Advertisement
Journal Articles
The regulation of bidirectional mitochondrial transport is coordinated with axonal outgrowth
R.L. Morris, P.J. Hollenbeck
Journal of Cell Science 1993 104: 917-927;

Summary

Although small molecules such as ATP diffuse freely in the cytosol, many types of cells nonetheless position their mitochondria in regions of intense ATP consumption. We reasoned that in the highly elongated axonal processes of growing neurons in culture, the active growth cone would form a focus of ATP consumption so distant from the cell body as to require the positioning of mitochondria nearby via regulated axonal transport. To test this hypothesis, we quantified the distribution and transport behavior of mitochondria in live, aerobically respiring chick sympathetic neurons. We found that in the distal region of actively growing axons, the distribution of mitochondria was highly skewed toward the growth cone, with a sevenfold higher density in the region immediately adjacent to the growth cone than in the region 100 microns away. When axonal outgrowth was blocked by substratum-associated barriers or mild cytochalasin E treatment, the gradient of mitochondrial distribution collapsed as mitochondria exited retrogradely from the distal region, becoming uniformly distributed along the axon within one hour. Analysis of individual mitochondrial behaviors revealed that mitochondrial movement everywhere was bidirectional but balanced so that net transport was anterograde in growing axons and retrograde in blocked axons. This reversal in net transport derived from two separate modulations of mitochondrial movement. First, moving mitochondria underwent a transition to a persistently stationary state in the region of active growth cones that was reversed when growth cone activity was halted. Second, the fraction of time that mitochondria spent moving anterogradely was sharply reduced in non-growing axons. Together, these could account for the formation of gradients of mitochondria in growing axons and their dissipation when outgrowth was blocked. This regulated transport behavior was not dependent upon the ability of mitochondria to produce ATP. Our data indicate that mitochondria possess distinct motor activities for both directions of movement and that mitochondrial transport in axons is regulated by both recruitment between stationary and moving states, and direct regulation of the anterograde motor.

