Evolution of the Molecules Coupling mRNA Transport with Translational Control in Metazoans

  • Paula Vazquez-PianzolaEmail author
  • Beat Suter
  • Greco HernándezEmail author


Restricting proteins to specific subcellular regions is fundamental for various cellular processes. Such compartmentalization seems particularly important in large eukaryotic cells, and, accordingly, localization processes have been characterized best during embryogenesis, oogenesis, and in neuronal cells. A key mechanism underlying this process is the transport of mRNAs by molecular motors. Equally relevant is the translational control of the cargo mRNA, and this points to the importance of molecules that couple transport of mRNAs to translational control. In this chapter, we review recent discoveries sheading light on the evolution of one of the best-characterized machineries that couples transport and translation of mRNAs in metazoans, namely the Drosophila Bic-D/Egl/Dyn RNA localization machinery.


Bicaudal-D Egalitarian Dynein Dynactin Kinesin Cytoskeleton Mrna transport Translational control 



P. V.-P. and B.S. were supported by the Swiss National Science Foundation and the Canton of Bern. G.H. was supported by the National Institute of Cancer (INCan), Mexico, and the National Council of Science and Technology (CONACyT, grant no. 168154 to G.H.), Mexico.


  1. 1.
    Jekely G. Origin and evolution of self-organizing cytoskeleton in the network of eukaryotic organelles. In: Keeling PJ, Koonin EV, editors. The origin and evolution of eukaryotes. Cold Spring Harbor, New York, U.S.A.: Cold Spring Harbor Laboratory Press; 2014. p. 199–218.Google Scholar
  2. 2.
    Vale RD. The molecular motor toolbox for intracellular transport. Cell. 2003;112:467–80.CrossRefPubMedGoogle Scholar
  3. 3.
    Wickstead B, Gull K, Richards TA. Patterns of kinesin evolution reveal a complex ancestral eukaryote with a multifunctional cytoskeleton. BMC Evol Biol. 2010;10:110.CrossRefPubMedPubMedCentralGoogle Scholar
  4. 4.
    Wickstead B, Gull K. The evolution of the cytoskeleton. J Cell Biol. 2011;194:513–25.CrossRefPubMedPubMedCentralGoogle Scholar
  5. 5.
    Koonin EV, Yutin N. The dispersed archaeal eukaryome and the complex archaeal ancestor of eukaryotes. Cold Spring Harb Perspect Biol. 2014;6:a016188.CrossRefPubMedPubMedCentralGoogle Scholar
  6. 6.
    Williams TA, Foster PG, Cox CJ, Embley M. An archaeal origin of eukaryotes supports only two primary domains of life. Nature. 2013;504:231–6.CrossRefPubMedGoogle Scholar
  7. 7.
    Koumandou VL, Wickstead B, Ginger ML, van der Giezen M, Dacks JB, Field MC. Molecular paleontology and complexity in the last eukaryotic common ancestor. Crit Rev Biochem Mol Biol. 2013;48:373–96.CrossRefPubMedPubMedCentralGoogle Scholar
  8. 8.
    Gibbons BH, Asai DJ, Tang WJ, Hays TS, Gibbons IR. Phylogeny and expression of axonemal and cytoplasmic dynein genes in sea urchins. Mol Biol Cell. 1994;5:57–70.CrossRefPubMedPubMedCentralGoogle Scholar
  9. 9.
    Kamal A, Goldstein LSB. Principles of cargo attachment to cytoplasmic motor proteins. Curr Opin Cell Biol. 2002;14:63–8.CrossRefPubMedGoogle Scholar
  10. 10.
    Karcher RL, Deacon SW, Geldfand VI. Motor–cargo interactions: the key to transport specificity. Trends Cell Biol. 2002;12:21–7.CrossRefPubMedGoogle Scholar
  11. 11.
    Czaplinski K, Singer R. Pathways for mRNA localization in the cytoplasm. Trends Biochem Sci. 2006;31:687–93.CrossRefPubMedGoogle Scholar
  12. 12.
