneous binding. The association of TRAK1 between CC3b and CC4 may induce a conformational change to release CC4 from motor domains, thus opening the folding within the stalk, as suggested by the decreased number of crosslinks within kinesin-1 after binding to TRAK1 (Figure 6E). Besides activating kinesin-1, TRAK1 also serves as a cargo adaptor for the dynein motor (Canty et al., 2021). However, the relationship between the kinesin-1 and dynein binding sites is less well understood. Here, after examining the XL-MS data, we realized that the kinesin-1 binding site on TRAK1 is in the upstream region of the CC1 box with only a little overlap with the CC1 box (Figure 6E). Given that the dynein–dynactin complex should interact with the downstream of the CC1 box (Canty et al., 2021; Reck-Peterson et al., 2018), our XL-MS data suggest that there are separate binding sites for kinesin-1 and dynein on TRAK1.

The mechanism of kinesin-1 heterotetramer activation needs further investigation. The TPR domain in KLC functions as a cargo binding platform by binding partner proteins such as Nesprin4 (Roux et al., 2009; Wilson and Holzbaur, 2015), SKIP (Pernigo et al., 2013), and JIP1/3 (Pernigo et al., 2018; Verhey et al., 2001; Figure 7). All these cargoes contain a W-acidic or Y-acidic motif to mediate cargo recognition by KLCs (Cross and Dodding, 2019). In cellular assays, a small molecule that mimics the TPR-W-acidic interaction can activate kinesin-1 (Randall et al., 2017), and attachment of the W-acidic motif to intracellular cargo can result in kinesin-dependent cargo displacement (Kawano et al., 2012). However, in microtubule binding and single-molecule motility assays, binding of the KLC partner proteins JIP1 and SKIP1 was insufficient to activate the kinesin-1 heterotetramer, whereas binding of Nesprin4 could activate when present at saturating concentrations (Blasius et al., 2007; Chiba et al., 2022). In our model, we placed one TPR domain near the CC2 and the other on top of the motor domain (Figure 4D and E), while we also noticed that crosslinked pairs between TPR and KHC are greater than the maximum distance allowed for the BS3 crosslinkers. We propose that cargo binding to TPR may induce a conformational change in KLC that loosens the folding of KHC, the full activation still needs other factors, such as binding a kinesin-1 partner protein (Blasius et al., 2007). High-resolution structures of inhibited kinesin-1 heterotetramer are needed for understanding the detailed molecular mechanism of kinesin-1 inhibition and activation.

Full activation of kinesin-1 for cargo transport appears to require not only cargo adaptor proteins but also microtubule-associated factors such as MAP7 (Figure 7). In cells, MAP7 and its Drosophila homolog ensconsin promote kinesin-1’s microtubule association and cargo transport (Barlan et al., 2013; Chaudhary et al., 2019; Gallaud et al., 2014; Metzger et al., 2012; Sung et al., 2008; Tymanskyj et al., 2018). Using in vitro assays, addition of MAP7 directly recruited kinesin-1 to microtubules and can thereby promote the full activation of TRAK1/2-bound kinesin-1 dimer (Canty et al., 2021) and Nesprin4-bound kinesin-1 heterotetramer (Chiba et al., 2022). Based on our hierarchical folding model for kinesin-1, we suggest that MAP7 facilitates the activation of folded kinesin-1 by removing the inhibitory effects of CC2 on the motor domain. This hypothesis is supported by the findings that (i) MAP7 binds residues 418–526, which span CC2, of KHC (Hooikaas et al., 2019), and (ii) MAP7 can increase the motility of KIF5B(1–560) (Hooikaas et al., 2019), which remains in a partly inhibited state (Figure 5—figure supplement 1D). MAP7’s ability to promote kinesin-1 activation is likely also facilitated by its ability to bind to the microtubules through its N-terminal microtubule-binding domain (Hooikaas et al., 2019).

In summary, our work combined AlphaFold structure prediction, XL-MS, and EM to obtain a comprehensive description of the autoinhibition mechanism of kinesin-1. We revised the previous model by showing that autoinhibited kinesin-1 adopts a hierarchical folding pattern, where multilayer intramolecular interactions are needed to promote kinesin-1 fold into the inhibited state. The activation of kinesin-1 thus required multiple protein partners to release the intramolecular folding. Given the limitations of our studies, we did realize that some crosslinked residues in our model have distances greater than the maximum distance allowed for the BS3 crosslinkers, especially for the crosslinked pairs between TPR and heavy chains, which suggest that our current model could be partially incorrect in specific positional details. Furthermore, we could not fully resolve the role of the tail and TPR domain in maintaining the autoinhibited state. High-resolution structures of kinesin-1 homodimers and heterotetramers are needed to completely understand the autoinhibition and activation mechanisms.

