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LIS1 promotes the formation of activated cytoplasmic dynein-1 complexes

Nature Cell Biology volume 22, pages 518–525 (2020)Cite this article

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Abstract

Cytoplasmic dynein-1 is a molecular motor that drives nearly all minus-end-directed microtubule-based transport in human cells, performing functions that range from retrograde axonal transport to mitotic spindle assembly1,2. Activated dynein complexes consist of one or two dynein dimers, the dynactin complex and an ‘activating adaptor’, and they show faster velocity when two dynein dimers are present3,4,5,6. Little is known about the assembly process of this massive ~4 MDa complex. Here, using purified recombinant human proteins, we uncover a role for the dynein-binding protein LIS1 in promoting the formation of activated dynein–dynactin complexes that contain two dynein dimers. Complexes activated by proteins representing three families of activating adaptors—BicD2, Hook3 and Ninl—all show enhanced motile properties in the presence of LIS1. Activated dynein complexes do not require sustained LIS1 binding for fast velocity. Using cryo-electron microscopy, we show that human LIS1 binds to dynein at two sites on the motor domain of dynein. Our research suggests that LIS1 binding at these sites functions in multiple stages of assembling the motile dynein–dynactin-activating adaptor complex.

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Fig. 1: LIS1 increases microtubule binding and velocity of activated dynein complexes.
Fig. 2: Human LIS1 binds to the human dynein motor domain at AAA3/4 and the stalk.
Fig. 3: LIS1 recruits a second dynein dimer to dynein–dynactin–BicD2-S complexes.
Fig. 4: LIS1 is not required to sustain the fast velocity of activated dynein complexes.
Fig. 5: LIS1 preferentially binds to open dynein and enhances the formation of complexes containing two open dynein dimers.

Data availability

No large datasets were generated during this study. Source data for Figs. 1–5 and Extended Data Figs. 1–4 are available with the paper. We encourage anyone who wishes to build on these studies or replicate them to contact the corresponding authors and we will share all plasmids used to generate the proteins used in these studies.

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Acknowledgements

We thank J. Christensen, E. Karasmanis, A. Kendrick, A. Roberts and J. Salogiannis for comments on the manuscript and discussions; staff at the Nikon Imaging Center at UC San Diego, where we collected data and received help with image analysis; staff at the UC San Diego cryo-EM facility, where cryo-EM data were collected; and staff at the Physics Computing Facility for IT support. S.L.R.-P. is supported by HHMI and NIH R01GM121772. Funding to S.L.R.-P. from the HHMI/Simons Faculty Scholars Program and R01GM107214 funded earlier parts of this research. A.E.L. is supported by R01GM107214; Z.M.H. by NSF graduate research fellowship DGE1144152; J.P.G. by the Molecular Biophysics Training Grant, NIH Grant T32 GM008326; R.W.B. is a Damon Runyon Fellow supported by the Damon Runyon Cancer Research Foundation DRG-#2285-17; and M.E.D. by NIH K99GM127757 and previously by a Jane Coffin Childs Memorial Fund Postdoctoral Fellowship 61-1552-T.

Author information

Author notes

  1. Zaw Min Htet

    Present address: Department of Molecular and Cell Biology, University of California Berkeley, Berkeley, CA, USA

  2. Richard W. Baker

    Present address: Department of Biochemistry and Biophysics, University of North Carolina, Chapel Hill, NC, USA

  3. Morgan E. DeSantis

    Present address: Department of Molecular, Cellular and Developmental Biology, University of Michigan, Ann Arbor, MI, USA

  4. These authors contributed equally: Zaw Min Htet, John P. Gillies.

Authors and Affiliations

  1. Department of Cellular and Molecular Medicine, University of California San Diego, La Jolla, CA, USA

    Zaw Min Htet, John P. Gillies, Richard W. Baker, Andres E. Leschziner, Morgan E. DeSantis & Samara L. Reck-Peterson

  2. Biophysics Graduate Program, Harvard Medical School, Boston, MA, USA

    Zaw Min Htet

  3. Division of Biological Sciences, Cell and Developmental Biology Section, University of California San Diego, La Jolla, CA, USA

    John P. Gillies & Samara L. Reck-Peterson

  4. Division of Biological Sciences, Molecular Biology Section, University of California San Diego, La Jolla, CA, USA

    Andres E. Leschziner

  5. Howard Hughes Medical Institute, Chevy Chase, MD, USA

    Samara L. Reck-Peterson

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Contributions

Z.M.H., J.P.G., M.E.D., A.E.L. and S.L.R.-P. designed the experiments. Z.M.H., J.P.G., M.E.D. and R.W.B. performed the experiments. Z.M.H., J.P.G., M.E.D., A.E.L. and S.L.R.-P. wrote the manuscript. All of the authors interpreted the data and reviewed and edited the manuscript.

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Correspondence to Morgan E. DeSantis or Samara L. Reck-Peterson.