REFERENCES

    1. Bereiter-Hahn J. and
    2. Vöth M.
    (1983). Metabolic control of shape and structure of mitochondria in situ. Biol.Cell 47, 309–322
    OpenUrl
    1. Bradley T. J. and
    2. Satir P.
    (1979). Evidence of microfilament-associated mitochondrial movement. J. Supramol. Struct 12, 165–175
    OpenUrlCrossRefPubMedWeb of Science
    1. Brady S. T.,
    2. Lasek R. J. and
    3. Allen R. D.
    (1982). Fast axonal transport in extruded axoplasm from squid giant axon. Science 218, 1129–1131
    OpenUrlAbstract/FREE Full Text
    1. Brady S. T.,
    2. Lasek R. J.,
    3. Allen R. D.,
    4. Yin H. L. and
    5. Stossel T. P.
    (1984). Gelsolin inhibition of fast axonal transport indicates a requirement for actin microfilaments. Nature 310, 56–58
    OpenUrlCrossRefPubMed
    1. Bray J. J.,
    2. Kon C. M. and
    3. Breckenridge B. McL
    (1971). Reversed polarity of rapid axonal transport in chicken motoneurons. Brain Res 33, 560–564
    OpenUrlCrossRefPubMedWeb of Science
    1. Burton P. R. and
    2. Paige J. L.
    (1981). Polarity of axoplasmic microtubules in the olfactory nerve of the frog. Proc. Nat. Acad. Sci. USA 78, 3269–3273
    OpenUrlAbstract/FREE Full Text
    1. Chilcote T. J. and
    2. Johnson K. A.
    (1990). Phosphorylation of Tetrahymena 22 S dynein. J. Biol. Chem 265, 17257–17266
    OpenUrlAbstract/FREE Full Text
    1. Cohan C. S.,
    2. Conner J. A. and
    3. Kater S. B.
    (1987). Electrically and chemically mediated increases in intracellular calcium in neuronal growth cones. J. Neurosci 7, 3588–3599
    OpenUrlAbstract
    1. Cole D. G.,
    2. Cande W. Z.,
    3. Baskin R. J.,
    4. Skoufias D. A.,
    5. Hogan C. J. and
    6. Schloley J. M.
    (1992). Isolation of sea urchin kinesin-related protein using peptide antibodies. J. Cell Sci 101, 291–301
    OpenUrlAbstract/FREE Full Text
    1. Cooper J. A.
    (1987). Effects of cytochalasin and phalloidin on actin. J. Cell Biol 105, 1473–1478
    OpenUrlFREE Full Text
    1. Corthesy-Theulaz I.,
    2. Pauloin A. and
    3. Pfeffer S. R.
    (1992). Cytoplasmic dynein participates in the centrosomal localization of the Golgi complex. J. Cell Biol 118, 1333–1345
    OpenUrlAbstract/FREE Full Text
    1. Edmonds B. and
    2. Koenig E.
    (1987). Powering of bulk transport (varicosities) and differential sensitivities of directional transport in growing axons. Brain Res 406, 288–293
    OpenUrlCrossRefPubMedWeb of Science
    1. Farshori P. Q. and
    2. Goode D.
    (1991). Kinesin is phosphorylated during stimulation of insulin secretion and associates with secretory granules in HIT cells. J. Cell Biol 115, 390–.
    OpenUrl
    1. Forman D. S.,
    2. Lynch K. J. and
    3. Smith R. S.
    (1987). Organelle dynamics in lobster axons: anterograde, retrograde, and stationary mitochondria. Brain Res 412, 96–106
    OpenUrlCrossRefPubMedWeb of Science
    1. Frizell M.,
    2. Mclean W. G. and
    3. Sjöstrand J.
    (1976). Retrograde axonal transport of rapidly migrating labelled proteins and glycoproteins in regenerating peripheral nerves. J. Neurochem 27, 191–196
    OpenUrlCrossRefPubMed
    1. Grafstein B. and
    2. Forman D. S.
    (1980). Intracellular transport in neurons. Physiol. Rev. 60, 1167–1283
    OpenUrlFREE Full Text
    1. Hamasaki T.,
    2. Barkalow K.,
    3. Richmond J. and
    4. Satir P.
    (1991). cAMP-stimulated phosphorylation of an axonemal polypeptide that copurifies with a 22 S dynein arm regulates microtubule translocation velocity and swimming speed in Paramecium. Proc. Nat. Acad. Sci. USA 88, 7918–7922
    OpenUrlAbstract/FREE Full Text
    1. Heidemann S. R.,
    2. Landers J. M. and
    3. Hamborg M. A.
    (1981). Polarity orientation of axonal microtubules. J. Cell Biol 91, 661–665
    OpenUrlAbstract/FREE Full Text
    1. Heggeness M. H.,
    2. Simon M. and
    3. Singer S. J.
    (1978). Association of mitochondria with microtubules in cultured cells. Proc. Nat. Acad. Sci. USA 75, 3863–3866
    OpenUrlAbstract/FREE Full Text
    1. Hirokawa N.
    (1982). Cross-linker system between neurofilaments, microtubules, and membranous organelles in frog axons revealed by the quick-freeze, deep-etching method. J. Cell Biol 94, 129–142
    OpenUrlAbstract/FREE Full Text
    1. Hirokawa N.,
    2. Sato-Yoshitake R.,
    3. Kobayashi N.,
    4. Pfister K. K.,
    5. Bloom G. S. and
    6. Brady S. T.
    (1991). Kinesin associates with anterogradely transported membranous organelles in vivo. J. Cell Biol 114, 295–302
    OpenUrlAbstract/FREE Full Text
    1. Ho W. C.,
    2. Allen V. J.,
    3. Van Meer G.,
    4. Berger E. G. and
    5. Kreis T. E.
    (1989). Reclustering of scattered Golgi elements occurs along microtubules. Eur. J. Cell Biol 48, 250–263
    OpenUrlPubMedWeb of Science
    1. Hollenbeck P. J. and
    2. Bray D.
    (1987). Rapidly transported oganelles containing membrane and cytoskeletal components: their relation to axonal growth. J. Cell Biol 105, 2827–2835
    OpenUrlAbstract/FREE Full Text
    1. Hollenbeck P. J.,
    2. Bray D. and
    3. Adams R. J.
    (1985). Effects of the uncoupling agents FCCP and CCCP on the saltatory movements of cytoplasmic organelles. Cell Biol. Int. Rep 9, 193–199
    OpenUrlCrossRefPubMedWeb of Science