The raw data from our XL-MS experiments were processed using pLink software. The crosslinked peptide pairs identified were outputted from pLink and organized into tabular formats that delineate the positions of the crosslinks. We have included files as Source Data for our manuscript. We also attached protein sequence information for each XL-MS experiment. Below are the instructions on how to interpret the data. KIF5B: We have performed three XL-MS experiments on KIF5B and chose one of them (KIF5B_xlink_sites_1) as representative data to present in our manuscript (Figure 1F, Figure 2). The XL-MS results are: KIF5B_pLINK_1, 2, and 3 for identified crosslinked peptide pairs; KIF5B_xlink_sites_1, 2, and 3 for crosslinked positions in tabular format. KIF5C: We performed one XL-MS analysis on KIF5C and presented it in our manuscript (Figure 1—figure supplement 1F). The XL-MS results are: KIF5C_pLINK for identified crosslinked peptide pairs; KIF5C_xlink_sites for crosslinked positions in tabular format.KIF5B-KLC1: We have performed two XL-MS on KIF5B-KLC1 and selected one of them ( KIF5B-KLC1_xlink_sites_1) as representative data to show in our manuscript (Figure 3—figure supplement 1F and Figure 3—figure supplement 2C). The XL-MS results are: KIF5B-KLC1_pLINK_1 and 2 for identified crosslinked peptide pairs; KIF5B-KLC1_xlink_sites_1 and 2 for crosslinked positions in tabular format. KIF5C-KLC1: We have performed two XL-MS on KIF5C-KLC1 and selected one of them ( KIF5C-KLC1_xlink_sites_1) as representative data to show in our manuscript (Figure 3F, Figure 4, and Figure 3—figure supplement 2D). The XL-MS results are: KIF5C-KLC1_pLINK_1 and 2 for identified crosslinked peptide pairs; KIF5C-KLC1_xlink_sites_1 and 2 for crosslinked positions in tabular format. KIF5B (1-565): We performed one XL-MS on KIF5B (1-565) and presented it in our manuscript (Figure 5—figure supplement 1D). The XL-MS results are: KIF5B(1_565)_pLINK for identified crosslinked peptide pairs; KIF5B(1_565)_xlink_sites for crosslinked positions in tabular format.KIF5B_IAK_AAA: We performed one XL-MS on KIF5B (IAK/AAA) and presented it in our manuscript (Figure 5—figure supplement 2C). The XL-MS results are: KIF5B_IAK_AAA_pLINK for identified crosslinked peptide pairs; KIF5B_IAK_AAA_xlink_sites for crosslinked positions in tabular format. TRAK1-KIF5B: We conducted one XL-MS analysis on TRAK1-KIF5B and presented it in our manuscript (Figure 6E). The XL-MS results are: TRAK1_KIF5B_pLINK for identified crosslinked peptide pairs; TRAK1_KIF5B_xlink_sites for crosslinked positions in tabular format.

1. 1. Amos LA

   (1987) Kinesin from pig brain studied by electron microscopy

   Journal of Cell Science 87:105–111.

   https://doi.org/10.1242/jcs.87.1.105

   • PubMed
   • Google Scholar
2. 1. Atherton J
   2. Farabella I
   3. Yu IM
   4. Rosenfeld SS
   5. Houdusse A
   6. Topf M
   7. Moores CA

   (2014) Conserved mechanisms of microtubule-stimulated ADP release, ATP binding, and force generation in transport kinesins

   eLife 3:e03680.

   https://doi.org/10.7554/eLife.03680

   • PubMed
   • Google Scholar
3. 1. Barlan K
   2. Lu W
   3. Gelfand VI

   (2013) The microtubule-binding protein ensconsin is an essential cofactor of kinesin-1

   Current Biology 23:317–322.

   https://doi.org/10.1016/j.cub.2013.01.008

   • PubMed
   • Google Scholar
4. 1. Blasius TL
   2. Cai D
   3. Jih GT
   4. Toret CP
   5. Verhey KJ

   (2007) Two binding partners cooperate to activate the molecular motor Kinesin-1

   The Journal of Cell Biology 176:11–17.

   https://doi.org/10.1083/jcb.200605099

   • PubMed
   • Google Scholar
5. 1. Cai D
   2. Hoppe AD
   3. Swanson JA
   4. Verhey KJ

   (2007a) Kinesin-1 structural organization and conformational changes revealed by FRET stoichiometry in live cells

   The Journal of Cell Biology 176:51–63.

   https://doi.org/10.1083/jcb.200605097

   • PubMed
   • Google Scholar
6. 1. Cai D
   2. Verhey KJ
   3. Meyhöfer E

   (2007b) Tracking single Kinesin molecules in the cytoplasm of mammalian cells

   Biophysical Journal 92:4137–4144.

   https://doi.org/10.1529/biophysj.106.100206

   • PubMed
   • Google Scholar
7. Preprint

   1. Canty JT
   2. Hensley A
   3. Yildiz A

   (2021) TRAK Adaptors Coordinate the Recruitment and Activation of Dynein and Kinesin to Control Mitochondrial Transport

   bioRxiv.

   https://doi.org/10.1101/2021.07.30.454553

   • Google Scholar
8. 1. Chaudhary AR
   2. Lu H
   3. Krementsova EB
   4. Bookwalter CS
   5. Trybus KM
   6. Hendricks AG

