Hox Genes pp 145-181 | Cite as

Transgenesis in Non-model Organisms: The Case of Parhyale

  • Zacharias Kontarakis
  • Anastasios PavlopoulosEmail author
Part of the Methods in Molecular Biology book series (MIMB, volume 1196)


One of the most striking manifestations of Hox gene activity is the morphological and functional diversity of arthropod body plans, segments, and associated appendages. Among arthropod models, the amphipod crustacean Parhyale hawaiensis satisfies a number of appealing biological and technical requirements to study the Hox control of tissue and organ morphogenesis. Parhyale embryos undergo direct development from fertilized eggs into miniature adults within 10 days and are amenable to all sorts of embryological and functional genetic manipulations. Furthermore, each embryo develops a series of specialized appendages along the anterior–posterior body axis, offering exceptional material to probe the genetic basis of appendage patterning, growth, and differentiation. Here, we describe the methodologies and techniques required for transgenesis-based gain-of-function studies of Hox genes in Parhyale embryos. First, we introduce a protocol for efficient microinjection of early-stage Parhyale embryos. Second, we describe the application of fast and reliable assays to test the activity of the Minos DNA transposon in embryos. Third, we present the use of Minos-based transgenesis vectors to generate stable and transient transgenic Parhyale. Finally, we describe the development and application of a conditional heat-inducible misexpression system to study the role of the Hox gene Ultrabithorax in Parhyale appendage specialization. Beyond providing a useful resource for Parhyalists, this chapter also aims to provide a road map for researchers working on other emerging model organisms. Acknowledging the time and effort that need to be invested in developing transgenic approaches in new species, it is all worth it considering the wide scope of experimentation that opens up once transgenesis is established.

Key words

Arthropods Crustaceans Parhyale hawaiensis Minos transgenesis Microinjections Heat-shock promoter Conditional gene misexpression Hox genes Ultrabithorax Appendage development 



We dedicate this chapter to the memory of Thanasis Loukeris, whose work paved the way for transgenic approaches in non-model organisms. We are grateful to Frederike Alwes for providing the drawings and photo shown in Fig. 2. Many protocols described in this chapter have been developed in close interaction with our Ph.D. supervisor and mentor Michalis Averof. Z.K. was supported by an EMBO long-term fellowship, and A.P. by a Marie Curie Intra-European fellowship and by the Howard Hughes Medical Institute.


