Abstract
The diploid pipid frog Xenopus tropicalis has recently emerged as a powerful new model system for combining genetic and genomic analysis of tetrapod development with embryological and biochemical assays. Its early development closely resembles that of its well-understood tetraploid relative Xenopus laevis, from which techniques and reagents can be readily transferred, but its compact genome is highly syntenic with those of amniotes. Genetic approaches are facilitated by the large number of embryos produced and the ease of haploid genetics and gynogenesis.
This is a preview of subscription content, log in via an institution to check access.
Access this chapter
Tax calculation will be finalised at checkout
Purchases are for personal use only
References
Gurdon, J. B. and Hopwood, N. (2000) The introduction of Xenopus laevis into developmental biology: of empire, pregnancy testing and ribosomal genes. Int. J. Dev. Biol. 44, 43–50.
Hellsten, U., Harland, R. M., Gilchrist, M. J., Hendrix, D., Jurka, J., Kapitonov, V., Ovcharenko, I., Putnam, N. H., Shu, S., Taher, L., Blitz, I. L., Blumberg, B., Dichmann, D. S., Dubchak, I., Amaya, E., Detter, J. C., Fletcher, R., Gerhard, D. S., Goodstein, D., Graves, T., Grigoriev, I. V., Grimwood, J., Kawashima, T., Lindquist, E., Lucas, S. M., Mead, P. E., Mitros, T., Ogino, H., Ohta, Y., Poliakov, A. V., Pollet, N., Robert, J., Salamov, A., Sater, A. K., Schmutz, J., Terry, A., Vize, P. D., Warren, W. C., Wells, D., Wills, A., Wilson, R. K., Zimmerman, L. B., Zorn, A. M., Grainger, R., Grammer, T., Khokha, M. K., Richardson, P. M., and Rokhsar, D. S. (2010) The genome of the Western clawed frog Xenopus tropicalis. Science 328, 633–6.
Goda, T., Abu-Daya, A., Carruthers, S., Clark, M. D., Stemple, D. L., and Zimmerman, L. B. (2006) Genetic screens for mutations affecting development of Xenopus tropicalis. PLoS Genet. 2, e91.
Grammer, T. C., Khokha, M. K., Lane, M. A., Lam, K., and Harland, R. M. (2005) Identification of mutants in inbred Xenopus tropicalis. Mech. Dev. 122, 263–72.
Noramly, S., Zimmerman, L., Cox, A., Aloise, R., Fisher, M., and Grainger, R. M. (2005) A gynogenetic screen to isolate naturally occurring recessive mutations in Xenopus tropicalis. Mech. Dev. 122, 273–87.
Abu-Daya, A., Sater, A. K., Wells, D. E., Mohun, T. J., and Zimmerman, L. B. (2009) Absence of heartbeat in the Xenopus tropicalis mutation muzak is caused by a nonsense mutation in cardiac myosin myh6. Dev. Biol. 336, 20–9.
Khokha, M. K., Krylov, V., Reilly, M. J., Gall, J. G., Bhattacharya, D., Cheung, C. Y. J., Kaufman, S., Lam, D. K., Macha, J., Ngo, C., Prakash, N., Schmidt, P., Tlapakova, T., Trivedi, T., Tumova, L., Abu-Daya, A., Geach, T., Vendrell, E., Ironfield, H., Sinzelle, L., Sater, A. K., Wells, D. E., Harland, R. M., and Zimmerman, L. B. (2009) Rapid gynogenetic mapping of Xenopus tropicalis mutations to chromosomes. Dev. Dyn. 238, 1398–46.
Reinschmidt, D., Friedman, J., Hauth, J., Ratner, E., Cohen, M., Miller, M., Krotoski, D., and Tompkins, R. (1985) Gene–centromere mapping in Xenopus laevis. J. Hered. 76, 345–7.
Tompkins, R. and Reinschmidt, D. (1991) Experimentally induced homozygosity in Xenopus laevis. Methods Cell Biol. 36, 35–44.
Kroll, K. L. and Amaya, E. (1996) Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signaling requirements during gastrulation. Development 122, 3173–83.
Yergeau, D. A. and Mead, P. E. (2007) Manipulating the Xenopus genome with transposable elements. Genome Biol. 8 Suppl 1, S11.
Ogino, H., McConnell, W. B., and Grainger, R. M. (2006) Highly efficient transgenesis in Xenopus tropicalis using I-SceI meganuclease. Mech. Dev. 123, 103–13.
Allen, B. G. and Weeks, D. L. (2005) Transgenic Xenopus laevis embryos can be generated using phiC31 integrase. Nat. Methods 2, 975–9.