   (2019) MAP7 regulates organelle transport by recruiting kinesin-1 to microtubules

   The Journal of Biological Chemistry 294:10160–10171.

   https://doi.org/10.1074/jbc.RA119.008052

   • PubMed
   • Google Scholar
9. 1. Chen ZL
   2. Meng JM
   3. Cao Y
   4. Yin JL
   5. Fang RQ
   6. Fan SB
   7. Liu C
   8. Zeng WF
   9. Ding YH
   10. Tan D
   11. Wu L
   12. Zhou WJ
   13. Chi H
   14. Sun RX
   15. Dong MQ
   16. He SM

   (2019) A high-speed search engine pLink 2 with systematic evaluation for proteome-scale identification of cross-linked peptides

   Nature Communications 10:3404.

   https://doi.org/10.1038/s41467-019-11337-z

   • Google Scholar
10. 1. Chiba K
    2. Ori-McKenney KM
    3. Niwa S
    4. McKenney RJ

    (2022) Synergistic autoinhibition and activation mechanisms control kinesin-1 motor activity

    Cell Reports 39:110900.

    https://doi.org/10.1016/j.celrep.2022.110900

    • Google Scholar
11. Conference

    1. Cianfrocco MA
    2. Wong-Barnum M
    3. Youn C
    4. Wagner R
    5. Leschziner A

    (2017) COSMIC2: A Science Gateway for Cryo-Electron Microscopy Structure Determination

    Proceedings of the Practice and Experience in Advanced Research Computing 2017 on Sustainability, Success and Impact, Article Article 22.

    https://doi.org/10.1145/3093338.3093390

    • Google Scholar
12. 1. Cockburn JJB
    2. Hesketh SJ
    3. Mulhair P
    4. Thomsen M
    5. O’Connell MJ
    6. Way M

    (2018) Insights into Kinesin-1 Activation from the Crystal Structure of KLC2 Bound to JIP3

    Structure 26:1486–1498.

    https://doi.org/10.1016/j.str.2018.07.011

    • PubMed
    • Google Scholar
13. 1. Coy DL
    2. Hancock WO
    3. Wagenbach M
    4. Howard J

    (1999) Kinesin’s tail domain is an inhibitory regulator of the motor domain

    Nature Cell Biology 1:288–292.

    https://doi.org/10.1038/13001

    • PubMed
    • Google Scholar
14. 1. Crimella C
    2. Baschirotto C
    3. Arnoldi A
    4. Tonelli A
    5. Tenderini E
    6. Airoldi G
    7. Martinuzzi A
    8. Trabacca A
    9. Losito L
    10. Scarlato M
    11. Benedetti S
    12. Scarpini E
    13. Spinicci G
    14. Bresolin N
    15. Bassi MT

    (2012) Mutations in the motor and stalk domains of KIF5A in spastic paraplegia type 10 and in axonal Charcot–Marie–Tooth type 2

    Clinical Genetics 82:157–164.

    https://doi.org/10.1111/j.1399-0004.2011.01717.x

    • Google Scholar
15. 1. Cross JA
    2. Dodding MP

    (2019) Motor-cargo adaptors at the organelle-cytoskeleton interface

    Current Opinion in Cell Biology 59:16–23.

    https://doi.org/10.1016/j.ceb.2019.02.010

    • PubMed
    • Google Scholar
16. 1. Delorenzi M
    2. Speed T

    (2002) An HMM model for coiled-coil domains and a comparison with PSSM-based predictions

    Bioinformatics 18:617–625.

    https://doi.org/10.1093/bioinformatics/18.4.617

    • PubMed
    • Google Scholar
17. 1. Diefenbach RJ
    2. Mackay JP
    3. Armati PJ
    4. Cunningham AL

    (1998) The C-terminal region of the stalk domain of ubiquitous human kinesin heavy chain contains the binding site for kinesin light chain

    Biochemistry 37:16663–16670.

    https://doi.org/10.1021/bi981163r

    • PubMed
    • Google Scholar
18. 1. Dietrich KA
    2. Sindelar CV
    3. Brewer PD
    4. Downing KH
    5. Cremo CR
    6. Rice SE

    (2008) The kinesin-1 motor protein is regulated by a direct interaction of its head and tail

    PNAS 105:8938–8943.

    https://doi.org/10.1073/pnas.0803575105

    • PubMed
    • Google Scholar
19. Preprint

    1. Evans R
    2. O’Neill M
    3. Pritzel A
    4. Antropova N
    5. Senior A
    6. Green T
    7. Žídek A
    8. Bates R
    9. Blackwell S
    10. Yim J
    11. Ronneberger O
    12. Bodenstein S
    13. Zielinski M
    14. Bridgland A
    15. Potapenko A
    16. Cowie A
    17. Tunyasuvunakool K
    18. Jain R
    19. Clancy E
    20. Kohli P
    21. Jumper J
    22. Hassabis D

    (2022) Protein Complex Prediction with AlphaFold-Multimer

    bioRxiv.