  1. 1.
    Brusca RC, Brusca GJ (2003) Invertebrates, 2nd edn. Sinauer Associates, Sunderland, MAGoogle Scholar
  2. 2.
    Browne WE, Price AL, Gerberding M et al (2005) Stages of embryonic development in the amphipod crustacean, Parhyale hawaiensis. Genesis 42:124–149PubMedCrossRefGoogle Scholar
  3. 3.
    Rehm EJ, Hannibal RL, Chaw RC et al (2009) The crustacean Parhyale hawaiensis: a new model for arthropod development. Cold Spring Harb Protoc, pdb.emo114Google Scholar
  4. 4.
    Hannibal RL, Price AL, Parchem RJ et al (2012) Analysis of snail genes in the crustacean Parhyale hawaiensis: insight into snail gene family evolution. Dev Genes Evol 222:139–151PubMedCrossRefGoogle Scholar
  5. 5.
    Prpic NM, Telford MJ (2008) Expression of homothorax and extradenticle mRNA in the legs of the crustacean Parhyale hawaiensis: evidence for a reversal of gene expression regulation in the pancrustacean lineage. Dev Genes Evol 218:333–339PubMedCentralPubMedCrossRefGoogle Scholar
  6. 6.
    Schaeper ND, Pechmann M, Damen WG et al (2010) Evolutionary plasticity of collier function in head development of diverse arthropods. Dev Biol 344:363–376PubMedCrossRefGoogle Scholar
  7. 7.
    Simanton W, Clark S, Clemons A et al (2009) Conservation of arthropod midline netrin accumulation revealed with a cross-reactive antibody provides evidence for midline cell homology. Evol Dev 11:260–268PubMedCentralPubMedCrossRefGoogle Scholar
  8. 8.
    Vargas-Vila MA, Hannibal RL, Parchem RJ et al (2010) A prominent requirement for single-minded and the ventral midline in patterning the dorsoventral axis of the crustacean Parhyale hawaiensis. Development 137:3469–3476PubMedCentralPubMedCrossRefGoogle Scholar
  9. 9.
    Browne WE, Schmid BG, Wimmer EA et al (2006) Expression of otd orthologs in the amphipod crustacean, Parhyale hawaiensis. Dev Genes Evol 216:581–595PubMedCrossRefGoogle Scholar
  10. 10.
    Hannibal RL, Price AL, Patel NH (2012) The functional relationship between ectodermal and mesodermal segmentation in the crustacean, Parhyale hawaiensis. Dev Biol 361:427–438PubMedCrossRefGoogle Scholar
  11. 11.
    Kontarakis Z, Pavlopoulos A, Kiupakis A et al (2011) A versatile strategy for gene trapping and trap conversion in emerging model organisms. Development 138:2625–2630PubMedCrossRefGoogle Scholar
  12. 12.
    Liubicich DM, Serano JM, Pavlopoulos A et al (2009) Knockdown of Parhyale Ultrabithorax recapitulates evolutionary changes in crustacean appendage morphology. Proc Natl Acad Sci U S A 106:13892–13896PubMedCentralPubMedCrossRefGoogle Scholar
  13. 13.
    Nestorov P, Battke F, Levesque MP et al (2013) The maternal transcriptome of the crustacean Parhyale hawaiensis is inherited asymmetrically to invariant cell lineages of the ectoderm and mesoderm. PLoS One 8:e56049PubMedCentralPubMedCrossRefGoogle Scholar
  14. 14.
    Pavlopoulos A, Kontarakis Z, Liubicich DM et al (2009) Probing the evolution of appendage specialization by Hox gene misexpression in an emerging model crustacean. Proc Natl Acad Sci U S A 106:13897–13902PubMedCentralPubMedCrossRefGoogle Scholar
  15. 15.
    Wolff C, Scholtz G (2002) Cell lineage, axis formation, and the origin of germ layers in the amphipod crustacean Orchestia cavimana. Dev Biol 250:44–58PubMedCrossRefGoogle Scholar
  16. 16.
    Wolff C, Scholtz G (2008) The clonal composition of biramous and uniramous arthropod limbs. Proc Biol Sci 275:1023–1028PubMedCentralPubMedCrossRefGoogle Scholar
  17. 17.
    Ito A, Aoki MN, Yahata K et al (2011) Embryonic development and expression analysis of Distal-less in Caprella scaura (Crustacea, Amphipoda, Caprellidea). Biol Bull 221:206–214PubMedGoogle Scholar
  18. 18.
    Aspiras AC, Prasad R, Fong DW et al (2012) Parallel reduction in expression of the eye development gene hedgehog in separately derived cave populations of the amphipod Gammarus minus. J Evol Biol 25:995–1001PubMedCrossRefGoogle Scholar
  19. 19.
    Alwes F, Hinchen B, Extavour CG (2011) Patterns of cell lineage, movement, and migration from germ layer specification to gastrulation in the amphipod crustacean Parhyale hawaiensis. Dev Biol 359:110–123PubMedCrossRefGoogle Scholar
  20. 20.
    Gerberding M, Browne WE, Patel NH (2002) Cell lineage analysis of the amphipod crustacean Parhyale hawaiensis reveals an early restriction of cell fates. Development 129:5789–5801PubMedCrossRefGoogle Scholar
  21. 21.
    Extavour CG (2005) The fate of isolated blastomeres with respect to germ cell formation in the amphipod crustacean Parhyale hawaiensis. Dev Biol 277:387–402PubMedCrossRefGoogle Scholar
  22. 22.
    Price AL, Modrell MS, Hannibal RL et al (2010) Mesoderm and ectoderm lineages in the crustacean Parhyale hawaiensis display intra-germ layer compensation. Dev Biol 341:256–266PubMedCrossRefGoogle Scholar
  23. 23.
    Rehm EJ, Hannibal RL, Chaw RC et al (2009) Injection of Parhyale hawaiensis blastomeres with fluorescently labeled tracers. Cold Spring Harb Protoc, pdb.prot5128Google Scholar
  24. 24.
    Rehm EJ, Hannibal RL, Chaw RC et al (2009) In situ hybridization of labeled RNA probes to fixed Parhyale hawaiensis embryos. Cold Spring Harb Protoc, pdb.prot5130Google Scholar
  25. 25.
    Rehm EJ, Hannibal RL, Chaw RC et al (2009) Antibody staining of Parhyale hawaiensis embryos. Cold Spring Harb Protoc, pdb.prot5129Google Scholar