    Kong J, Lasko P. Translational control in cellular and developmental processes. Nat Rev Genet. 2012;13:383–94.CrossRefPubMedGoogle Scholar
  13. 13.
    Vazquez-Pianzola P, Suter B. Conservation of the RNA transport machineries and their coupling to translation control across eukaryotes. Comp Funct Genom. 2012;2012:287852.CrossRefGoogle Scholar
  14. 14.
    Rodriguez AJ, Czaplinski K, Condeelis JS, Singer RH. Mechanisms and cellular roles of local protein synthesis in mammalian cells. Curr Opin Cell Biol. 2008;20:144–9.CrossRefPubMedPubMedCentralGoogle Scholar
  15. 15.
    Weis BL, Schleiff E, Zerges W. Protein targeting to subcellular organelles via MRNA localization. Biochim Biophys Acta. 2013;1833:260–73.CrossRefPubMedGoogle Scholar
  16. 16.
    Lasko P. Translational control during early development. In: Hershey JWB, editors. Progress in Molecular Biology and Translational Science vol 90. Burlington: Academic Press; 2009. p. 211–254.Google Scholar
  17. 17.
    Richter JD, Lasko P. Translational control in oocyte development. Cold Spring Harb Perspect Biol. 2011;3:a002758.CrossRefPubMedPubMedCentralGoogle Scholar
  18. 18.
    McLaughlin JM, Bratu DP. Drosophila melanogaster oogenesis: an overview. Methods Mol Biol. 2015;1328:1–20.CrossRefPubMedGoogle Scholar
  19. 19.
    King ML, Messitt TJ, Mowry KL. Putting RNAs in the right place at the right time: RNA localization in the frog oocyte. Biol Cell. 2005;97:19–33.CrossRefPubMedGoogle Scholar
  20. 20.
    Lécuyer E, Yoshida H, Parthasarathy N, Alm C, Babak T, Cerovina T, Hughes TR, Tomancak P, Krause HM. Global analysis of mRNA localization reveals a prominent role in organizing cellular architecture and function. Cell. 2007;131:174–87.CrossRefPubMedGoogle Scholar
  21. 21.
    Kumano G. Polarizing animal cells via mRNA localization in oogenesis and early development. Dev Growth Differ. 2011;54:1–18.CrossRefPubMedGoogle Scholar
  22. 22.
    Jambor H, Surendranath V, Kalinka AT, Mejstrik P, Saalfeld S, Tomancak P. Systematic imaging reveals features and changing localization of mRNAs in Drosophila development. eLife. 2015;4.Google Scholar
  23. 23.
    Taliaferro JM, Wang ET, Burge CB. Genomic analysis of RNA localization. RNA Biol. 2014;11:1040–50.CrossRefPubMedPubMedCentralGoogle Scholar
  24. 24.
    Lécuyer E, Yoshida H, Krause HM. Global implications of mRNA localization pathways in cellular organization. Curr Opin Cell Biol. 2009;21:409–15.CrossRefPubMedGoogle Scholar
  25. 25.
    Mardakheh FK, Paul A, Kümper S, Sadok A, Paterson H, Mccarthy A, Yuan Y, Marshall CJ. Global analysis of mRNA, translation, and protein localization: local translation is a key regulator of cell protrusions. Dev Cell. 2015;35:344–57.CrossRefPubMedPubMedCentralGoogle Scholar
  26. 26.
    Condeelis J, Singer RH. How and why does beta-actin mRNA target? Biol Cell. 2005;97:97–110.CrossRefPubMedGoogle Scholar
  27. 27.
    Eom T, Antar LN, Singer RH, Bassell GJ. Localization of a beta-actin messenger ribonucleoprotein complex with zipcode-binding protein modulates the density of dendritic filopodia and filopodial synapses. J Neurosci. 2003;23:10433–44.PubMedGoogle Scholar
  28. 28.
    Zhang HL, Eom T, Oleynikov Y, Shenoy SM, Liebelt DA, Dictenberg JB, Singer RH, Bassell GJ. Neurotrophin-induced transport of a beta-actin mRNP complex increases beta-actin levels and stimulates growth cone motility. Neuron. 2001;31:261–75.CrossRefPubMedGoogle Scholar
  29. 29.