    https://doi.org/10.1101/2021.10.04.463034

    • Google Scholar
20. 1. Fenton AR
    2. Jongens TA
    3. Holzbaur ELF

    (2021) Mitochondrial adaptor TRAK2 activates and functionally links opposing kinesin and dynein motors

    Nature Communications 12:4578.

    https://doi.org/10.1038/s41467-021-24862-7

    • PubMed
    • Google Scholar
21. 1. Ferro LS
    2. Fang Q
    3. Eshun-Wilson L
    4. Fernandes J
    5. Jack A
    6. Farrell DP
    7. Golcuk M
    8. Huijben T
    9. Costa K
    10. Gur M
    11. DiMaio F
    12. Nogales E
    13. Yildiz A

    (2022) Structural and functional insight into regulation of kinesin-1 by microtubule-associated protein MAP7

    Science 375:326–331.

    https://doi.org/10.1126/science.abf6154

    • PubMed
    • Google Scholar
22. 1. Friedman DS
    2. Vale RD

    (1999) Single-molecule analysis of kinesin motility reveals regulation by the cargo-binding tail domain

    Nature Cell Biology 1:293–297.

    https://doi.org/10.1038/13008

    • PubMed
    • Google Scholar
23. 1. Fu M
    2. Holzbaur ELF

    (2013) JIP1 regulates the directionality of APP axonal transport by coordinating kinesin and dynein motors

    The Journal of Cell Biology 202:495–508.

    https://doi.org/10.1083/jcb.201302078

    • PubMed
    • Google Scholar
24. 1. Gabler F
    2. Nam S-Z
    3. Till S
    4. Mirdita M
    5. Steinegger M
    6. Söding J
    7. Lupas AN
    8. Alva V

    (2020) Protein Sequence analysis using the MPI Bioinformatics Toolkit

    Current Protocols in Bioinformatics 72:e108.

    https://doi.org/10.1002/cpbi.108

    • PubMed
    • Google Scholar
25. 1. Gallaud E
    2. Caous R
    3. Pascal A
    4. Bazile F
    5. Gagné JP
    6. Huet S
    7. Poirier GG
    8. Chrétien D
    9. Richard-Parpaillon L
    10. Giet R

    (2014) Ensconsin/Map7 promotes microtubule growth and centrosome separation in Drosophila neural stem cells

    The Journal of Cell Biology 204:1111–1121.

    https://doi.org/10.1083/jcb.201311094

    • PubMed
    • Google Scholar
26. Preprint

    1. Graham M
    2. Combe C
    3. Kolbowski L
    4. Rappsilber J

    (2019) xiView: A Common Platform for the Downstream Analysis of Crosslinking Mass Spectrometry Data

    bioRxiv.

    https://doi.org/10.1101/561829

    • Google Scholar
27. 1. Hackney DD
    2. Stock MF

    (2000) Kinesin’s IAK tail domain inhibits initial microtubule-stimulated ADP release

    Nature Cell Biology 2:257–260.

    https://doi.org/10.1038/35010525

    • PubMed
    • Google Scholar
28. 1. Henrichs V
    2. Grycova L
    3. Barinka C
    4. Nahacka Z
    5. Neuzil J
    6. Diez S
    7. Rohlena J
    8. Braun M
    9. Lansky Z

    (2020) Mitochondria-adaptor TRAK1 promotes kinesin-1 driven transport in crowded environments

    Nature Communications 11:3123.

    https://doi.org/10.1038/s41467-020-16972-5

    • PubMed
    • Google Scholar
29. 1. Hirokawa N
    2. Pfister KK
    3. Yorifuji H
    4. Wagner MC
    5. Brady ST
    6. Bloom GS

    (1989) Submolecular domains of bovine brain kinesin identified by electron microscopy and monoclonal antibody decoration

    Cell 56:867–878.

    https://doi.org/10.1016/0092-8674(89)90691-0

    • PubMed
    • Google Scholar
30. 1. Hirokawa N
    2. Noda Y
    3. Tanaka Y
    4. Niwa S

    (2009) Kinesin superfamily motor proteins and intracellular transport

    Nature Reviews. Molecular Cell Biology 10:682–696.

    https://doi.org/10.1038/nrm2774

    • PubMed
    • Google Scholar
31. 1. Hooikaas PJ
    2. Martin M
    3. Mühlethaler T
    4. Kuijntjes G-J
    5. Peeters CAE
    6. Katrukha EA
    7. Ferrari L
    8. Stucchi R
    9. Verhagen DGF
    10. van Riel WE
    11. Grigoriev I
    12. Altelaar AFM
    13. Hoogenraad CC
    14. Rüdiger SGD
    15. Steinmetz MO
    16. Kapitein LC
    17. Akhmanova A

    (2019) MAP7 family proteins regulate kinesin-1 recruitment and activation

    The Journal of Cell Biology 218:1298–1318.

    https://doi.org/10.1083/jcb.201808065

    • PubMed
    • Google Scholar
32. 1. Jon Kull F
    2. Sablin EP
    3. Lau R
    4. Fletterick RJ
    5. Vale RD