    1. Hollenbeck P. J. and
    2. Swanson J. A.
    (1990). Radial extension of macrophage tubular lysosomes supported by kinesin. Nature 346, 864–866
    OpenUrlCrossRefPubMedWeb of Science
    1. Hughes A.
    (1953). The growth of embryonic neurites. J. Anat 87, 150–162
    OpenUrlPubMed
    1. Johnson G. D.,
    2. Nogueira Araujo G. M. and
    3. de C.
    (1981). A simple method for reducing the fading of immunofluorescence during microscopy. J. Immunol. Meth 43, 349–350
    OpenUrlCrossRefPubMedWeb of Science
    1. Johnson L.V.,
    2. Walsh M. L.,
    3. Bockus B. J. and
    4. Chen L. B.
    (1981). Monitoring of relative mitochondrial membrane potential in living cells by fluorescence microscopy. J. Cell Biol 88, 526–535
    OpenUrlAbstract/FREE Full Text
    1. Kuhn-Nentwig L. and
    2. Kadenbach B.
    (1985). Isolation and properties of cytochrome C oxidase from rat liver and quantitation of immunological differences between isozymes from various rat tissues with subunit-specific antisera. Eur. J. Biochem 149, 147–158
    OpenUrlPubMedWeb of Science
    1. Kuznetsov S. A.,
    2. Langford G. M. and
    3. Weiss D. G.
    (1992). Actin-dependent organelle movement in squid axoplasm. Nature 356, 722–725
    OpenUrlCrossRefPubMedWeb of Science
    1. Lander A. D.,
    2. Fujii D.K.,
    3. Gospodarowicz D. and
    4. Reichardt L. F.
    (1982). Characterization of a factor that promotes neurite outgrowth: evidence linking activity to a heparin sulfate proteoglycan. J. Cell Biol 94, 574–585
    OpenUrlAbstract/FREE Full Text
    1. Lankford K. L.,
    2. DeMello F. G. and
    3. Klein W. L.
    (1988). D1 -type dopamine receptors inhibit growth cone motility in cultured retina neurons: evidence that neurotransmitters act as morphogenic growth regulators in the developing central nervous system. Proc. Nat. Acad. Sci. USA 85, 2839–2843
    OpenUrlAbstract/FREE Full Text
    1. Lankford K. L. and
    2. Letourneau P. C.
    (1991). Roles of actin filaments and three second-messenger systems in short-term regulation of chick dorsal root ganglion neurite outgrowth. Cell Motil. Cytoskel 20, 7–29
    OpenUrlCrossRefPubMedWeb of Science
    1. Lee C.,
    2. Ferguson M. and
    3. Chen L. B.
    (1989). Construction of the endoplasmic reticulum. J. Cell Biol 109, 2045–2055
    OpenUrlAbstract/FREE Full Text
    1. Leterrier J.-F.,
    2. Linden M. and
    3. Nelson B. D.
    (1990). How do microtubules interact in vitro with purified subcellular organelles?. Biochem. J 269, 556–558
    OpenUrlFREE Full Text
    1. Lin A.,
    2. Krockmalnic G. and
    3. Penman S.
    (1990). Imaging cytoskeleton-mitochondrial membrane attachments by embedment-free electron microscopy of saponin-extracted cells. Proc. Nat. Acad. Sci. USA 87, 8565–8569
    OpenUrlAbstract/FREE Full Text
    1. Luzikov V. N.,
    2. Zubatov A. S. and
    3. Rainina E. I.
    (1973). Formation and degradation of mitochondria in the cell. J. Bioenergetics 5, 129–149
    OpenUrl
    1. Luzikov V. N.,
    2. Zubatov A. S.,
    3. Rainina E. I. and
    4. Bakeyeva L. E.
    (1971). Degradation and restoration of mitochondria upon deaeration and subsequent aeration of aerobically grown Saccharomyces cerevisiae cells. Biochim. Biophys. Acta 245, 321–334
    OpenUrlPubMed
    1. Magrassi L.,
    2. Purves D. and
    3. Lichtman J.W.
    (1987). Fluorescent probes that stain living nerve terminals. J. Neurosci 7, 1207–1214
    OpenUrlAbstract
    1. Maness P. F.,
    2. Aubry M.,
    3. Shores C. G.,
    4. Frame L. and
    5. Pfenninger K. H.
    (1988). c-src gene product in developing rat brain is enriched in nerve growth cone membranes. Proc. Nat. Acad. Sci. USA 85, 5001–5005
    OpenUrlAbstract/FREE Full Text
    1. Mattson M. P.,
    2. Taylor-Hunter A. and
    3. Kater S. B.
    (1988). Neurite outgrowth in individual neurons of a neuronal population is differentially regulated by calcium and cAMP. J. Neurosci 8, 1704–1711
    OpenUrlAbstract
    1. Martz D.,
    2. Lasek R. J.,
    3. Brady S. T. and
    4. Allen R. D.
    (1984). Mitochondrial motility in axons: membranous organelles may interactwith the force generating system through multiple surface binding sites. Cell Motil 4, 89–101
    OpenUrlCrossRefPubMedWeb of Science
    1. McLean I. W. and
    2. Nakane P. K.
    (1974). Periodate-lysine-paraformaldehyde fixative: a new fixative for immunoelectron microscopy. J. Histochem. Cytochem 22, 1077–1083
    OpenUrlAbstract/FREE Full Text
    1. Miller R. H. and
    2. Lasek R. J.
    (1985). Cross-bridges mediate anterograde and retrograde vesicle transport along microtubules in squid axoplasm. J. Cell Biol 101, 2181–2193
    OpenUrlAbstract/FREE Full Text
    1. Paschal B. M.,
    2. Shpetner H. S. and
    3. Vallee R. B.
    (1987). MAP 1C is a microtubule-activated ATP-ase which translocates microtubules in vitro and has dynein-like properties. J. Cell Biol 105, 1273–1282
    OpenUrlAbstract/FREE Full Text
    1. Porter M. E.,
    2. Scholey J. M.,
    3. Stemple D. L.,
    4. Vigers G. P.,
    5. Vale R. D.,
    6. Sheetz M. P. and
    7. McIntosh J. R.
    (1987). Characterization of the microtubule movement produced by sea urchin egg kinesin. J. Biol. Chem 262, 2794–2802
    OpenUrlAbstract/FREE Full Text
    1. Price M. G. and