    Leung KM, van Horck FP, Lin AC, Allison R, Standart N, Holt CE. Asymmetrical beta-actin mRNA translation in growth cones mediates attractive turning to netrin-1. Nat Neurosci. 2006;9:1247–56.CrossRefPubMedPubMedCentralGoogle Scholar
  30. 30.
    Donnelly CJ, Fainzilber M, Twiss JL. Subcellular communication through RNA transport and localized protein synthesis. Traffic. 2010;11:1498–505.CrossRefPubMedPubMedCentralGoogle Scholar
  31. 31.
    Hüttelmaier S, Zenklusen D, Lederer M, Dictenberg J, Lorenz M, Meng X, Bassell GJ, Condeelis J, Singer RH. Spatial regulation of beta-actin translation by Src-dependent phosphorylation of ZBP1. Nature. 2005;438:512–5.CrossRefPubMedGoogle Scholar
  32. 32.
    Ishizuka A, Siomi MC, Siomi H. A Drosophila fragile X protein interacts with components of RNAi and ribosomal proteins. Genes Dev. 2002;16:2497–508.CrossRefPubMedPubMedCentralGoogle Scholar
  33. 33.
    Chen E, Sharma MR, Shi X, Agrawal RK, Joseph S. Fragile X mental retardation protein regulates translation by binding directly to the ribosome. Mol Cell. 2014;54:407–17.CrossRefPubMedPubMedCentralGoogle Scholar
  34. 34.
    Bechara EG, Didiot MC, Melko M, Davidovic L, Bensaid M, Martin P, Castets M, Pognonec P, Khandjian EW, Moine H. A novel function for fragile X mental retardation protein in translational activation. PLoS Biol. 2009;7:e16.CrossRefPubMedGoogle Scholar
  35. 35.
    Dictenberg JB, Swanger SA, Antar LN, Singer RH, Bassell GJ. A direct role for FMRP in activity-dependent dendritic mRNA transport links filopodial-spine morphogenesis to fragile X syndrome. Dev Cell. 2008;14:926–39.CrossRefPubMedPubMedCentralGoogle Scholar
  36. 36.
    Bassell GJ, Warren ST. Fragile X syndrome: loss of local mRNA regulation alters synaptic development and function. Neuron. 2008;60:201–14.CrossRefPubMedPubMedCentralGoogle Scholar
  37. 37.
    Huber F, Boire A, López MP, Koenderink GH. Cytoskeletal crosstalk: when three different personalities team up. Curr Opin Cell Biol. 2015;32:39–47.CrossRefPubMedGoogle Scholar
  38. 38.
    Alfaro-Aco R, Petry S. Building the microtubule cytoskeleton piece by piece. J Biol Chem. 2015;290:17154–62.CrossRefPubMedPubMedCentralGoogle Scholar
  39. 39.
    De la Cruz EM, Gardel ML. Actin mechanics and fragmentation. J Biol Chem. 2015;290:17137–44.CrossRefGoogle Scholar
  40. 40.
    Lowery J, Kuczmarski ER, Herrmann H, Goldman RD. Intermediate filaments play a pivotal role in regulating cell architecture and function. J Biol Chem. 2015;290:17145–53.CrossRefPubMedPubMedCentralGoogle Scholar
  41. 41.
    Shih YL, Rothfield L. The bacterial cytoskeleton. Microbiol Mol Biol Rev. 2006;70:729–54.CrossRefPubMedPubMedCentralGoogle Scholar
  42. 42.
    Löwe J, Amos LA. Evolution of cytomotive filaments: the cytoskeleton from prokaryotes to eukaryotes. Int J Biochem Cell Biol. 2009;41:323–9.CrossRefPubMedGoogle Scholar
  43. 43.
    Koonin EV. Origin of eukaryotes from within archaea, archaeal eukaryome and bursts of gene gain: eukaryogenesis just made easier? Philos Trans R Soc Lond B Biol Sci. 2015;370:20140333.CrossRefPubMedPubMedCentralGoogle Scholar
  44. 44.