    (1996) Crystal structure of the kinesin motor domain reveals a structural similarity to myosin

    Nature 380:550–555.

    https://doi.org/10.1038/380550a0

    • Google Scholar
33. 1. Jumper J
    2. Evans R
    3. Pritzel A
    4. Green T
    5. Figurnov M
    6. Ronneberger O
    7. Tunyasuvunakool K
    8. Bates R
    9. Žídek A
    10. Potapenko A
    11. Bridgland A
    12. Meyer C
    13. Kohl SAA
    14. Ballard AJ
    15. Cowie A
    16. Romera-Paredes B
    17. Nikolov S
    18. Jain R
    19. Adler J
    20. Back T
    21. Petersen S
    22. Reiman D
    23. Clancy E
    24. Zielinski M
    25. Steinegger M
    26. Pacholska M
    27. Berghammer T
    28. Bodenstein S
    29. Silver D
    30. Vinyals O
    31. Senior AW
    32. Kavukcuoglu K
    33. Kohli P
    34. Hassabis D

    (2021) Highly accurate protein structure prediction with AlphaFold

    Nature 596:583–589.

    https://doi.org/10.1038/s41586-021-03819-2

    • PubMed
    • Google Scholar
34. 1. Kaan HYK
    2. Hackney DD
    3. Kozielski F

    (2011) The structure of the kinesin-1 motor-tail complex reveals the mechanism of autoinhibition

    Science 333:883–885.

    https://doi.org/10.1126/science.1204824

    • PubMed
    • Google Scholar
35. 1. Kanai Y
    2. Okada Y
    3. Tanaka Y
    4. Harada A
    5. Terada S
    6. Hirokawa N

    (2000) KIF5C, a novel neuronal kinesin enriched in motor neurons

    The Journal of Neuroscience 20:6374–6384.

    https://doi.org/10.1523/JNEUROSCI.20-17-06374.2000

    • PubMed
    • Google Scholar
36. 1. Kawano T
    2. Araseki M
    3. Araki Y
    4. Kinjo M
    5. Yamamoto T
    6. Suzuki T

    (2012) A small peptide sequence is sufficient for initiating kinesin-1 activation through part of TPR region of KLC1

    Traffic 13:834–848.

    https://doi.org/10.1111/j.1600-0854.2012.01350.x

    • PubMed
    • Google Scholar
37. 1. Kelliher MT
    2. Yue Y
    3. Ng A
    4. Kamiyama D
    5. Huang B
    6. Verhey KJ
    7. Wildonger J

    (2018) Autoinhibition of kinesin-1 is essential to the dendrite-specific localization of Golgi outposts

    The Journal of Cell Biology 217:2531–2547.

    https://doi.org/10.1083/jcb.201708096

    • PubMed
    • Google Scholar
38. 1. Kimanius D
    2. Dong L
    3. Sharov G
    4. Nakane T
    5. Scheres SHW

    (2021) New tools for automated cryo-EM single-particle analysis in RELION-4.0

    The Biochemical Journal 478:4169–4185.

    https://doi.org/10.1042/BCJ20210708

    • PubMed
    • Google Scholar
39. 1. Kuznetsov SA
    2. Vaisberg EA
    3. Shanina NA
    4. Magretova NN
    5. Chernyak VY
    6. Gelfand VI

    (1988) The quaternary structure of bovine brain kinesin

    The EMBO Journal 7:353–356.

    https://doi.org/10.1002/j.1460-2075.1988.tb02820.x

    • PubMed
    • Google Scholar
40. 1. Merkley ED
    2. Rysavy S
    3. Kahraman A
    4. Hafen RP
    5. Daggett V
    6. Adkins JN

    (2014) Distance restraints from crosslinking mass spectrometry: mining a molecular dynamics simulation database to evaluate lysine-lysine distances

    Protein Science 23:747–759.

    https://doi.org/10.1002/pro.2458

    • PubMed
    • Google Scholar
41. 1. Metzger T
    2. Gache V
    3. Xu M
    4. Cadot B
    5. Folker ES
    6. Richardson BE
    7. Gomes ER
    8. Baylies MK

    (2012) MAP and kinesin-dependent nuclear positioning is required for skeletal muscle function

    Nature 484:120–124.

    https://doi.org/10.1038/nature10914

    • PubMed
    • Google Scholar
42. 1. Mirdita M
    2. Schütze K
    3. Moriwaki Y
    4. Heo L
    5. Ovchinnikov S
    6. Steinegger M

    (2022) ColabFold: making protein folding accessible to all

    Nature Methods 19:679–682.

    https://doi.org/10.1038/s41592-022-01488-1

    • PubMed
    • Google Scholar
43. 1. Monroy BY
    2. Sawyer DL
    3. Ackermann BE
    4. Borden MM
    5. Tan TC
    6. Ori-McKenney KM

    (2018) Competition between microtubule-associated proteins directs motor transport

    Nature Communications 9:1487.