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Extended data

Extended Data Fig. 1 Effect of Lis1 on the motility and microtubule binding of activated dynein complexes.

a, SDS-PAGE gel stained with Sypro Red of human dynein, dynactin and the activating adaptors BicD2-S (aa 25-398), BicD2-L (aa 1-598), Hook3 (aa 1-552), and Ninl (aa 1-702) used in this study. The dynein heavy chain was tagged with the SNAP tag, the dynactin subunit p62 with the HaloTag, and each activating adaptor with the HaloTag. The dynein light chains are too small to be seen on this low percentage gel. SDS-PAGE gels were run after all protein purifications. b, Example microscopy images for microtubule binding density data in the absence (white circles) or presence (black circles) of 300 nM Lis1 presented in Fig. 1d, e. Microtubules in magenta and dynein or activating adaptor foci in green. Scale bars are 10 µm. c, Example kymographs of dynein–dynactin–activating adaptor complexes in the absence (white circles) or presence (black circles) of 300 nM Lis1. Scale bars are 10 µm (x) and 20 sec (y). d, Percent processive runs of dynein–dynactin–activating adaptor complexes in standard motility buffer in the absence (white circles) or presence (black circles) of 300 nM Lis1. Statistical analysis was performed on data pooled from all replicates with χ2 test. e, Immunoblots of cell lysates from human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs, as well as either scramble siRNA or Lis1 siRNA 1 or 2. Blots were performed for each biorep with similar results. f, Peroxisome velocity in human U2OS cells with scrambled or Lis1 siRNA pool knockdown. The median and interquartile range are shown. At least 7 peroxisome motility events were measured per cell. g, Immunoblots of cell lysates from human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs and scramble or Lis1 siRNA pool. Two bio-replicates (1 and 2) are shown. An anti-V5 antibody detects BicD2-S-V5-FRB, an anti-Lis1 antibody assesses the efficiency of Lis1 knockdown, and an anti-actin antibody serves as a loading control for immunoblots shown in e and g. Statistical data and unprocessed gel and blot images are available as source data for Extended Data Fig. 1.

Source data

Extended Data Fig. 2 Characterization of the dynein binding interface and dimerization domain of Lis1.

a, Example SEC-MALS traces with Lis1 dimer (orange), dynein monomer (grey), and dynein monomer with Lis1 dimer (black). The intensity of the UV signal (solid line) and the molecular weight fit (dashed line) are shown. Dimeric Halo-tagged-Lis1 is expected to be 161.4 kDa and monomeric dynein is expected to be 380.4 kDa. In this experiment we observe Halo-tagged-Lis1 to be 157.6 kDa, monomeric dynein to be 489.5 kDa and the Lis1-dynein complex to be 700.1 kDa. The high apparent molecular weight of monomeric dynein may be due to a self-association species that appears as a shoulder in the UV trace. The experiment was repeated in triplicate yielding similar results, giving a stoichiometry of 1.2 ± 0.3 Lis1 dimers per dynein monomer. Based on this data we cannot rule out that some dynein monomers are bound to two Lis1 dimers (which has been reported to occur34), but our data suggest that most dynein monomers bind a single Lis1 dimer, and that Lis1 does not tether two dynein monomers. b, Single-molecule velocity of dynein–dynactin–BicD2-S complexes with increasing concentrations of Lis1. The median and interquartile range are shown. c, Single-molecule velocity of dynein–dynactin–Hook3 complexes in the presence of a higher ionic strength buffer in the absence (white circles) or presence (black circles) of 300 nM Lis1 or Lis1-5A. The data in the presence and absence of WT Lis1 was also presented in Fig. 1g. The median and interquartile range are shown. d, Example kymographs of dynein–dynactin–Hook3 complexes in a higher ionic strength buffer in the absence or presence of 300 nM Lis1 or Lis1-5A. Scale bars are 10 µm (x) and 20 sec (y). Data is quantified in Extended Data Fig. 2c. e, Example SEC-MALS trace of Lis1ΔN (orange). The intensity of the UV signal (solid line) and the molecular weight fit (dashed line) are shown. Monomeric Halo-tagged-Lis1ΔN is expected to be 71.5kDa. In this experiment we observe Halo-tagged-Lis1ΔN to have a monomer peak at 72.0 kDa and a dimer peak at 141.2 kDa. The experiment was repeated in triplicate yielding similar results. Statistical data is available as source data for Extended Data Fig. 2.

Source data

Extended Data Fig. 3 Quantification of the velocity of one-color and two-color activated dynein complexes in the presence of absence of Lis1.

a, Single-molecule velocity of dynein–dynactin–BicD2-S complexes in the absence (white circles) or presence (black circles) of 300 nM Lis1 with colocalized dynein (two color) or without observed colocalization (one color). The median and interquartile range are shown. Statistical data is available as source data for Extended Data Fig. 3.