    Makarova KS, Yutin N, Bell SD, Koonin EV. Evolution of diverse cell division and vesicle formation systems in Archaea. Nat Rev Microbiol. 2010;8:731–41.CrossRefPubMedPubMedCentralGoogle Scholar
  45. 45.
    Ettema TJ, Lindås AC, Bernander R. An actin-based cytoskeleon in Archaea. Mol Microbiol. 2011;80:1052–6.CrossRefPubMedGoogle Scholar
  46. 46.
    Wickstead B, Gull K. Dyneins across eukaryotes: a comparative genomic analysis. Traffic. 2007;8:1708–21.CrossRefPubMedPubMedCentralGoogle Scholar
  47. 47.
    Dienstbier M, Li X. Bicaudal-D and its role in cargo sorting by microtubule-based motors. Biochem Soc Trans. 2009;37:1066–71.CrossRefPubMedGoogle Scholar
  48. 48.
    Bullock SL. Translocation of mRNAs by molecular motors: think complex. Semin Cell Dev Biol. 2007;18:194–201.CrossRefPubMedGoogle Scholar
  49. 49.
    Claußen M, Suter B. BicD-dependent localization processes: from Drosophilia development to human cell biology. Ann Anat. 2005;187:539–53.CrossRefPubMedGoogle Scholar
  50. 50.
    Dienstbier M, Boehl F, Li X, Bullock SL. Egalitarian is a selective RNA-binding protein linking mRNA localization signals to the dynein motor. Genes Dev. 2009;23:1546–58.CrossRefPubMedPubMedCentralGoogle Scholar
  51. 51.
    Ran B, Bopp R, Suter B. Null alleles reveal novel requirement for Bic-D during Drosophila oogenesis and zygotic development. Development. 1994;120:1233–42.PubMedGoogle Scholar
  52. 52.
    Suter B, Steward R. Requirement for phosphorylation and localization of the Bicaudal-D protein in Drosophila oocyte differentiation. Cell. 1991;67:917–26.CrossRefPubMedGoogle Scholar
  53. 53.
    Schüpbach T, Wieschaus E. Female sterile mutations on the second chromosome of Drosophila melanogaster: II Mutations blocking oogenesis or altering egg morphology. Genetics. 1991;129:1119–36.PubMedPubMedCentralGoogle Scholar
  54. 54.
    Theurkauf WE, Alberts BM, Jan YN, Jongens TA. A central role for microtubules in the differentiation of Drosophila oocytes. Development. 1993;118:1169–80.PubMedGoogle Scholar
  55. 55.
    Clark A, Meignin C, Davis I. A Dynein-dependent shortcut rapidly delivers axis determination transcripts into the Drosophila oocyte. Development. 2007;134:1955–65.CrossRefPubMedPubMedCentralGoogle Scholar
  56. 56.
    Hughes JR, Bullock SL, Ish-Horowicz D. Inscuteable mRNA localization is dynein-dependent and regulates apicobasal polarity and spindle length in Drosophila neuroblasts. Curr Biol. 2004;14:1950–6.CrossRefPubMedGoogle Scholar
  57. 57.
    Bullock SL, Ish-Horowicz D. Conserved signals and machinery for RNA transport in Drosophila oogenesis and embryogenesis. Nature. 2001;414:611–6.CrossRefPubMedGoogle Scholar
  58. 58.
    Bullock SL, Ringel I, Ish-Horowicz D, Lukavsky PJ. A-form RNA helices are required for cytoplasmic mRNA transport in Drosophila. Nat Struct Mol Biol. 2010;17:703–9.CrossRefPubMedPubMedCentralGoogle Scholar
  59. 59.
    Jambor H, Mueller S, Bullock SL, Ephrussi A. A stem-loop structure directs oskar mRNA to microtubule minus ends. RNA. 2014;20:429–39.CrossRefPubMedPubMedCentralGoogle Scholar
  60. 60.
    Hachet O, Ephrussi A. Splicing of oskar RNA in the nucleus is coupled to its cytoplasmic localization. Nature. 2004;428:959–63.CrossRefPubMedGoogle Scholar
  61. 61.