    https://doi.org/10.1038/s41467-018-03909-2

    • PubMed
    • Google Scholar
44. 1. Navone F
    2. Niclas J
    3. Hom-Booher N
    4. Sparks L
    5. Bernstein HD
    6. McCaffrey G
    7. Vale RD

    (1992) Cloning and expression of a human kinesin heavy chain gene: interaction of the COOH-terminal domain with cytoplasmic microtubules in transfected CV-1 cells

    The Journal of Cell Biology 117:1263–1275.

    https://doi.org/10.1083/jcb.117.6.1263

    • PubMed
    • Google Scholar
45. 1. Nicolas A
    2. Kenna KP
    3. Renton AE
    4. Ticozzi N
    5. Faghri F
    6. Chia R
    7. Dominov JA
    8. Kenna BJ
    9. Nalls MA
    10. Keagle P
    11. Rivera AM
    12. van Rheenen W
    13. Murphy NA
    14. van Vugt J
    15. Geiger JT
    16. Van der Spek RA
    17. Pliner HA
    18. Shankaracharya S

    (2018) Genome-wide analyses identify KIF5A as a Novel ALS Gene

    Neuron 97:1268–1283.

    https://doi.org/10.1016/j.neuron.2018.02.027

    • Google Scholar
46. 1. Pernigo S
    2. Lamprecht A
    3. Steiner RA
    4. Dodding MP

    (2013) Structural basis for kinesin-1:cargo recognition

    Science 340:356–359.

    https://doi.org/10.1126/science.1234264

    • PubMed
    • Google Scholar
47. 1. Pernigo S
    2. Chegkazi MS
    3. Yip YY
    4. Treacy C
    5. Glorani G
    6. Hansen K
    7. Politis A
    8. Bui S
    9. Dodding MP
    10. Steiner RA

    (2018) Structural basis for isoform-specific kinesin-1 recognition of Y-acidic cargo adaptors

    eLife 7:e38362.

    https://doi.org/10.7554/eLife.38362

    • PubMed
    • Google Scholar
48. 1. Pettersen EF
    2. Goddard TD
    3. Huang CC
    4. Meng EC
    5. Couch GS
    6. Croll TI
    7. Morris JH
    8. Ferrin TE

    (2021) UCSF ChimeraX: Structure visualization for researchers, educators, and developers

    Protein Science 30:70–82.

    https://doi.org/10.1002/pro.3943

    • PubMed
    • Google Scholar
49. 1. Poirier K
    2. Lebrun N
    3. Broix L
    4. Tian G
    5. Saillour Y
    6. Boscheron C
    7. Parrini E
    8. Valence S
    9. Pierre BS
    10. Oger M
    11. Lacombe D
    12. Geneviève D
    13. Fontana E
    14. Darra F
    15. Cances C
    16. Barth M
    17. Bonneau D
    18. Bernadina BD
    19. N’guyen S
    20. Gitiaux C
    21. Parent P
    22. des Portes V
    23. Pedespan JM
    24. Legrez V
    25. Castelnau-Ptakine L
    26. Nitschke P
    27. Hieu T
    28. Masson C
    29. Zelenika D
    30. Andrieux A
    31. Francis F
    32. Guerrini R
    33. Cowan NJ
    34. Bahi-Buisson N
    35. Chelly J

    (2013) Mutations in TUBG1, DYNC1H1, KIF5C and KIF2A cause malformations of cortical development and microcephaly

    Nature Genetics 45:639–647.

    https://doi.org/10.1038/ng.2613

    • PubMed
    • Google Scholar
50. 1. Punjani A
    2. Rubinstein JL
    3. Fleet DJ
    4. Brubaker MA

    (2017) cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination

    Nature Methods 14:290–296.

    https://doi.org/10.1038/nmeth.4169

    • PubMed
    • Google Scholar
51. 1. Rahman A
    2. Friedman DS
    3. Goldstein LS

    (1998) Two kinesin light chain genes in mice. Identification and characterization of the encoded proteins

    The Journal of Biological Chemistry 273:15395–15403.

    https://doi.org/10.1074/jbc.273.25.15395

    • PubMed
    • Google Scholar
52. 1. Randall TS
    2. Yip YY
    3. Wallock-Richards DJ
    4. Pfisterer K
    5. Sanger A
    6. Ficek W
    7. Steiner RA
    8. Beavil AJ
    9. Parsons M
    10. Dodding MP

    (2017) A small-molecule activator of kinesin-1 drives remodeling of the microtubule network

    PNAS 114:13738–13743.

    https://doi.org/10.1073/pnas.1715115115

    • PubMed
    • Google Scholar
53. 1. Reck-Peterson SL
    2. Redwine WB
    3. Vale RD
    4. Carter AP

    (2018) The cytoplasmic dynein transport machinery and its many cargoes

    Nature Reviews Molecular Cell Biology 19:382–398.

    https://doi.org/10.1038/s41580-018-0004-3

    • Google Scholar
54. 1. Rohou A
    2. Grigorieff N

    (2015) CTFFIND4: Fast and accurate defocus estimation from electron micrographs

    Journal of Structural Biology 192:216–221.