    2. Gomer R. H.
    (1989). Mitoskelin: a mitochondrial protein found in cytoskeletal preparations. Cell Motil 13, 274–287
    OpenUrl
    1. Rehder V. and
    2. Kater S. B.
    (1992). Regulation of neuronal growth cone filopodia by intracellular calcium. J. Neurosci 12, 3175–3186
    OpenUrlAbstract
    1. Reinsch S. S.,
    2. Mitchison T. J. and
    3. Kirschner M.
    (1991). Microtubule polymer assembly and transport during axonal elongation. J. Cell Biol 115, 365–379
    OpenUrlAbstract/FREE Full Text
    1. Rendon A.,
    2. Jung D. and
    3. Jancsik V.
    (1990). Interaction of microtubules and microtubule-associated proteins (MAPs) with rat brain mitochondria. Biochem. J 269, 555–556
    OpenUrlFREE Full Text
    1. Sahenk Z. and
    2. Brown A.
    (1991). Weak-base amines inhibit the anterograde-to-retrograde conversion of axonally transported vesicles in nerve terminals. J. Neurocytol 20, 365–375
    OpenUrlCrossRefPubMed
    1. Sahenk Z. and
    2. Lasek R. J.
    (1988). Inhibition of proteolysis blocks anterograde-retrograde conversion of axonally transported vesicles. Brain Res 460, 199–203
    OpenUrlCrossRefPubMedWeb of Science
    1. Sato-Yoshitake R.,
    2. Yorifuji H.,
    3. Inagaki M. and
    4. Hirokawa N.
    (1991). Phosphorylation of kinesin regulates its binding to synaptic vesicles. J. Cell Biol 115, 390–.
    OpenUrl
    1. Sawin K. E.,
    2. Mitchison T. J. and
    3. Wordeman L. G.
    (1992). Evidence for kinesin-related proteins in the mitotic apparatus using peptide antibodies. J. Cell Sci 101, 303–313
    OpenUrlAbstract/FREE Full Text
    1. Saxton W. M.,
    2. Porter M. E.,
    3. Cohn S. A.,
    4. Scholey J. M.,
    5. Raff E. C. and
    6. McIntosh J. R.
    (1988). Drosophila kinesin: characterization of microtubule motility and ATPase. Proc. Nat. Acad. Sci. USA 85, 1109–1113
    OpenUrlAbstract/FREE Full Text
    1. Schnapp B. and
    2. Reese T.S.
    ) (1982). Cytoplasmic structure in rapid-frozen axons. J. Cell Biol 94, 667–679
    OpenUrlAbstract/FREE Full Text
    1. Schnapp B. and
    2. Reese T.S.
    ) (1989). Dynein is the motor for retrograde axonal transport of organelles. Proc. Nat. Acad. Sci. USA 86, 1548–1552
    OpenUrlAbstract/FREE Full Text
    1. Schroer T. A. and
    2. Sheetz M. P.
    (1991). Functions of microtubule-based motors. Annu. Rev. Physiol 53, 629–652
    OpenUrlCrossRefPubMedWeb of Science
    1. Schroer T. A.,
    2. Steuer E. R. and
    3. Sheetz M. P.
    (1989). Cytoplasmic dynein is a minus end-directed motor for membranous organelles. Cell 56, 937–946
    OpenUrlCrossRefPubMedWeb of Science
    1. Smith R. S.
    (1988). Studies on the mechanism of the reversal of rapid organelle transport in myelinated axons of Xenopus laevis. Cell Motil 10, 296–308
    OpenUrlCrossRef
    1. Smith R. S. and
    2. Snyder R. E.
    (1991). Reversal of rapid axonal transport at a lesion: leupeptin inhibits reversed protein transport, but does not inhibit reversed organelle transport. Brain Res 552, 215–227
    OpenUrlCrossRefPubMed
    1. Stromer M. H. and
    2. Bendayan M.
    (1990). Immunocytochemical identification of cytoskeletal linkages to smooth muscle cell nuclei and mitochondria. Cell Motil 17, 11–18
    OpenUrl
    1. Summerhayes I. C.,
    2. Wong D. and
    3. Chen L. B.
    (1983). Effect of microtubules and intermediate filaments on mitochondrial distribution. J. Cell Sci 61, 87–105
    OpenUrlAbstract/FREE Full Text
    1. Takenaka T.,
    2. Kawakami T.,
    3. Hikawa N. and
    4. Gotoh H.
    (1990). Axoplasmic transport of mitochondria in cultured dorsal root ganglion cells. Brain Res 528, 285–290
    OpenUrlCrossRefPubMedWeb of Science
    1. Toyoshima I.,
    2. Yu H.,
    3. Steuer E. R. and
    4. Sheetz M. P.
    (1992). Kinectin, a major kinesin-binding protein on ER. J. Cell Biol 118, 1121–1132
    OpenUrlAbstract/FREE Full Text
    1. Tsukita S. and
    2. Ishikawa H.
    (1980). The movement of membranous organelles in axons. J. Cell Biol 84, 513–30
    OpenUrlAbstract/FREE Full Text
    1. Vale R. D.,
    2. Reese T. S. and
    3. Sheetz M. P.
    (1985). Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility. Cell 42, 39–50
    OpenUrlCrossRefPubMedWeb of Science
    1. Vale R. D.,
    2. Schnapp B. J.,
    3. Mitchison T.,
    4. Steuer E.,
    5. Reese T. S. and
    6. Morris R. L.
    P. J. Hollenbeck927Regulation of mitochondrial transportSheetz, M. P (1985). Different axoplasmic proteins generate movement in opposite directions along microtubules in vitro. Cell 43, 623–632
    OpenUrlCrossRefPubMedWeb of Science
    1. Vallee R. B. and
    2. Bloom G. S.
    (1991). Mechanisms of fast and slow axonal transport. Annu. Rev. Neurosci 14, 59–92
    OpenUrlCrossRefPubMedWeb of Science
    1. Van Blerkom J.
    (1991). Microtubule mediation of cytoplasmic and nuclear maturation during the early stages of resumed meiosis in cultured mouse oocytes. Proc. Nat. Acad. Sci. USA 88, 5031–5035
    OpenUrlAbstract/FREE Full Text
    1. Zielinski B. S.,
    2. Getchell M. L. and
    3. Getchell T. V.
    (1988). Ultrastructural characteristics of sustentacular cells in control and odorant-treated olfactory mucosae of the salamander. Anat. Rec 221, 769–779
    OpenUrlCrossRefPubMed
Back to top