    Jambor H, Brunel C, Ephrussi A. Dimerization of oskar 3′ UTRs promotes hitchhiking for RNA localization in the Drosophila oocyte. RNA. 2011;17:2049–57.CrossRefPubMedPubMedCentralGoogle Scholar
  62. 62.
    Tanaka T, Kato Y, Matsuda K, Hanyu-Nakamura K, Nakamura A. Drosophila Mon2 couples Oskar induced endocytosis with actin remodeling for cortical anchorage of the germ plasm. Development. 2011;138:2523–32.CrossRefPubMedGoogle Scholar
  63. 63.
    Vanzo N, Oprins A, Xanthakis D, Ephrussi A, Rabouille C. Stimulation of endocytosis and actin dynamics by Oskar polarizes the Drosophila oocyte. Dev Cell. 2007;12:543–55.CrossRefPubMedGoogle Scholar
  64. 64.
    Zimyanin VL. In vivo imaging of oskar mRNA transport reveals the mechanism of posterior localization. Cell. 2008;134:843–53.CrossRefPubMedPubMedCentralGoogle Scholar
  65. 65.
    Krauss J. Lopez de Quinto S, Nusslein-Volhard C, Ephrussi A. Myosin-V regulates oskar mRNA localization in the Drosophila oocyte. Curr Biol. 2009;19:1058–63.CrossRefPubMedGoogle Scholar
  66. 66.
    Cook HA, Koppetsch BS, Wu J, Theurkauf WE. The Drosophila SDE3 homolog armitage is required for oskar mRNA silencing and embryonic axis specification. Cell. 2004;116:817–29.CrossRefPubMedGoogle Scholar
  67. 67.
    Pane A, Wehr K, Schüpbach T. zucchini and squash encode two putative nucleases required for rasiRNA production in the Drosophila germline. Dev Cell. 2007;12:851–62.CrossRefPubMedPubMedCentralGoogle Scholar
  68. 68.
    Lim AK, Kai T. Unique germ-line organelle, nuage, functions to repress selfish genetic elements in Drosophila melanogaster. Proc Natl Acad Sci USA. 2007;104:6714–9.CrossRefPubMedPubMedCentralGoogle Scholar
  69. 69.
    Nakamura A, Amikura R, Hanyu K, Kobayashi S. Me31B silences translation of oocyte-localizing RNAs through the formation of cytoplasmic RNP complex during Drosophila oogenesis. Development. 2001;128:3233–42.PubMedGoogle Scholar
  70. 70.
    Besse F. Lopez de Quinto S, Marchand V, Trucco A, Ephrussi A. Drosophila PTB promotes formation of high-order RNP particles and represses oskar translation. Genes Dev. 2009;23:195–207.CrossRefPubMedPubMedCentralGoogle Scholar
  71. 71.
    Nakamura A, Sato K, Hanyu-Nakamura K. Drosophila cup is an eIF4E binding protein that associates with Bruno and regulates oskar mRNA translation in oogenesis. Dev Cell. 2004;6:69–78.CrossRefPubMedGoogle Scholar
  72. 72.
    Chekulaeva M, Hentze MW, Ephrussi A. Bruno acts as a dual repressor of oskar translation, promoting mRNA oligomerization and formation of silencing particles. Cell. 2006;124:521–33.CrossRefPubMedGoogle Scholar
  73. 73.
    Yano T, Lopez de Quinto S, Matsui Y, Shevchenko A, Ephrussi A. Hrp48, a Drosophila hnRNPA/B homolog, binds and regulates translation of oskar mRNA. Dev Cell. 2004;6:637–648.Google Scholar
  74. 74.
    Clouse KN, Ferguson SB, Schupbach T. Squid, Cup, and PABP55B function together to regulate gurken translation in Drosophila. Dev Biol. 2008;313:713–24.CrossRefPubMedGoogle Scholar
  75. 75.
    Vazquez-Pianzola P, Urlaub H, Suter B. Pabp binds to the osk 3′UTR and specifically contributes to osk mRNA stability and oocyte accumulation. Dev Biol. 2011;357:404–18.CrossRefPubMedGoogle Scholar
  76. 76.