    https://doi.org/10.1016/j.jsb.2015.08.008

    • PubMed
    • Google Scholar
55. 1. Roux KJ
    2. Crisp ML
    3. Liu Q
    4. Kim D
    5. Kozlov S
    6. Stewart CL
    7. Burke B

    (2009) Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization

    PNAS 106:2194–2199.

    https://doi.org/10.1073/pnas.0808602106

    • PubMed
    • Google Scholar
56. 1. Schwarz TL

    (2013) Mitochondrial trafficking in neurons

    Cold Spring Harbor Perspectives in Biology 5:a011304.

    https://doi.org/10.1101/cshperspect.a011304

    • PubMed
    • Google Scholar
57. 1. Seeger MA
    2. Zhang Y
    3. Rice SE

    (2012) Kinesin tail domains are intrinsically disordered

    Proteins 80:2437–2446.

    https://doi.org/10.1002/prot.24128

    • PubMed
    • Google Scholar
58. 1. Seiler S
    2. Kirchner J
    3. Horn C
    4. Kallipolitou A
    5. Woehlke G
    6. Schliwa M

    (2000) Cargo binding and regulatory sites in the tail of fungal conventional kinesin

    Nature Cell Biology 2:333–338.

    https://doi.org/10.1038/35014022

    • PubMed
    • Google Scholar
59. 1. Solon AL
    2. Tan Z
    3. Schutt KL
    4. Jepsen L
    5. Haynes SE
    6. Nesvizhskii AI
    7. Sept D
    8. Stumpff J
    9. Ohi R
    10. Cianfrocco MA

    (2021) Kinesin-binding protein remodels the kinesin motor to prevent microtubule binding

    Science Advances 7:eabj9812.

    https://doi.org/10.1126/sciadv.abj9812

    • PubMed
    • Google Scholar
60. 1. Stock MF
    2. Guerrero J
    3. Cobb B
    4. Eggers CT
    5. Huang TG
    6. Li X
    7. Hackney DD

    (1999) Formation of the compact confomer of kinesin requires a COOH-terminal heavy chain domain and inhibits microtubule-stimulated ATPase activity

    The Journal of Biological Chemistry 274:14617–14623.

    https://doi.org/10.1074/jbc.274.21.14617

    • PubMed
    • Google Scholar
61. 1. Suloway C
    2. Pulokas J
    3. Fellmann D
    4. Cheng A
    5. Guerra F
    6. Quispe J
    7. Stagg S
    8. Potter CS
    9. Carragher B

    (2005) Automated molecular microscopy: the new Leginon system

    Journal of Structural Biology 151:41–60.

    https://doi.org/10.1016/j.jsb.2005.03.010

    • PubMed
    • Google Scholar
62. 1. Sun F
    2. Zhu C
    3. Dixit R
    4. Cavalli V

    (2011) Sunday Driver/JIP3 binds kinesin heavy chain directly and enhances its motility

    The EMBO Journal 30:3416–3429.

    https://doi.org/10.1038/emboj.2011.229

    • PubMed
    • Google Scholar
63. 1. Sung HH
    2. Telley IA
    3. Papadaki P
    4. Ephrussi A
    5. Surrey T
    6. Rørth P

    (2008) Drosophila ensconsin promotes productive recruitment of Kinesin-1 to microtubules

    Developmental Cell 15:866–876.

    https://doi.org/10.1016/j.devcel.2008.10.006

    • PubMed
    • Google Scholar
64. 1. Twelvetrees AE
    2. Lesept F
    3. Holzbaur ELF
    4. Kittler JT

    (2019) The adaptor proteins HAP1a and GRIP1 collaborate to activate the kinesin-1 isoform KIF5C

    Journal of Cell Science 132:jcs215822.

    https://doi.org/10.1242/jcs.215822

    • PubMed
    • Google Scholar
65. 1. Tymanskyj SR
    2. Yang BH
    3. Verhey KJ
    4. Ma L

    (2018) MAP7 regulates axon morphogenesis by recruiting kinesin-1 to microtubules and modulating organelle transport

    eLife 7:e36374.

    https://doi.org/10.7554/eLife.36374

    • PubMed
    • Google Scholar
66. 1. van Spronsen M
    2. Mikhaylova M
    3. Lipka J
    4. Schlager MA
    5. van den Heuvel DJ
    6. Kuijpers M
    7. Wulf PS
    8. Keijzer N
    9. Demmers J
    10. Kapitein LC
    11. Jaarsma D
    12. Gerritsen HC
    13. Akhmanova A
    14. Hoogenraad CC

    (2013) TRAK/Milton motor-adaptor proteins steer mitochondrial trafficking to axons and dendrites

    Neuron 77:485–502.

    https://doi.org/10.1016/j.neuron.2012.11.027

    • PubMed
    • Google Scholar
67. 1. Verhey KJ
    2. Lizotte DL
    3. Abramson T
    4. Barenboim L
    5. Schnapp BJ
    6. Rapoport TA