 Download PDF

Email
Journal Articles
The regulation of bidirectional mitochondrial transport is coordinated with axonal outgrowth
R.L. Morris, P.J. Hollenbeck
Journal of Cell Science 1993 104: 917-927;
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
Citation Tools
Alerts

Article navigation

Other journals from The Company of Biologists

Advertisement

2020 at The Company of Biologists

Despite the challenges of 2020, we were able to bring a number of long-term projects and new ventures to fruition. While we look forward to a new year, join us as we reflect on the triumphs of the last 12 months.

Mole – The Corona Files

"This is not going to go away, 'like a miracle.' We have to do magic. And I know we can."

Mole continues to offer his wise words to researchers on how to manage during the COVID-19 pandemic.

Cell scientist to watch – Christine Faulkner

In an interview, Christine Faulkner talks about where her interest in plant science began, how she found the transition between Australia and the UK, and shares her thoughts on virtual conferences.

Read & Publish participation extends worldwide

“The clear advantages are rapid and efficient exposure and easy access to my article around the world. I believe it is great to have this publishing option in fast-growing fields in biomedical research.”

Dr Jaceques Behmoaras (Imperial College London) shares his experience of publishing Open Access as part of our growing Read & Publish initiative. We now have over 60 institutions in 12 countries taking part – find out more and view our full list of participating institutions.

JCS and COVID-19

For more information on measures Journal of Cell Science is taking to support the community during the COVID-19 pandemic, please see here.

If you have any questions or concerns, please do not hestiate to contact the Editorial Office.