    Olesnicky EC, Brent AE, Tonnes L, Walker M, Pultz MA, Leaf D, Desplan C. A caudal mRNA gradient controls posterior development in the wasp Nasonia. Development. 2006;133:3973–82.CrossRefPubMedGoogle Scholar
  77. 77.
    Kardon JR, Vale RD. Regulators of the cytoplasmic dynein motor. Nat Rev Mol Cell Biol. 2009;10:854–65.CrossRefPubMedPubMedCentralGoogle Scholar
  78. 78.
    Vazquez-Pianzola P, Adam J, Haldemann D, Hain D, Urlaub H, Suter B. Clathrin heavy chain plays multiple roles in polarizing the Drosophila oocyte downstream of Bic-D. Development. 2014;141:1915–26.CrossRefPubMedGoogle Scholar
  79. 79.
    Aguirre-Chen C, Bulow HE, Kaprielian ZC. elegans bicd-1, homolog of the Drosophila dynein accessory factor Bicaudal D, regulates the branching of PVD sensory neuron dendrites. Development. 2011;138:507–18.CrossRefPubMedPubMedCentralGoogle Scholar
  80. 80.
    Fridolfsson HN, Starr DA. Kinesin-1 and dynein at the nuclear envelope mediate the bidirectional migrations of nuclei. J Cell Biol. 2010;191:115–28.CrossRefPubMedPubMedCentralGoogle Scholar
  81. 81.
    Jaarsma D, van den Berg R, Wulf PS, van Erp S, Keijzer N, Schlager MA, de Graaff E, De Zeeuw CI, Pasterkamp RJ, Akhmanova A, Hoogenraad CC. A role for Bicaudal-D2 in radial cerebellar granule cell migration. Nat Commun. 2014;5:3411.PubMedGoogle Scholar
  82. 82.
    Terenzio M, Golding M, Russell MR, Wicher KB, Rosewell I, Spencer-Dene B, Ish-Horowicz D, Schiavo G. Bicaudal-D1 regulates the intracellular sorting and signalling of neurotrophin receptors. EMBO J. 2014;33:1582–98.CrossRefPubMedPubMedCentralGoogle Scholar
  83. 83.
    Lipka J, Kuijpers M, Jaworski J, Hoogenraad CC. Mutations in cytoplasmic dynein and its regulators cause malformations of cortical development and neurodegenerative diseases. Biochem Soc Trans. 2013;41:1605–12.CrossRefPubMedGoogle Scholar
  84. 84.
    Neveling K, Martinez-Carrera LA, Holker I, Heister A, Verrips A, Hosseini-Barkooie SM, Gilissen C, Vermeer S, Pennings M, Meijer R, te Riele M, Frijns CJ, Suchowersky O, MacLaren L, Rudnik-Schoneborn S, Sinke RJ, Zerres K, Lowry RB, Lemmink HH, Garbes L, Veltman JA, Schelhaas HJ, Scheffer H, Wirth B. Mutations in BICD2, which encodes a golgin and important motor adaptor, cause congenital autosomal-dominant spinal muscular atrophy. Am J Hum Genet. 2013;92:946–54.CrossRefPubMedPubMedCentralGoogle Scholar
  85. 85.
    Peeters K, Litvinenko I, Asselbergh B, Almeida-Souza L, Chamova T, Geuens T, Ydens E, Zimon M, Irobi J, De Vriendt E, De Winter V, Ooms T, Timmerman V, Tournev I, Jordanova A. Molecular defects in the motor adaptor BICD2 cause proximal spinal muscular atrophy with autosomal-dominant inheritance. Am J Hum Genet. 2013;92:955–64.CrossRefPubMedPubMedCentralGoogle Scholar
  86. 86.
    Hoogenraad CC, Akhmanova A, Howell SA, Dortland BR, De Zeeuw CI, Willemsen R, Visser P, Grosveld F, Galjart N. Mammalian Golgi-associated Bicaudal-D2 functions in the dynein-dynactin pathway by interacting with these complexes. EMBO J. 2001;20:4041–54.CrossRefPubMedPubMedCentralGoogle Scholar
  87. 87.