    (1998) Light chain-dependent regulation of Kinesin’s interaction with microtubules

    The Journal of Cell Biology 143:1053–1066.

    https://doi.org/10.1083/jcb.143.4.1053

    • PubMed
    • Google Scholar
68. 1. Verhey KJ
    2. Meyer D
    3. Deehan R
    4. Blenis J
    5. Schnapp BJ
    6. Rapoport TA
    7. Margolis B

    (2001) Cargo of kinesin identified as JIP scaffolding proteins and associated signaling molecules

    The Journal of Cell Biology 152:959–970.

    https://doi.org/10.1083/jcb.152.5.959

    • PubMed
    • Google Scholar
69. 1. Weijman JF
    2. Yadav SKN
    3. Surridge KJ
    4. Cross JA
    5. Borucu U
    6. Mantell J
    7. Woolfson DN
    8. Schaffitzel C
    9. Dodding MP

    (2022) Molecular architecture of the autoinhibited kinesin-1 lambda particle

    Science Advances 8:eabp9660.

    https://doi.org/10.1126/sciadv.abp9660

    • PubMed
    • Google Scholar
70. 1. Willemsen MH
    2. Ba W
    3. Wissink-Lindhout WM
    4. de Brouwer APM
    5. Haas SA
    6. Bienek M
    7. Hu H
    8. Vissers LELM
    9. van Bokhoven H
    10. Kalscheuer V
    11. Nadif Kasri N
    12. Kleefstra T

    (2014) Involvement of the kinesin family members KIF4A and KIF5C in intellectual disability and synaptic function

    Journal of Medical Genetics 51:487–494.

    https://doi.org/10.1136/jmedgenet-2013-102182

    • PubMed
    • Google Scholar
71. 1. Wilson MH
    2. Holzbaur ELF

    (2015) Nesprins anchor kinesin-1 motors to the nucleus to drive nuclear distribution in muscle cells

    Development 142:218–228.

    https://doi.org/10.1242/dev.114769

    • PubMed
    • Google Scholar
72. 1. Wong YL
    2. Rice SE

    (2010) Kinesin’s light chains inhibit the head- and microtubule-binding activity of its tail

    PNAS 107:11781–11786.

    https://doi.org/10.1073/pnas.1005854107

    • Google Scholar
73. 1. Zhu H
    2. Lee HY
    3. Tong Y
    4. Hong BS
    5. Kim KP
    6. Shen Y
    7. Lim KJ
    8. Mackenzie F
    9. Tempel W
    10. Park HW

    (2012) Crystal structures of the tetratricopeptide repeat domains of kinesin light chains: insight into cargo recognition mechanisms

    PLOS ONE 7:e33943.

    https://doi.org/10.1371/journal.pone.0033943

    • PubMed
    • Google Scholar
74. 1. Zimmermann L
    2. Stephens A
    3. Nam S-Z
    4. Rau D
    5. Kübler J
    6. Lozajic M
    7. Gabler F
    8. Söding J
    9. Lupas AN
    10. Alva V

    (2018) A completely reimplemented MPI Bioinformatics Toolkit with A New HHpred Server at its Core

    Journal of Molecular Biology 430:2237–2243.

    https://doi.org/10.1016/j.jmb.2017.12.007

    • PubMed
    • Google Scholar

Author details

1. Zhenyu Tan

   1. Department of Biophysics, University of Michigan, Ann Arbor, United States
   2. Life Sciences Institute, University of Michigan, Ann Arbor, United States

   Contribution

   Conceptualization, Formal analysis, Investigation, Methodology, Writing - original draft, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-6491-5806

2. Yang Yue

   Department of Cell & Developmental Biology, University of Michigan, Ann Arbor, United States

   Contribution

   Formal analysis, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

3. Felipe Leprevost

   Department of Pathology, University of Michigan, Ann Arbor, United States

   Contribution

   Software, Formal analysis, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

4. Sarah Haynes

   Department of Pathology, University of Michigan, Ann Arbor, United States

   Contribution

   Software, Formal analysis, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

5. Venkatesha Basrur

   Department of Pathology, University of Michigan, Ann Arbor, United States

   Contribution

   Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

6. Alexey I Nesvizhskii

   1. Department of Pathology, University of Michigan, Ann Arbor, United States
   2. Department of Computational Medicine and Bioinformatics, University of Michigan, Ann Arbor, United States

   Contribution

   Supervision, Funding acquisition, Writing - review and editing

   Competing interests

   No competing interests declared

7. Kristen J Verhey

   Department of Cell & Developmental Biology, University of Michigan, Ann Arbor, United States

   Contribution

   Conceptualization, Formal analysis, Supervision, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-9329-4981

8. Michael A Cianfrocco

   1. Life Sciences Institute, University of Michigan, Ann Arbor, United States
   2. Department of Biological Chemistry, University of Michigan, Ann Arbor, United States

   Contribution

   Conceptualization, Data curation, Software, Formal analysis, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing

   For correspondence

   mcianfro@umich.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2067-4999

• 1,325

  views

• 96

  downloads

• 7

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.