  26. 26.
    Rehm EJ, Hannibal RL, Chaw RC et al (2009) Fixation and dissection of Parhyale hawaiensis embryos. Cold Spring Harb Protoc, pdb.prot5127Google Scholar
  27. 27.
    Ozhan-Kizil G, Havemann J, Gerberding M (2009) Germ cells in the crustacean Parhyale hawaiensis depend on Vasa protein for their maintenance but not for their formation. Dev Biol 327:230–239PubMedCrossRefGoogle Scholar
  28. 28.
    Pavlopoulos A, Averof M (2005) Establishing genetic transformation for comparative developmental studies in the crustacean Parhyale hawaiensis. Proc Natl Acad Sci U S A 102:7888–7893PubMedCentralPubMedCrossRefGoogle Scholar
  29. 29.
    Chaw RC, Patel NH (2012) Independent migration of cell populations in the early gastrulation of the amphipod crustacean Parhyale hawaiensis. Dev Biol 371:94–109PubMedCrossRefGoogle Scholar
  30. 30.
    Blythe MJ, Malla S, Everall R et al (2012) High through-put sequencing of the Parhyale hawaiensis mRNAs and microRNAs to aid comparative developmental studies. PLoS One 7:e33784PubMedCentralPubMedCrossRefGoogle Scholar
  31. 31.
    Parchem RJ, Poulin F, Stuart AB et al (2010) BAC library for the amphipod crustacean, Parhyale hawaiensis. Genomics 95:261–267PubMedCrossRefGoogle Scholar
  32. 32.
    Zeng V, Villanueva KE, Ewen-Campen BS et al (2011) De novo assembly and characterization of a maternal and developmental transcriptome for the emerging model crustacean Parhyale hawaiensis. BMC Genomics 12:581PubMedCentralPubMedCrossRefGoogle Scholar
  33. 33.
    Dohle W, Scholtz G (1988) Clonal analysis of the crustacean segment: the discordance between genealogical and segmental borders. Development 104(Suppl):147–160Google Scholar
  34. 34.
    Scholtz G, Patel NH, Dohle W (1994) Serially homologous engrailed stripes are generated via different cell lineages in the germ band of amphipod crustaceans (Malacostraca, Peracarida). Int J Dev Biol 38:471–478PubMedGoogle Scholar
  35. 35.
    Klinakis AG, Loukeris TG, Pavlopoulos A et al (2000) Mobility assays confirm the broad host-range activity of the Minos transposable element and validate new transformation tools. Insect Mol Biol 9:269–275PubMedCrossRefGoogle Scholar
  36. 36.
    Pavlopoulos A, Oehler S, Kapetanaki MG et al (2007) The DNA transposon Minos as a tool for transgenesis and functional genomic analysis in vertebrates and invertebrates. Genome Biol 8(Suppl 1):S2PubMedCentralPubMedCrossRefGoogle Scholar
  37. 37.
    Berghammer AJ, Klingler M, Wimmer EA (1999) A universal marker for transgenic insects. Nature 402:370–371PubMedCrossRefGoogle Scholar
  38. 38.
    Horn C, Schmid BG, Pogoda FS et al (2002) Fluorescent transformation markers for insect transgenesis. Insect Biochem Mol Biol 32:1221–1235PubMedCrossRefGoogle Scholar
  39. 39.
    Wimmer EA (2003) Innovations: applications of insect transgenesis. Nat Rev Genet 4:225–232PubMedCrossRefGoogle Scholar
  40. 40.
    Pavlopoulos A (2011) Identification of DNA sequences that flank a known region by inverse PCR. Methods Mol Biol 772:267–275PubMedCrossRefGoogle Scholar
  41. 41.
    Sambrook J, Russell DW (2001) Molecular cloning: a laboratory manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NYGoogle Scholar
  42. 42.
    Klinakis AG, Zagoraiou L, Vassilatis DK et al (2000) Genome-wide insertional mutagenesis in human cells by the Drosophila mobile element Minos. EMBO Rep 1:416–421PubMedCentralPubMedCrossRefGoogle Scholar
  43. 43.
    Pavlopoulos A, Berghammer AJ, Averof M et al (2004) Efficient transformation of the beetle Tribolium castaneum using the Minos transposable element: quantitative and qualitative analysis of genomic integration events. Genetics 167:737–746PubMedCentralPubMedCrossRefGoogle Scholar
  44. 44.
    Kapetanaki MG, Loukeris TG, Livadaras I et al (2002) High frequencies of Minos transposon mobilization are obtained in insects by using in vitro synthesized mRNA as a source of transposase. Nucleic Acids Res 30:3333–3340PubMedCentralPubMedCrossRefGoogle Scholar
  45. 45.
    Arca B, Zabalou S, Loukeris TG et al (1997) Mobilization of a Minos transposon in Drosophila melanogaster chromosomes and chromatid repair by heteroduplex formation. Genetics 145:267–279PubMedCentralPubMedGoogle Scholar
  46. 46.
    Horn C, Wimmer EA (2000) A versatile vector set for animal transgenesis. Dev Genes Evol 210:630–637PubMedCrossRefGoogle Scholar
  47. 47.
    Miller DF, Holtzman SL, Kaufman TC (2002) Customized microinjection glass capillary needles for P-element transformations in Drosophila melanogaster. Biotechniques 33:366–367, 369–370, 372 passimGoogle Scholar
  48. 48.
    Horn C, Jaunich B, Wimmer EA (2000) Highly sensitive, fluorescent transformation marker for Drosophila transgenesis. Dev Genes Evol 210:623–629PubMedCrossRefGoogle Scholar
  49. 49.
    Tour E, Hittinger CT, McGinnis W (2005) Evolutionarily conserved domains required for activation and repression functions of the Drosophila Hox protein Ultrabithorax. Development 132:5271–5281PubMedCrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media New York 2014

Authors and Affiliations

  1. 1.Max Planck Institute for Heart and Lung ResearchBad NauheimGermany
  2. 2.Max Planck Institute of Molecular Cell Biology and GeneticsDresdenGermany
  3. 3.Howard Hughes Medical InstituteAshburnUSA

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