Hirsch, N., Zimmerman, L. B., Gray, J., Chae, J., Curran, K. L., Fisher, M., Ogino, H., and Grainger, R. M. (2002) Xenopus tropicalis transgenic lines and their use in the study of embryonic induction. Dev. Dyn. 225, 522–35.
Chae, J., Zimmerman, L. B., and Grainger, R. M. (2002) Inducible control of tissue-specific transgene expression in Xenopus tropicalis transgenic lines. Mech. Dev. 117, 235–41.
Hartley, K. O., Nutt, S. L., and Amaya, E. (2002) Targeted gene expression in transgenic Xenopus using the binary Gal4-UAS system. Proc. Natl. Acad. Sci. USA 99, 1377–82.
Ryffel, G. U., Werdien, D., Turan, G., Gerhards, A., Goosses, S., and Senkel, S. (2003) Tagging muscle cell lineages in development and tail regeneration using Cre recombinase in transgenic Xenopus. Nucleic Acids Res. 31, e44.
Lehman, C. W. and Carroll, D. (1991) Homologous recombination catalyzed by a nuclear extract from Xenopus oocytes. Proc. Natl. Acad. Sci. USA 88, 10840–4.
Doyon, Y., McCammon, J. M., Miller, J. C., Faraji, F., Ngo, C., Katibah, G. E., Amora, R., Hocking, T. D., Zhang, L., Rebar, E. J., Gregory, P. D., Urnov, F. D., and Amacher, S. L. (2008) Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Nat. Biotechnol. 26, 702–8.
Carruthers, S. and Stemple, D. L. (2006) Genetic and genomic prospects for Xenopus tropicalis research. Sem. Cell Dev. Biol. 17, 146–53.
Sargent, M. G. and Mohun, T. J. (2005) Cryopreservation of sperm of Xenopus laevis and Xenopus tropicalis. Genesis 41, 41–6.
Bahary, N., Davidson, A., Ransom, D., Shepard, J., Stern, H., Trede, N., Zhou, Y., Barut, B., and Zon, L. I. (2004) The Zon laboratory guide to positional cloning in zebrafish. Methods Cell Biol. 77, 305–29.
Bassam, B. J., Caetano-Anolles, G., and Gresshoff, P. M. (1991) Fast and sensitive silver staining of DNA in polyacrylamide gels. Anal Biochem 196, 80–3.
Streisinger, G., Singer, F., Walker, C., Knauber, D., and Dower, N. (1986) Segregation analyses and gene–centromere distances in zebrafish. Genetics 112, 311–19.
Vos, P., Hogers, R., Bleeker, M., Reijans, M., van de Lee, T., Hornes, M., Frijters, A., Pot, J., Peleman, J., Kuiper, M., et al. (1995) AFLP: a new technique for DNA fingerprinting. Nucleic Acids Res. 23, 4407–14.
Neff, M. M., Neff, J. D., Chory, J., and Pepper, A. E. (1998) dCAPS, a simple technique for the genetic analysis of single nucleotide polymorphisms: experimental applications in Arabidopsis thaliana genetics. Plant J. Cell Mol. Biol. 14, 387–92.
Konieczny, A. and Ausubel, F. M. (1993) A procedure for mapping Arabidopsis mutations using co-dominant ecotype-specific PCR-based markers. Plant J. Cell Mol. Biol. 4, 403–10.
Chang, Y. -F., Imam, J. S., and Wilkinson, M. F. (2007) The nonsense-mediated decay RNA surveillance pathway. Annu. Rev. Biochem. 76, 51–74.
Acknowledgements
A large number of colleagues have adapted and developed the protocols outlined in this chapter. In addition to all past and present members of the Zimmerman laboratory, the authors would particularly like to thank Rob Grainger (University of Virginia), Richard Harland (UC Berkeley) and Mustafa Khokha (Yale). T.J.G. and L.B.Z. are funded by the Medical Research Council, UK.
Author information
Authors and Affiliations
Corresponding author
Editor information
Editors and Affiliations
Rights and permissions
Copyright information
© 2011 Springer Science+Business Media, LLC
About this protocol
Cite this protocol
Geach, T.J., Zimmerman, L.B. (2011). Developmental Genetics in Xenopus tropicalis . In: Pelegri, F. (eds) Vertebrate Embryogenesis. Methods in Molecular Biology, vol 770. Humana Press, Totowa, NJ. https://doi.org/10.1007/978-1-61779-210-6_4
Download citation
DOI: https://doi.org/10.1007/978-1-61779-210-6_4
Published:
Publisher Name: Humana Press, Totowa, NJ
Print ISBN: 978-1-61779-209-0
Online ISBN: 978-1-61779-210-6
eBook Packages: Springer Protocols