    Kriventseva EV, Tegenfeldt F, Petty TJ, Waterhouse RM, Simão FA, Pozdnyakov IA, Ioannidis P, Zdobnov EM. OrthoDB v8: update of the hierarchical catalog of orthologs and the underlying free software. Nucleic Acids Res. 2015;43:D250–6.CrossRefPubMedGoogle Scholar
  88. 88.
    Waterhouse RM, Tegenfeldt F, Li J, Zdobnov EM, Kriventseva EV. OrthoDB: a hierarchical catalog of animal, fungal and bacterial orthologs. Nucleic Acids Res. 2013;41:D358–65.CrossRefPubMedGoogle Scholar
  89. 89.
    Waterhouse RM, Zdobnov EM, Tegenfeldt F, Li J, Kriventseva EV. OrthoDB: the hierarchical catalog of eukaryotic orthologs in 2011. Nucleic Acids Res. 2011;39:D283–8.CrossRefPubMedGoogle Scholar
  90. 90.
    Schlager MA, Kapitein LC, Grigoriev I, Burzynski GM, Wulf PS, Keijzer N, de Graaff E, Fukuda M, Shepherd IT, Akhmanova A, Hoogenraad CC. Pericentrosomal targeting of Rab6 secretory vesicles by Bicaudal-D-related protein 1 (BICDR-1) regulates neuritogenesis. EMBO J. 2010;29:1637–51.CrossRefPubMedPubMedCentralGoogle Scholar
  91. 91.
    Preechaphol R, Klinbunga S, Khamnamtong B, Menasveta P. Isolation and characterization of genes functionally involved in ovarian development of the giant tiger shrimp Penaeus monodon by suppression subtractive hybridization (SSH). Genet Mol Biol. 2010;33:676–85.CrossRefPubMedPubMedCentralGoogle Scholar
  92. 92.
    Bianco A, Dienstbier M, Salter HK, Gatto G, Bullock SL. Bicaudal-D regulates fragile X mental retardation protein levels, motility, and function during neuronal morphogenesis. Curr Biol. 2010;20:1487–92.CrossRefPubMedPubMedCentralGoogle Scholar
  93. 93.
    Guerra N, Vega-Sendino M, Pérez-Morgado MI, Ramos E, Soto M, Gonzalez VM, Martín ME. Identification and functional characterization of a poly(A)-binding protein from Leishmania infantum (LiPABP). FEBS Lett. 2011;585:193–8.CrossRefPubMedGoogle Scholar
  94. 94.
    Burgess HM, Gray NK. mRNA-specific regulation of translation by poly(A)-binding proteins. Biochem Soc Trans. 2010;38:1517–22.CrossRefPubMedGoogle Scholar
  95. 95.
    Mangus DA, Evans MC, Jacobson A. Poly(A)-binding proteins: multifunctional scaffolds for post-transcriptional control of gene expression. Genom Biol. 2003;4:223.CrossRefGoogle Scholar
  96. 96.
    Gallie DR, Liu R. Phylogenetic analysis reveals dynamic evolution of the poly(A)-binding protein gene family in plants. BMC Evol Biol. 2014;14:238.CrossRefPubMedPubMedCentralGoogle Scholar
  97. 97.
    Hernández G, Altmann M, Lasko P. Origins and evolution of the mechanisms regulating translation initiation in eukaryotes. Trends Biochem Sci. 2010;35:63–73.CrossRefPubMedGoogle Scholar
  98. 98.
    Jimenez-Lopez D, Bravo J, Guzman P. Evolutionary history exposes radical diversification among classes of interaction partners of the MLLE domain of plant poly(A)-binding proteins. BMC Evol Biol. 2015;15:195.CrossRefPubMedPubMedCentralGoogle Scholar
  99. 99.
    Arn EA, Cha BJ, Theurkauf WE, Macdonald PM. Recognition of a bicoid mRNA localization signal by a protein complex containing Swallow, Nod, and RNA binding proteins. Dev Cell. 2003;4:41–51.CrossRefPubMedGoogle Scholar
  100. 100.