Source data

Extended Data Fig. 4 Characterization of Lis1 binding to microtubules and activated dynein complexes.

a, Example microscopy images for imaging 50 nM TMR–Lis1 (magenta in merge) in the presence of microtubules (green in merge). Lis1 does not colocalize with microtubules. Scale bars are 10 µm. The experiment was repeated in triplicate yielding similar results. b, Single-molecule velocity of dynein–dynactin–BicD2-S complexes in the absence (white circles) or presence (black circles) of 50 nM TMR–Lis1 or TMR–Lis1-5A. The median and interquartile range are shown. c, Representative kymographs of Alexa647–dynein–dynactin–BicD2-S complexes in the presence of 50 nM TMR–Lis1-5A. Scale bars are 10 µm (x) and 20 sec (y). Statistical data is available as source data for Extended Data Fig. 4.

Source data

Supplementary information

Reporting Summary

Supplementary Video 1

Single-molecule imaging of dynein–dynactin–BicD2-S complexes. Single-molecule imaging of dynein–dynactin–BicD2-S complexes in the absence (top) or presence (bottom) of 300 nM LIS1 in standard motility buffer. TMR–dynein is shown in grey and the microtubule is shown in magenta. Frames were taken every 300 ms for 3 min. The video frame rate is 20 frames per s. Data are quantified in Fig. 1g.

Supplementary Video 2

Single-molecule imaging of dynein–dynactin–BicD2-L complexes. Single-molecule imaging of dynein–dynactin–BicD2-L complexes in the absence (top) or presence (bottom) of 300 nM LIS1 in standard motility buffer. TMR–dynein is shown in grey and the microtubule is shown in magenta. Frames were taken every 300 ms for 3 min. The video frame rate is 20 frames per s. Data are quantified in Fig. 1g.

Supplementary Video 3

Single-molecule imaging of dynein–dynactin–Hook3 complexes. Single-molecule imaging of dynein–dynactin–Hook3 complexes in the absence (top) or presence (bottom) of 300 nM LIS1 in standard motility buffer. TMR–dynein is shown in grey and the microtubule is shown in magenta. Frames were taken every 300 ms for 3 min. The video frame rate is 20 frames per s. Data are quantified in Fig. 1g.

Supplementary Video 4

Single-molecule imaging of dynein–dynactin–Ninl complexes. Single-molecule imaging of dynein–dynactin–Ninl complexes in the absence (top) or presence (bottom) of 300 nM LIS1 in standard motility buffer. TMR–dynein is shown in grey and the microtubule is shown in magenta. Frames were taken every 300 ms for 3 min. The video frame rate is 20 frames per s. Data are quantified in Fig. 1g.

Supplementary Video 5

Single-molecule imaging of dynein–dynactin–Hook3 complexes in higher-ionic-strength buffer. Single-molecule imaging of dynein–dynactin–Hook3 complexes in the absence (top) or presence (bottom) of 300 nM LIS1 in a higher-ionic-strength motility buffer. TMR–dynein is shown in grey and the microtubule is shown in magenta. Frames were taken every 300 ms for 3 min. Video frame rate is 20 frames per s. Data are quantified in Fig. 1h.

Supplementary Video 6

Peroxisome dynamics before the addition of rapalog with scrambled siRNA. Human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs and a scrambled siRNA. mEmerald dynamics are visualized before addition of rapalog. Frames were taken every 500 ms for 2 min. The video frame rate is 20 frames per s. Two biological replicates were collected yielding similar results.

Supplementary Video 7

Peroxisome dynamics after the addition of rapalog with scrambled siRNA. Human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs and a scrambled siRNA. mEmerald dynamics are visualized after addition of rapalogue. The orange arrows indicate examples of motile peroxisomes. Frames were taken every 500 ms for 2 min. The video frame rate is 20 frames per s. Data are quantified in Extended Data Fig. 1f.

Supplementary Video 8

Peroxisome dynamics before the addition of rapalog with PAFAH1B1 siRNA. Human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs and PAFAH1B1 siRNA pool. mEmerald dynamics are visualized before addition of rapalog. Frames were taken every 500 ms for 2 min. The video frame rate is 20 frames per s. Two biological replicates were collected yielding similar results.

Supplementary Video 9

Peroxisome dynamics after the addition of rapalog with PAFAH1B1 siRNA. Human U2OS cells co-transfected with PEX3-mEmerald-FKBP and BicD2-S-V5-FRB constructs and PAFAH1B1 siRNA pool. mEmerald dynamics are visualized after addition of rapalog. Orange arrows indicate examples of motile peroxisomes. Frames were taken every 500 ms for 2 min. The video frame rate is 20 frames per s. Data are quantified in Extended Data Fig. 1f.

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Htet, Z.M., Gillies, J.P., Baker, R.W. et al. LIS1 promotes the formation of activated cytoplasmic dynein-1 complexes. Nat Cell Biol 22, 518–525 (2020). https://doi.org/10.1038/s41556-020-0506-z

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