    Mohr E, Fuhrmann C, Richter D. VP-RBP, a protein enriched in brain tissue, specifically interacts with the dendritic localizer sequence of rat vasopressin mRNA. Eur J Neurosci. 2001;13:1107–12.CrossRefPubMedGoogle Scholar
  101. 101.
    Mohr E, Prakash N, Vieluf K, Fuhrmann C, Buck F, Richter D. Vasopressin mRNA localization in nerve cells: characterization of cis-acting elements and trans-acting factors. Proc Natl Acad Sci USA. 2001;98:7072–9.CrossRefPubMedPubMedCentralGoogle Scholar
  102. 102.
    Mohr E, Richter D. Subcellular vasopressin mRNA trafficking and local translation in dendrites. J Neuroendocrinol. 2004;16:333–9.CrossRefPubMedGoogle Scholar
  103. 103.
    Coffee RL, Tessier CR, Woodruff EA, Broadie K. Fragile X mental retardation protein has a unique, evolutionarily conserved neuronal function not shared with FXR1P or FXR2P. Dis Model Mech. 2010;3:471–85.CrossRefPubMedPubMedCentralGoogle Scholar
  104. 104.
    Tucker B, Richards R, Lardelli M. Expression of three zebrafish orthologs of human FMR1-related genes and their phylogenetic relationships. Dev Genes Evol. 2004;214:567–74.CrossRefPubMedGoogle Scholar
  105. 105.
    Kirkpatrick LL, McIlwain KA, Nelson DL. Comparative genomic sequence analysis of the FXR gene family: FMR1, FXR1, and FXR2. Genomics. 2001;78:169–77.CrossRefPubMedGoogle Scholar
  106. 106.
    Munro TP, Kwon S, Schnapp BJ. St Johnston D. A repeated IMP-binding motif controls oskar mRNA translation and anchoring independently of Drosophila melanogaster IMP. J Cell Biol. 2006;172:577–88.CrossRefPubMedPubMedCentralGoogle Scholar
  107. 107.
    Geng C, Macdonald PM. Imp associates with squid and Hrp48 and contributes to localized expression of gurken in the oocyte. Mol Cell Biol. 2006;26:9508–16.CrossRefPubMedPubMedCentralGoogle Scholar
  108. 108.
    Nielsen FC, Nielsen J, Christiansen J. A family of IGF-II mRNA binding proteins (IMP) involved in RNA trafficking. Scand J Clin Lab Invest Suppl. 2001;234:93–9.CrossRefPubMedGoogle Scholar
  109. 109.
    Paquin N, Chartrand P. Local regulation of mRNA translation: new insights from the bud. Trends Cell Biol. 2008;18:105–11.CrossRefPubMedGoogle Scholar
  110. 110.
    Asai DJ, Wilkes DE. The dynein heavy chain family. J Eukaryot Microbiol. 2004;51:23–9.CrossRefPubMedGoogle Scholar
  111. 111.
    Wilkes DE, Watson HE, Mitchell DR, Asai DJ. Twenty-five dyneins in Tetrahymena: a re-examination of the multidynein hypothesis. Cell Motil Cytoskeleton. 2008;65:342–51.CrossRefPubMedGoogle Scholar
  112. 112.
    Odronitz F, Becker S, Kollmar M. Reconstructing the phylogeny of 21 completely sequenced arthropod species based on their motor proteins. BMC Genom. 2009;10:173.CrossRefGoogle Scholar
  113. 113.
    Lawrence CJ, Morris NR, Meagher RB, Dawe RK. Dyneins have run their curse in plant lineage. Traffic. 2001;2:362–3.CrossRefPubMedGoogle Scholar
  114. 114.
    Rajagopalan V, Wilkes DE. Evolution of the dynein heavy chain family in ciliates. J Eukaryot Microbiol. 2015;63:138–41.CrossRefPubMedGoogle Scholar

Copyright information

© Springer International Publishing Switzerland 2016

Authors and Affiliations

  1. 1.Institute of Cell BiologyUniversity of BernBernSwitzerland
  2. 2.Division of Basic ResearchNational Institute of Cancer (INCan)Mexico CityMexico

Personalised recommendations